Bioreactors in Stem Cell Biology: Methods and Protocols [2 ed.] 1071620177, 9781071620175

Table of contents :
Preface
Contents
Contributors
Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines
1 Introduction
2 Materials
2.1 Equipment
2.2 Bioreactor
2.3 Cell Culture Media, Feeds, and Supplements
2.4 Antifoam
2.5 Controller
2.6 Probes
2.7 Other Supplements
3 Methods
3.1 Cell Expansion
3.2 Probe Calibration and Sterilization
3.3 Bioreactor Preparation
3.4 Bioreactor Setpoints
3.5 Media Preparation/Hold
3.6 Inoculation
3.7 Fed-Batch (Sampling and Feeding)
3.8 Harvest and Clarification
4 Notes
References
Volumetric Mass Transfer Coefficient Measurement in a Stirred Tank Reactor
1 Introduction
1.1 The Measurement of kLa
1.2 The Unsteady-State (Dynamic) Method
1.3 The Steady-State Method
2 Materials
2.1 Bioreactor
2.2 Microorganism
2.3 Growth Media
2.4 Spectrophotometer
2.5 Chronometer
3 Methods
3.1 Installation of the Bioreactor
3.2 Preparation of the Growth Media
3.3 Preparation of Microorganism
3.4 Operation of the STR
3.5 Measurement of the Oxygen Transfer Coefficient: kLa
4 Notes
References
Fabrication Protocol for Thermoplastic Microfluidic Devices: Nanoliter Volume Bioreactors for Cell Culturing
1 Introduction
2 Materials
3 Methods
3.1 Thermoplastic Chip Design and Preparing Its Mold
3.1.1 Photolithography
3.1.2 Electrochemical Wet Etching
3.2 Fabrication of Electrodes
3.2.1 Creating an Electrode Design
3.2.2 Chromium and Gold Deposition on Blank COP Substrate
3.2.3 Cr/Au Etching to Form Electrodes on Coated COP Sample
3.3 Drilling and Planarization
3.4 Hot Embossing
3.5 Thermo-Compression Bonding
3.6 Establishing the Experiment
3.6.1 Microfluidic Device Without Electrodes
3.6.2 Microfluidic Device with Electrodes
4 Notes
References
Expansion of Human Pluripotent Stem Cells in Stirred Tank Bioreactors
1 Introduction
2 Materials
2.1 L7 TFO2 hPSC Complete Medium
2.2 Passaging Solutions
2.3 Buffers and Reagents
2.4 Tubing
2.5 Bioreactor
3 Methods
3.1 Extracellular Matrix Coating of T-75 and 1-Layer Cell Stack
3.2 hPSC Culture on T-75 Flask
3.3 hPSC Culture on 2D Seed Train
3.4 Microcarrier Coating
3.5 Bioreactor Setup
3.6 Cell Inoculum Introduction
3.7 Cell Culture Medium Perfusion Setup
3.8 Cell Count during Expansion
3.9 Cell Harvest
4 Notes
References
High-Efficiency Differentiation of Human Pluripotent Stem Cells to Hematopoietic Stem/Progenitor Cells in Random Positioning M
1 Introduction
2 Materials
2.1 Cell Lines
2.2 Cell Culture Medium and Reagents
2.3 Bioreactor Equipment
2.4 Immunostaining and Flow Cytometry
2.5 Other Material
3 Methods
3.1 Feeder-Free Expansion of Human PSCs
3.2 Mesoderm Induction from hPSCs
3.3 Generation of Hemogenic Endothelium Progenitor
3.4 Generation of Hematopoietic Progenitor Cells
3.5 Characterization of Differentiated Cells Derived from hESCs in Bioreactor
3.5.1 Immunofluorescence Staining
3.5.2 Flow Cytometry
4 Notes
References
Integrating Human-Induced Pluripotent Stem Cell Expansion Capability and Cardiomyocyte Differentiation Potential in a Microcar
1 Introduction
2 Materials
2.1 General Equipment and Consumables
2.2 Media and Consumables for hiPSC Cultures
2.3 Reagents and Consumables for Microcarrier Cultures
2.4 Media and Small Molecules for CM Differentiation
2.5 Antibodies and Reagents
2.6 Preparation of Extracellular Matrices for hPSCs Cultures
2.7 Preparation of Cell Culture Medium
2.8 Preparation of Small Molecules
2.9 Preparation of Microcarriers
3 Methods
3.1 hiPSC Culture
3.2 Screening for High Cardiac Differentiation Potency hiPSC Lines (Fig. 1)
3.2.1 hiPSC Seeding for CM Differentiation
3.2.2 CM Differentiation (Monolayer Method)
3.2.3 Characterization of hiPSC-Derived CM from Monolayer by Flow Cytometry (Fig. 2)
3.3 Testing of Cell Expansion in Microcarrier Cultures
3.3.1 Preparation of GelTrex-Coated Microcarriers
3.3.2 Seeding Cells into Spinner Flask
3.3.3 Cell Counting and Imaging during Cultivation
3.3.4 Evaluation the Pluripotency of hiPSCs in Spinner Flask Cultures by Flow Cytometry (Fig. 3)
3.4 Cardiac Differentiation in Microcarrier Spinner Cultures (Fig. 4)
4 Notes
References
Chemically Defined, Xeno-Free Expansion of Human Mesenchymal Stem Cells (hMSCs) on Benchtop-Scale Using a Stirred Single-Use B
1 Introduction
1.1 Expansion of hMSCs in Stirred Single-Use Bioreactors
1.2 Chemically Defined, Serum-Free Expansion of hMSCs
1.3 General Procedure for and Results of the Serum- and Xeno-Free Expansion of Human Mesenchymal Stem Cells from Adipose Tissu...
2 Materials
2.1 hASC Inoculum Production in T75-Flasks
2.2 hASC Expansion in Spinner Flasks
2.3 hASC Expansion in the BioBLU
2.4 Process Analytics
3 Methods
3.1 T75-Flask-Based Inoculum Production
3.2 hASC Expansion in Corning´s Spinner Flasks
3.3 hASC Expansion in Eppendorf´s Instrumented BioBLU 0.3c
3.4 Sampling and Quality Control
4 Notes
References
Large-Scale Expansion of Umbilical Cord Mesenchymal Stem Cells with Microcarrier Tablets in Bioreactor
1 Introduction
2 Materials
2.1 Cell Culture Reagents
2.2 3D TableTrix Microcarrier Tablets
2.3 3D FloTrix Digest Solution
2.4 Cell Quality Assessment-Related Reagents
2.5 Other Reagents
3 Methods
3.1 Setting up 3D FloTrix vivaSPIN Bioreactor on Day 1
3.2 Inoculation on Day 0
3.3 Cell Culture, Medium Replenishment, and Growth Monitoring on Days 1-3
3.4 Cell Harvesting on Day 4
4 Notes
References
Optimized Method to Improve Cell Activity in 3D Scaffolds Under a Dual Real-Time Dynamic Bioreactor System
1 Introduction
2 Materials
2.1 Bioreactor System
3 Methods
3.1 Sample Preparation
3.2 Sample Transfer and Chamber Assembly
3.3 Mechanical Compression Load and Fluid Flow Parameters
4 Notes
References
In Vitro 3D Mechanical Stimulation to Tendon-Derived Stem Cells by Bioreactor
1 Introduction
2 Materials
2.1 Isolation of Mice TDSCs
2.2 Mechanical Stimulation
2.2.1 Scaffold-Free Mechanical Stimulation
2.2.2 Scaffold-Based Mechanical Stimulation
2.3 Extraction of RNA for Validation of the System
3 Methods
3.1 Isolation of Mice TDSCs
3.2 Mechanical Stimulation
3.2.1 Scaffold-Free Mechanical Stimulation
3.2.2 Scaffold-Based Mechanical Stimulation
3.3 Extraction of RNA for Validation of the System
4 Notes
References
Microcarrier-Supported Culture of Chondrocytes in Continuously Rocked Disposable Bioreactor
1 Introduction
2 Materials
2.1 Microcarrier Conditioning
2.2 Preparation of CP5 Cell Inoculum
2.3 Cell Culture
2.4 Analytical Methods
2.4.1 Preparation of Samples for Analysis
2.4.2 Cell Staining
2.4.3 Activity of Intracellular Oxidoreductases
2.4.4 Glucose Consumption Rate
2.4.5 Activity of Lactate Dehydrogenase (LDH)
2.5 General Description of the Setup
2.5.1 ReadyToProcess WAVE 25 Bioreactor System
3 Methods
3.1 Rehydration, Conditioning, and Sterilization of Microcarriers
3.2 Preparation of CP5 Chondrocyte Inoculum
3.3 Maintaining CP5 Cells in Bioreactor System
3.4 Analytical Methods
3.4.1 Preparation of Samples for Analysis
3.4.2 Cell Density and Viability
3.4.3 Metabolic Activity of Cells Adhered to Microcarriers
3.4.4 Specific Glucose Consumption Rate
3.4.5 Activity of Lactate Dehydrogenase
3.4.6 DO and pH Level Measurement
4 Notes
References
Tracheal In Vitro Reconstruction Using a Decellularized Bio-Scaffold in Combination with a Rotating Bioreactor
1 Introduction
2 Materials
2.1 Porcine Trachea Collection
2.2 Generation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold
2.3 Human Chondrocyte Propagation and Maintenance
2.4 Repopulation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold with Human Chondrocytes and Bioreactor Setup
3 Methods
3.1 Porcine Trachea Collection
3.2 Generation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold
3.3 Human Chondrocyte Propagation and Maintenance
3.4 Repopulation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold with Human Chondrocytes and Bioreactor Setup
4 Notes
References
Bioreactor-Based De-epithelialization of Long-Segment Tracheal Grafts
1 Introduction
2 Materials
2.1 Bioreactor Main Components (Fig. 1)
2.2 Bioreactor Extra Components
2.3 Other Materials
2.4 Tools
2.5 Devices
2.6 Antibiotics, Media, and Detergents
3 Methods
3.1 Preparation of Antibiotics, Media, and Detergents (See Notes 1 and 2)
3.1.1 Cocktail of Antibiotics
3.1.2 DMEM + Antibiotics
3.1.3 1% Sodium Dodecyl Sulfate
3.1.4 1% Triton X-100
3.2 Graft Procurement and Preparation
3.3 Bioreactor Setup for De-epithelialization
3.3.1 Detergent Circuitries
3.3.2 Bioreactor Setup
3.3.3 Positioning the Trachea
3.3.4 Motor and Pumps Installation
3.4 Tracheal Graft De-epithelialization
3.5 Decontamination of the De-epithelialized Graft
4 Notes
References
Production of Extracellular Vesicles Using a CELLine Adherent Bioreactor Flask
1 Introduction
2 Materials
2.1 Cell Culture
2.2 EV Isolation Materials
2.3 Characterization Materials
3 Methods
3.1 Preparation of CELLine AD 1000 Bioreactor Flask (Figs. 1 and 2)
3.2 Inoculation of Cells (See Note 4)
3.3 Adapting Cells to Long-Term Bioreactor Media (See Note 6)
3.4 Continually Harvesting EVs and Monitoring Shed Cells from the Bioreactor (Fig. 2)
3.5 EV Isolation and Purification (See Note 10)
3.6 Characterizing EVs
3.7 Imaging the Bioreactor Growth Surface
3.8 EV-Associated RNA
4 Notes
References
Extracellular Vesicle Collection from Human Stem Cells Grown in Suspension Bioreactors
1 Introduction
2 Materials
2.1 Materials for hiPSC Differentiation in Planar and Bioreactor Cultures
2.2 Materials for hMSC Expansion in Planar and Bioreactor Cultures
2.3 Bioreactor Preparation
2.4 Materials for EV Isolation from Human Stem Cells Grown in Bioreactor Cultures
3 Methods
3.1 Culture and Expansion of hiPSCs in Planar Culture
3.2 Differentiation of NPC Organoid from hiPSCs in Spinner Flasks
3.3 Expansion of hMSCs in Planar Culture
3.4 Expansion of hMSCs in PBS-VW Bioreactors
3.5 EV Isolation from the Collected Media
4 Notes
References
Bacterial Nanocellulose-Based Grafts for Cell Colonization Studies: An In Vitro Bioreactor Perfusion Model
1 Introduction
2 Materials
2.1 Bioreactor Setup
2.2 Assays and Staining Solutions
2.3 Cell Culture
2.4 Antibodies
2.5 Chemicals and Solutions
2.6 Consumables
2.7 Hardware
3 Methods
3.1 General Considerations
3.2 Bioreactor Setup
3.3 Connection of Small Diameter Vascular Grafts to Bioreactor Chamber
3.4 Cell Seeding
3.5 Lactate Monitoring
3.6 WST-1 Proliferation Assay
3.6.1 Perform WST-1 Assay (Modified from Manufacturer´s Instructions)
3.7 Termination of Experiment
3.8 AcLDL Uptake Assay (Modified from Manufacturer´s Instructions)
3.9 Phalloidin-F-Actin Staining
3.10 Acridine Orange Staining
3.11 CD31 Immunofluorescence Staining
4 Notes
References
A Guideline to Set Up Cascaded Continuous Cultivation with E. coli Bl21 (DE3)
1 Introduction
2 Materials
2.1 Host Cells
2.2 Required Media
2.3 Required Devices for Cultivation
2.4 Required Equipment for Process Analysis
2.4.1 Biomass Determination
2.4.2 Viable Biomass Determination
2.4.3 Metabolite Determination
2.4.4 Product Determination
3 Methods
3.1 Cultivation Setup
3.2 Cultivation Scheme
3.2.1 Preculture and Batch Phase
3.2.2 Continuous Adaptation Phase
3.2.3 Induction Phase
3.3 Sampling and Analysis
3.3.1 Biomass Determination
3.3.2 Determination of Metabolite Accumulation
3.3.3 Product Determination
3.4 Calculation of Flow and Substrate Uptake Rates
3.4.1 Calculation of Flow Rates
3.4.2 Calculation of Substrate Uptake Rates
4 Notes
5 General Notes on Process Intensification for Cascaded Continuous Cultivation
5.1 Dilution Rate
5.2 Substrate Uptake Rate
5.3 Cultivation Temperature
References
Applying Stirred Perfusion to 3D Tissue Equivalents to Mimic the Dynamic In Vivo Microenvironment
1 Introduction
2 Materials
2.1 Cell Culture
2.1.1 HepG2 Tissue Model
2.1.2 Intestinal Tissue Model
Stromal Compartment
Epithelial Compartment
2.2 Processing and Analysis of Samples
3 Methods
3.1 Growth of HepG2 Liver Models for Drug Toxicity Testing
3.1.1 Revival of HepG2 Cells
3.1.2 Preparation of Alvetex Strata
3.1.3 Seeding HepG2 Cells onto the Alvetex Strata
3.1.4 Preparation of the Perfusion Bioreactors
3.1.5 Moving HepG2 Models into the Perfusion System
3.2 Creation of Perfused Intestinal Models
3.2.1 Revival of Cells
3.2.2 Preparation of Alvetex Scaffold
3.2.3 Seeding HDFn Cells onto the Alvetex Scaffold
3.2.4 Seeding Caco2 Cells onto the Stromal Compartments
3.2.5 Preparation of the Perfusion Bioreactors
3.2.6 Moving Intestinal Models into the Perfusion System
3.3 Processing and Analysis of the Perfused Models
3.3.1 Processing Samples for Paraffin Wax Embedding
3.3.2 Generation of Transverse Sections of Tissue Models
3.3.3 Histological Analysis
3.3.4 Immunofluorescent Analysis
3.3.5 MTT Assay
4 Notes
References
Bioreactor-Based Adherent Cells Harvesting from Microcarriers with 3D Printed Inertial Microfluidics
1 Introduction
2 Materials
2.1 3D Printing
2.2 PDMS Device Making (in Replacement of Direct-Printed Device)
2.3 Cell Harvesting
3 Methods
3.1 Device Design
3.2 Direct Fabrication
3.3 Mold Fabrication
3.4 Operation
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2436

Kursad Turksen Editor

Bioreactors in Stem Cell Biology Methods and Protocols Second Edition

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-bystep 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 Pub Med.

Bioreactors in Stem Cell Biology Methods and Protocols Second Edition

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2017-5 ISBN 978-1-0716-2018-2 (eBook) https://doi.org/10.1007/978-1-0716-2018-2 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022 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: Artwork created by Kursad Turksen. 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.

Preface The use of bioreactors in cell biology is becoming more commonplace as attempts are made to scale up production of various types of cells for regenerative medicine and pharmaceutical purposes. Given the rapidity with which bioreactor technologies are advancing and being applied, I have attempted to bring together some of the latest developments and protocols in this second edition on this topic. The protocols in this volume are all new protocols and reflect novel technology development and potential uses that I hope will benefit both established investigators and newcomers to the field. Once again, the protocols gathered here are faithful to the mission statement of the Methods in Molecular Biology series: They are well-established and described in an easy-tofollow, step-by-step fashion so as to be valuable for not only experts but also novices in the stem cell field. That goal is achieved because of the generosity of the contributors who have carefully described their protocols in this volume, and we are very grateful for their efforts. Our thanks as well go to Dr. John Walker, the Editor-in-Chief of the Methods in Molecular Biology series, for giving us the opportunity to create and now update this volume and for supporting us along the way. We are also grateful to Patrick Marton, the Executive Editor of Methods in Molecular Biology and the Springer Protocols collection, for his continuous support from idea to completion of this volume. A special thank you goes to Anna Rakovsky, Assistant Editor for Methods in Molecular Biology, for continuous support from the beginning to the end of this project. We would also like to thank David C. Casey, the Editor of Methods in Molecular Biology, for his outstanding editorial work during the production of this volume. Finally, we would like to thank Daniel Ignatius Jagadisan, Sarumathi Hemachandrane, Anand Ventakachalam, and the rest of the production crew for their work in putting together an outstanding volume. Ottawa, ON, Canada

Kursad Turksen

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ange´lique Schmid, Emanuel Kreidl, Martin Bertschinger, and Patrick Vetsch Volumetric Mass Transfer Coefficient Measurement in a Stirred Tank Reactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aysegul Inam, Ezgi Rojda Taymaz, Mehmet Emin Uslu, Baris Binay, and Irem Deniz Fabrication Protocol for Thermoplastic Microfluidic Devices: Nanoliter Volume Bioreactors for Cell Culturing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elif Gencturk, Senol Mutlu, and Kutlu O. Ulgen Expansion of Human Pluripotent Stem Cells in Stirred Tank Bioreactors. . . . . . . . . . . Marites T. Woon, Puspa R. Pandey, and Inbar Friedrich Ben-Nun High-Efficiency Differentiation of Human Pluripotent Stem Cells to Hematopoietic Stem/Progenitor Cells in Random Positioning Machine Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaohua Lei, Chiyuan Ma, Yujing Cao, Yue Xiong, Jian V. Zhang, and Enkui Duan Integrating Human-Induced Pluripotent Stem Cell Expansion Capability and Cardiomyocyte Differentiation Potential in a Microcarrier Suspension Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Valerie Ho, Gerine Tong, Alan Lam, Shaul Reuveny, and Steve Oh Chemically Defined, Xeno-Free Expansion of Human Mesenchymal Stem Cells (hMSCs) on Benchtop-Scale Using a Stirred Single-Use Bioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Misha Teale, Valentin Jossen, Dieter Eibl, and Regine Eibl Large-Scale Expansion of Umbilical Cord Mesenchymal Stem Cells with Microcarrier Tablets in Bioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huanye Xu, Zhongxiao Cong, Yuanyuan Zhang, Wei Liu, Xiaojun Yan, and Yanan Du Optimized Method to Improve Cell Activity in 3D Scaffolds Under a Dual Real-Time Dynamic Bioreactor System. . . . . . . . . . . . . . . . . . . . . . . . . . . . Flavia Pedrini, Moema A. Hausen, and Eliana A. R. Duek In Vitro 3D Mechanical Stimulation to Tendon-Derived Stem Cells by Bioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ziming Chen, Peilin Chen, Rui Ruan, and Minghao Zheng Microcarrier-Supported Culture of Chondrocytes in Continuously Rocked Disposable Bioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kamil Wierzchowski and Maciej Pilarek

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Tracheal In Vitro Reconstruction Using a Decellularized Bio-Scaffold in Combination with a Rotating Bioreactor. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Georgia Pennarossa, Matteo Ghiringhelli, Fulvio Gandolfi, and Tiziana A. L. Brevini Bioreactor-Based De-epithelialization of Long-Segment Tracheal Grafts . . . . . . . . . . . Alba E. Marin-Araujo, Siba Haykal, and Golnaz Karoubi Production of Extracellular Vesicles Using a CELLine Adherent Bioreactor Flask . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anastasiia Artuyants, Vanessa Chang, Gabrielle Reshef, Cherie Blenkiron, Lawrence W. Chamley, Euphemia Leung, and Colin L. Hisey Extracellular Vesicle Collection from Human Stem Cells Grown in Suspension Bioreactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xuegang Yuan, Xingchi Chen, Changchun Zeng, David G. Meckes Jr, and Yan Li Bacterial Nanocellulose-Based Grafts for Cell Colonization Studies: An In Vitro Bioreactor Perfusion Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Max Wacker, Jan Riedel, Priya Veluswamy, Maximilian Scherner, ¨ lsmann Jens Wippermann, Heike Walles, and Jo¨rn Hu A Guideline to Set Up Cascaded Continuous Cultivation with E. coli Bl21 (DE3) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julian Kopp and Oliver Spadiut Applying Stirred Perfusion to 3D Tissue Equivalents to Mimic the Dynamic In Vivo Microenvironment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Henry W. Hoyle, Claire L. Mobbs, and Stefan A. Przyborski Bioreactor-Based Adherent Cells Harvesting from Microcarriers with 3D Printed Inertial Microfluidics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lin Ding, Reza Moloudi, and Majid Ebrahimi Warkiani Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors ANASTASIIA ARTUYANTS • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Obstetrics and Gynaecology, University of Auckland, Auckland, New Zealand MARTIN BERTSCHINGER • Ichnos Sciences SA, La Chaux-de-Fonds, Switzerland BARIS BINAY • Department of Bioengineering, Gebze Technical University, Kocaeli, Turkey CHERIE BLENKIRON • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Molecular Medicine and Pathology, University of Auckland, Auckland, New Zealand; Auckland Cancer Society Research Centre, University of Auckland, Auckland, New Zealand TIZIANA A. L. BREVINI • Laboratory of Biomedical Embryology, Department of Health, ` degli Studi Animal Science and Food Safety and Center for Stem Cell Research, Universita di Milano, Milan, Italy YUJING CAO • State Key Laboratory of Stem Cell and Reproductive Biology, Institute of Zoology, Chinese Academy of Sciences, Beijing, China LAWRENCE W. CHAMLEY • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Obstetrics and Gynaecology, University of Auckland, Auckland, New Zealand VANESSA CHANG • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Obstetrics and Gynaecology, University of Auckland, Auckland, New Zealand PEILIN CHEN • Centre for Orthopaedic Translational Research, Medical School, University of Western Australia, Nedlands, WA, Australia XINGCHI CHEN • Department of Chemical & Biomedical Engineering, FAMU-FSU College of Engineering, Florida State University, Tallahassee, FL, USA ZIMING CHEN • Centre for Orthopaedic Translational Research, Medical School, University of Western Australia, Nedlands, WA, Australia ZHONGXIAO CONG • Beijing CytoNiche Biotechnology Co., Ltd., Beijing, China IREM DENIZ • Department of Bioengineering, Faculty of Engineering, Manisa Celal Bayar University, Manisa, Turkey LIN DING • School of Biomedical Engineering, University of Technology Sydney, Sydney, NSW, Australia YANAN DU • Department of Biomedical Engineering, School of Medicine, Tsinghua-PKU Center for Life Sciences, Tsinghua University, Beijing, China ENKUI DUAN • State Key Laboratory of Stem Cell and Reproductive Biology, Institute of Zoology, Chinese Academy of Sciences, Beijing, China ELIANA A. R. DUEK • Postgraduate Program in Biotechnology and Environmental Monitoring, Federal University of Sa˜o Carlos (UFSCar), Sorocaba, Brazil; Surgery Department, Faculty of Medical Sciences and Health, Pontifical Catholic University of Sa˜o Paulo (PUC/SP), Sorocaba, Brazil DIETER EIBL • Centre for Biochemical Engineering and Cell Cultivation Techniques, Institute of Chemistry and Biotechnology, Zurich University of Applied Sciences, W€ adenswil, Switzerland

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Contributors

REGINE EIBL • Centre for Biochemical Engineering and Cell Cultivation Techniques, Institute of Chemistry and Biotechnology, Zurich University of Applied Sciences, W€ adenswil, Switzerland INBAR FRIEDRICH BEN-NUN • Cell and Gene Therapy Research and Development, Lonza Inc., Rockville, MD, USA FULVIO GANDOLFI • Laboratory of Biomedical Embryology, Department of Agricultural and ` degli Studi di Environmental Sciences—Production, Landscape, Agroenergy, Universita Milano, Milan, Italy ELIF GENCTURK • Biosystems Engineering Laboratory, Department of Chemical Engineering, Bogazici University, Istanbul, Turkey MATTEO GHIRINGHELLI • Sohnis Research laboratory for Cardiac Electrophysiology and Regenerative Medicine, The Rappaport Faculty of Medicine and Research Institute, Technion-Israel Institute of Technology, Haifa, Israel MOEMA A. HAUSEN • Surgery Department, Faculty of Medical Sciences and Health, Pontifical Catholic University of Sa˜o Paulo (PUC/SP), Sorocaba, Brazil SIBA HAYKAL • Latner Research Laboratories, Division of Thoracic Surgery, University Health Network, Toronto, ON, Canada; Division of Plastic & Reconstructive Surgery, University Health Network, University of Toronto, Toronto, ON, Canada; Institute of Medical Sciences, University of Toronto, Toronto, ON, Canada COLIN L. HISEY • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Obstetrics and Gynaecology, University of Auckland, Auckland, New Zealand VALERIE HO • Bioprocessing Technology Institute, A*STAR Research Entities, Singapore, Singapore HENRY W. HOYLE • Department of Biosciences, Durham University, Durham, UK JO¨RN HU¨LSMANN • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany AYSEGUL INAM • Department of Bioengineering, Faculty of Engineering, Manisa Celal Bayar University, Manisa, Turkey VALENTIN JOSSEN • Centre for Biochemical Engineering and Cell Cultivation Techniques, Institute of Chemistry and Biotechnology, Zurich University of Applied Sciences, W€ adenswil, Switzerland GOLNAZ KAROUBI • Latner Research Laboratories, Division of Thoracic Surgery, University Health Network, Toronto, ON, Canada; Department of Laboratory Medicine and Pathobiology, University of Toronto, Toronto, ON, Canada; Department of Mechanical and Industrial Engineering, University of Toronto, Toronto, ON, Canada JULIAN KOPP • Research Division Integrated Bioprocess Development, Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria EMANUEL KREIDL • Ichnos Sciences SA, La Chaux-de-Fonds, Switzerland ALAN LAM • Bioprocessing Technology Institute, A*STAR Research Entities, Singapore, Singapore XIAOHUA LEI • Center for Energy Metabolism and Reproduction, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China EUPHEMIA LEUNG • Auckland Cancer Society Research Centre, University of Auckland, Auckland, New Zealand YAN LI • Department of Chemical & Biomedical Engineering, FAMU-FSU College of Engineering, Florida State University, Tallahassee, FL, USA WEI LIU • Beijing CytoNiche Biotechnology Co., Ltd., Beijing, China

Contributors

xi

CHIYUAN MA • Center for Energy Metabolism and Reproduction, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China ALBA E. MARIN-ARAUJO • Latner Research Laboratories, Division of Thoracic Surgery, University Health Network, Toronto, ON, Canada DAVID G. MECKES JR • Department of Biomedical Sciences, Florida State University College of Medicine, Tallahassee, FL, USA CLAIRE L. MOBBS • Department of Biosciences, Durham University, Durham, UK REZA MOLOUDI • Skeletal Biology and Engineering Research Centre, KU Leuven, Leuven, Belgium SENOL MUTLU • BUMEMS Laboratory, Department of Electrical and Electronics Engineering, Bogazici University, Istanbul, Turkey STEVE OH • Bioprocessing Technology Institute, A*STAR Research Entities, Singapore, Singapore PUSPA R. PANDEY • Cell and Gene Therapy Research and Development, Lonza Inc., Rockville, MD, USA FLAVIA PEDRINI • Postgraduate Program in Biotechnology and Environmental Monitoring, Federal University of Sa˜o Carlos (UFSCar), Sorocaba, Brazil GEORGIA PENNAROSSA • Laboratory of Biomedical Embryology, Department of Health, ` degli Studi Animal Science and Food Safety and Center for Stem Cell Research, Universita di Milano, Milan, Italy MACIEJ PILAREK • Warsaw University of Technology, Faculty of Chemical and Process Engineering, Warsaw, Poland STEFAN A. PRZYBORSKI • Department of Biosciences, Durham University, Durham, UK; REPROCELL Europe Ltd., NETPark Incubator, Sedgefield, UK GABRIELLE RESHEF • Hub for Extracellular Vesicle Investigations, University of Auckland, Auckland, New Zealand; Department of Obstetrics and Gynaecology, University of Auckland, Auckland, New Zealand SHAUL REUVENY • Bioprocessing Technology Institute, A*STAR Research Entities, Singapore, Singapore JAN RIEDEL • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany RUI RUAN • Centre for Orthopaedic Translational Research, Medical School, University of Western Australia, Nedlands, WA, Australia MAXIMILIAN SCHERNER • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany ANGE´LIQUE SCHMID • Ichnos Sciences SA, La Chaux-de-Fonds, Switzerland OLIVER SPADIUT • Research Division Integrated Bioprocess Development, Institute of Chemical, Environmental and Bioscience Engineering, TU Wien, Vienna, Austria EZGI ROJDA TAYMAZ • Department of Bioengineering, Faculty of Engineering, Manisa Celal Bayar University, Manisa, Turkey MISHA TEALE • Centre for Biochemical Engineering and Cell Cultivation Techniques, Institute of Chemistry and Biotechnology, Zurich University of Applied Sciences, W€ adenswil, Switzerland GERINE TONG • Bioprocessing Technology Institute, A*STAR Research Entities, Singapore, Singapore KUTLU O. ULGEN • Biosystems Engineering Laboratory, Department of Chemical Engineering, Bogazici University, Istanbul, Turkey

xii

Contributors

MEHMET EMIN USLU • Department of Bioengineering, Faculty of Engineering, Manisa Celal Bayar University, Manisa, Turkey PRIYA VELUSWAMY • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany PATRICK VETSCH • Ichnos Sciences SA, La Chaux-de-Fonds, Switzerland MAX WACKER • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany HEIKE WALLES • Core Facility Tissue Engineering, Otto-von-Guericke University Magdeburg, Magdeburg, Germany MAJID EBRAHIMI WARKIANI • School of Biomedical Engineering, University of Technology Sydney, Sydney, NSW, Australia KAMIL WIERZCHOWSKI • Warsaw University of Technology, Faculty of Chemical and Process Engineering, Warsaw, Poland JENS WIPPERMANN • Department of Cardiothoracic Surgery, University Hospital Magdeburg, Magdeburg, Germany MARITES T. WOON • Cell and Gene Therapy Research and Development, Lonza Inc., Rockville, MD, USA YUE XIONG • Center for Energy Metabolism and Reproduction, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China HUANYE XU • Beijing CytoNiche Biotechnology Co., Ltd., Beijing, China XIAOJUN YAN • Beijing CytoNiche Biotechnology Co., Ltd., Beijing, China XUEGANG YUAN • Department of Chemical & Biomedical Engineering, FAMU-FSU College of Engineering, Florida State University, Tallahassee, FL, USA; The National High Magnetic Field Laboratory, Florida State University, Tallahassee, FL, USA CHANGCHUN ZENG • Department of Industrial & Manufacturing Engineering, FAMUFSU College of Engineering, Florida State University, Tallahassee, FL, USA JIAN V. ZHANG • Center for Energy Metabolism and Reproduction, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China YUANYUAN ZHANG • Beijing CytoNiche Biotechnology Co., Ltd., Beijing, China MINGHAO ZHENG • Centre for Orthopaedic Translational Research, Medical School, University of Western Australia, Nedlands, WA, Australia

Methods in Molecular Biology (2022) 2436: 1–15 DOI 10.1007/7651_2021_441 © Springer Science+Business Media, LLC 2021 Published online: 06 October 2021

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines Ange´lique Schmid, Emanuel Kreidl, Martin Bertschinger, and Patrick Vetsch Abstract Bioreactors are manufactured apparatuses that allow the generation of a specific environment for the highly controlled cultivation of living cells. Originally used for microbial production systems, they have found widespread applications in fields as diverse as vaccine production, plant cell cultivation, and the growth of human brain organoids and exist in equally diverse designs (Chu and Robinson, Curr Opin Biotechnol 12 (2):180–187, 2001; Qian et al., Nat Protoc 13:565–580, 2018). Manufacturing of biologics is currently mostly performed using a stirred tank bioreactor and CHO host cells and represents the most “classical” bioreactor production process. In this chapter, we will therefore use the cultivation of suspension Chinese hamster ovary (CHO) cells for recombinant protein production in a stirred tank bioreactor as an example. However, general guidelines provided in this chapter are transferable to different bioreactor types and host cells (Li et al., MAbs 2(5):466–479, 2010). The preparation and operation of a bioreactor (also referred to as upstream process in a biotechnological/industrial setting) is comprised of three main steps: expansion (generation of biomass), production (batch, fed-batch, or continuous process), and harvest. The expansion of cells can last from few days to weeks depending on the number of cells at the start, the cellular doubling time, and the required biomass to inoculate the production bioreactor. The production phase lasts a few weeks and is a highly sensitive phase as the concentration of different chemicals and physical parameters need to be tightly controlled. Finally, the harvest will allow the separation of the product of interest from large particles and then the desired material (cell culture supernatant or cells) is transferred to the downstream process. The raw materials used during the upstream phase (all three steps) need to be aligned with the final purpose of the manufactured product, as the presence of residual impurities may have an impact on suitability of the final product for a desired purpose. Key words Bioreactor, Mammalian cell culture, Feeding strategy, Fed-batch, Recombinant protein production

1

Introduction For the production of complex therapeutic proteins like recombinant antibodies, mammalian cell lines have been the workhorse of the industry for three last decades [1, 2]. Chinese hamster ovary (CHO) cells in particular have found widespread application as they have a history of safe use, they exhibit a robust growth capacity with reasonable doubling times, they can be readily modified to express

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transgenes of interest, and their products show post-translational modifications compatible with in-human use [3]. A number of different modes of operation are currently employed, from microcarrier systems to different variants of perfusion processes. However, despite an increasing interest of perfusion technologies in mammalian cell culture, the classic fed-batch cultivation of suspension cell lines in stirred tank bioreactors is currently the most widely used technique, especially at larger scales [4, 5]. As a result, a number of “off-the-shelf” solutions ranging from optimized cell culture media to complete expression platforms providing the host cells, required consumables, and the recipes for bioreactor cultivation were developed. Using these technologies, secreted recombinant proteins can be produced with relative ease in volumetric concentrations ranging from hundreds of milligrams per liter for difficult to express proteins to several grams per liter of culture medium for easy to express proteins, making the approach attractive both for pure research and biotechnological use. The workflow for cell cultivation in bioreactors generally starts by thawing of cells expressing a recombinant protein of interest. The generation of such cell populations is described elsewhere [6]. Cryovials of the cell populations are frozen and are referred to as cell bank. Following thawing of the cell bank, the cells are cultivated in vessels of increasing size, typically starting with small shake flasks, with the aim of maintaining the cells in logarithmic growth (the seed train) until they are available in sufficient amounts to inoculate the production bioreactor(s) [7]. In order to reduce heterogeneity in the cell population, the cells used for manufacturing of biologics are usually clonally derived, meaning a cell population started from a single cell. Such a cell population is often referred to as “clone,” although there is significant heterogeneity even in such a population. Cells are then cultivated under tightly controlled conditions for 1–3 weeks with nutrients added in the form of feeds either at pre-defined intervals or based on consumption by the cells. Animal derived component free (usually abbreviated ADCF or ADF) and fully chemically defined media (usually abbreviated CD) and feeds have largely replaced previous media including bovine or calf serum or hydrolysates, as they reduce the risk of introduction of pathogens, show lower lot-to-lot variability and simplify downstream purification and testing [8]. In recent years, single-use technologies have found widespread application both in production and research as they allow to skip time- and labor-intensive cleaning and sterilization steps, reduce turnaround time for equipment and aim to improve reproducibility [9]. Importantly, the risk of crosscontamination with material of a previous batch is completely removed. The general operation and the required equipment only differ in minor details between single- and multi-use approaches (see Note 1).

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

2 2.1

3

Materials Equipment

Laminar flow hood. Shaking CO2 incubator. Autoclave. Cell counting system (either automatized or microscope). Single- or multi-use bioreactor (see below). Bioreactor control unit (see below). Metabolic analyzer (see Note 2). Dissolved oxygen and pH probes (see below). Weldable and silicone/bioprene tubing, clamps, connectors (see Note 3). Tubing welder (see Note 4). A source of pressurized air, oxygen and CO2. pH meter. Gas analyzer (pCO2 measurement).

2.2

Bioreactor

2.3 Cell Culture Media, Feeds, and Supplements

Glass, stainless steel as well as single-use (plastic) bioreactors are available in a large range of sizes and designs from various manufacturers, which has to be considered when many runs are planned in a short time frame [10]. The exact design characteristics of a bioreactor combined with the particular setpoints (rpm of the impeller, gassing strategy) have a significant impact on various parameters influencing the behavior of cells from shear stress to mass transfer coefficients and potential dead zones [11, 12]. Literature on the subject is extensive with the described findings needing to be verified for the particular setup, application, and/or clone. Bioreactors using pitched blade impellers and macrosparger are often successfully used in bench top (i.e., less than ~10 L working volume) setups used for mammalian cell cultivation. See Fig. 1 for a representation of a benchtop bioreactor. Basal media formulations allow the expansion of the cells from the cryovial to inoculation of the bioreactor. They also allow a shortterm cultivation of cells in the bioreactor. Feeds are concentrated nutrient solutions added during the process to replenish consumed media components. They allow to cultivate cells to high cell densities of tens of millions of cells per mL but also aim to drive cells towards a metabolism favorable for production and secretion of the protein of interest [13]. Highly potent cultivation media and feeds are nowadays available from a number of different suppliers. Small quantities are usually ordered in form of chemically defined liquid formulations.

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Fig. 1 (a) Sampling port—allows for the removal of cells and culture liquid for analysis using a syringe. The risk of contamination can be reduced by spraying the port with ethanol before and after sampling. (b) Filters attached to gas inlet and exhaust. Care needs to be taken to prevent moisture from accumulating on the filters which would lead to clogging. (c) Headplate with connections for liquid addition and ports for probe insertion. Missing of damaged O-rings on the probes result in a high contamination risk. (d) Impeller—various designs are available resulting in different mixing characteristics and shear stress to the cells. Shown here: Rushton type impeller more commonly used for microbial applications. (e) Sparger—allows for efficient transfer of gases to the cell culture liquid. Different designs result in different bubbles sizes greatly influencing gas transfer and shear stress. Shown here: macrosparger

For larger quantities, powder formulations may be considered. Following the protocol of the provider, a liquid formulation can then be generated by the addition of cell culture grade water. Generally, media and feeds were designed simultaneously to enable the cellular metabolism to secrete high quantities of recombinant protein. Usually, such medium feed combinations are provided with basic protocols describing cultivation conditions and timing of feed addition, allowing to quickly obtain reasonable results. Media and feeds can have a major impact on the performance of a specific cell population with regard to doubling time, production level, and product quality [14]. It may thus be advisable to perform

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

5

a pre-screening for best conditions at a smaller scale like shake flasks (SF) or spin tubes prior to advancing to the bioreactor stage [15] (see Note 5). Furthermore, it is often necessary to provide additional nourishment in the form of a concentrated glucose (e.g., 45% w/v) solution to keep this nutrient from being fully depleted (see Note 6). Several CHO-derived cells require the addition of glutamine. Depending on the cell type and the way the gene of interest was introduced, additional metabolic supplements may be required (HAT medium, Folate). Due to the additional stress that the cells may be exposed to in bioreactors, supplementation of the media with a shear protectant such as Pluronic might be required [16]. 2.4

Antifoam

Generally, antifoaming agents need to be added to the bioreactor to control the level of foam generated by stirring and gas addition through the sparger [17]. As these chemicals can have a negative impact on the cells and downstream purification steps, their addition should be limited to the minimum necessary [18] (see Note 7).

2.5

Controller

Controllers are available from many different manufacturers and can often be combined with equipment (bioreactor units, probes) from another supplier. While these devices can be operated in different modes with various degrees of complexity and automation, their core function is to integrate readouts from the bioreactor, like vessel temperature and amount of dissolved oxygen (DO) and adjust outputs to reach a particular setpoint. To provide an example, a specific setpoint in DO can be maintained by changing the rotation speed of the impeller or the amount or composition of the injected gasses (or both), until the desired DO has been reached. Furthermore, controllers allow for the automatic addition of feeds and other additives and the control of the pH as well as changes in cultivation temperature at a pre-defined time or based on process data (see Note 8). Table 1 is meant to serve as a general guideline and starting point for operation parameters.

2.6

Probes

Levels of dissolved oxygen (DO) and pH are typically measured by probes inserted into the bioreactor. They are available using different technologies and in single- and multi-use format. The latter need to be cleaned and sterilized prior to use and might require additional calibration and quality control steps at regular intervals. Conditions for long-term storage also need to be considered. Some manufacturers of single-use bioreactors now also offer pre-installed measurement capabilities which however usually require dedicated equipment [19]. In addition, a number of other technologies

Ange´lique Schmid et al.

6

Table 1 Summary table of important parameters Parameter type

Parameter

Ranges/guidelines (for a 3 L culture)

Physical parameters

DO percentage

20–50% of dissolved oxygen in the culture (see Subheading 3.4, step 4) In headspace around 100 ccm and through a macrosparger 50 ccm (see Subheading 3.4, step 2) 5–8% in headspace for equilibration. Could be used in sparger to regulate the pH (see Subheading 3.4, step 2) 37  C could be shifted down (see Subheading 3.4, step 1) Around 200 rpm depending on the coupling system (see Subheading 3.4, step 3) Around 6.7–7.5 (see Subheading 3.4, step 5) Will be based on the final bioreactor volume minus the volume of feeds and inoculum. Pay attention that this volume still needs to be above a limit defined by the supplier mainly to guaranty immersion of probes and the complete immersion of the impeller to control generation of foam (see Subheading 3.5, step 1) pCO2 level needs to be carefully monitored especially during the scale-up. Sensitivity to this parameter will differ from a cell line to another, usually 40–50 mmHg [27]

Air injection CO2 injection Temperature Agitation pH regulation Initial volume

pCO2

Biological parameters

Biomass

Metabolites concentration/ consumption Feeding strategy

Feeds percentage

Addition strategy

Key components adjustment Foam

Foam

Initial VCC to be determined regarding process performances (0.3 to multiple million per mL in case of perfusion) (see Subheading 3.6, step 2) Carbon source (e.g., glucose) and by-products (e.g., lactate, ammonia) need to be carefully monitored (see Note 2) Depending on the cell line and produced molecule, feeds percentage could vary. An experiment to determine the best strategy should be performed in advance. Ranges to be tested are usually provided by the supplier (see Subheading 2.3) As for the percentage, addition strategy will be based on the cell’s needs and need to be tested (see Subheading 2.3) Additional feeding of key metabolites in the culture could be made on top of feeds addition (see Note 6) Foam level needs to be controlled to avoid clogging of exhaust filter (see Subheading 3.7, step 4)

ranging from capacitance measurements to Raman spectroscopy and enzymatic glucose measurements have become available in probe format allowing to follow numerous parameters online [20]. 2.7 Other Supplements

pH regulation could be performed by addition of acid or base, typically HCl and NaOH (see Note 9).

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

3 3.1

7

Methods Cell Expansion

3.2 Probe Calibration and Sterilization

1. Vials of frozen cells are thawed and the cells are expanded to have enough biomass to inoculate the fed-batch bioreactor. Usually, expansion is performed using shake flasks or other vessels of increasing size. Duration between passages is 2–4 days. Initial VCC (Viable Cells Concentration) for passages will depend on the passage duration and is typically in the range of 0.3–1 million cells per mL (see Notes 10 and 11). 1. When not using (pre-installed) single-use probes, pH probe calibration is performed prior to sterilization (see Note 12). 2. For DO probes, it is advisable to confirm that they are in proper working condition (typically by checking the electric signal in air) before the sterilization (see Note 13). 3. Depending on the type of measurement system and/or bioreactor used, the same might need to be performed for temperature- and other probes. 4. Probes are then packaged in autoclaving bags and sterilized (see Note 14). In case of glass vessel use, probes could be inserted before the sterilization of the bioreactor and sterilized with it (see Note 15).

3.3 Bioreactor Preparation

1. In case of single-use bioreactors, pre-sterilized probes need to be inserted aseptically. Clamp all lines and remove protection to insert both DO and pH probes under a laminar flow hood. 2. For both single-use and glass vessels, transfer the bioreactor next to the control tower. Connect the overlay gas line to the filter, same for the sparger line. Unclamp the exhaust gas line if necessary. Add air (at a constant minimal flow) through the overlay to maintain an overpressure inside the bioreactor. 3. Connect pH and DO probes to the control towers and confirm proper signal transmission (see Note 16). 4. Put in place the motor (see Note 17), temperature probe, and electrical heating blanket/heating system around the bioreactor. Controlling the bioreactor temperature with a doublejacket filled with pre-heated water is also possible for glass vessels. 5. To avoid clogging of the exhaust filter due to humidity of the exhaust gas, a filter heater or condenser around the exhaust system needs to be applied.

3.4 Bioreactor Setpoints

1. The temperature should be tightly controlled within the bioreactor. Usually a setpoint of 37  C is used for CHO cells. Modification of this setpoint during the culture (typically

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towards a lower temperature) can be useful with a shift to 34 or 32  C at a pre-defined timepoint being a widespread approach [21]. 2. The ability to inject gases in response to the demands of the culture is one of the main advantages of using a bioreactor. Injection of (a combination of) air, oxygen, and CO2 into the headspace serves to adjust pH of the media, limit foam accumulation and may fulfill the initial oxygen requirements of the cells. It is typically not sufficient to do so once higher cell densities have been reached. Instead, gas needs to be injected directly into the culture medium. Spargers of different design exist with open pipe-, macro- (such as a ring sparger) and micro-spargers being the most popular ones. The choice of sparger needs to be made based on the oxygen requirements of the cells, sensitivity to bubble bursting, and potential equipment limitations (capacity of the gas source). Micro-spargers provide significantly better aeration due to the much higher total surface volume of the large amount of small bubbles. Conversely, this results in higher stress for the cells (see Note 18). 3. Stirring is important to ensure homogeneity of the culture and to avoid the formation of gradients within the bioreactors. Stirring also has an impact on shear stress. The appropriate agitation rate needs to be determined based on the system. Engineering parameters such as the maximum tip speed, mixing time, and the power input per volume can support the selection of these parameters [22]. 4. Establishing a regulation cascade for dissolved oxygen ensures that enough of this key substrate for growth and production is available to the cells independent of their density and metabolism. Typical setpoints are between 20 and 50% (compared to concentration in the medium equilibrated with air under normal pressure). This parameter is typically controlled through agitation rate and/or injection of air or oxygen through the sparger. The cascade needs to be well defined to ensure the setpoint can be met and to avoid spikes (see Note 19). 5. Active or passive pH regulation can be used. Cell culture pH is commonly between 6.7 and 7.5. With a passive approach, pH of the culture is usually dropping down during the first few days due to the generation of acidic by-products. Shifts in metabolism and addition of feeds then typically limit a further drop in pH. Active pH control injecting CO2 in the headspace or in the culture medium or removal of CO2 using sparging (in case of small adjustments) or acid/base addition can be used to maintain pH at a desired setpoint. The impact of the latter on the osmolality of the culture should not be overlooked.

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

3.5 Media Preparation/Hold

9

1. Basal culture media should be aseptically transferred into the bioreactor. Shear protectant can be added to reduce stress to the cells during the culture. When determining the amount of basal media to be used, it is important to consider the maximum working volume of the bioreactor (usually around 2/3 of the vessel volume) as well as the volume added by the inoculum and by addition of feeds (up to ~40% of the final volume) and acid/base (if applicable). Depending on the percentage of volume removed for sampling during the process, it might be required to adjust the feeding volume accordingly (see Note 24). 2. Before the inoculation, it is recommended to equilibrate media inside the bioreactor. Temperature, as well as DO and pH should reach their equilibrium or setpoint [23] (see Note 20). 3. DO probes should be calibrated with two points using air saturated media as the upper point and “oxygen free media” (either by removal of oxygen using nitrogen sparging or mimicking this by disconnection of the probe) as 0% point (see Note 21). 4. pH probes should be re-calibrated using an offline measurement as autoclavation can have an impact on the calibration. 5. Feeds, as well as other supplements, need to be aseptically connected to the bioreactor. This can also be delayed until the actual addition of the feed (s) is required (see Note 3).

3.6

Inoculation

1. At the time of inoculation, pH, pCO2, and metabolite concentrations should be carefully checked and potentially regulated to ensure a correct environment allowing cells to grow. Typical physical parameters listed in Table 1 should be observed at this particular step of a fed-batch process. 2. After counting cells from the expansion, the required amount of inoculum to reach the target cell concentration is added to the bioreactor. Adjusting the concentration of the cells to a pre-determined value so that the same volume can be transferred for each inoculation simplifies this process and increases reproducibility (see Notes 22 and 23). 3. Post-inoculation sampling is performed when the culture is homogenized (after around 30 min). Homogeneity of the bioreactor is important to ensure a representative sample is obtained and to determine initial condition of the culture. If the target seeding density is not reached, cells or media (depending if the VCC is too high or too low) could be added. In this case, the increase of the initial bioreactor volume should be taken into account and the feed volume should be adapted.

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3.7 Fed-Batch (Sampling and Feeding)

1. During the culture, which usually lasts around 2 weeks, sampling is done every day to acquire process understanding and to allow comparison with other bioreactors (see Note 24). 2. After a few days, some of the nutrients contained in the basal media are completely consumed by the cells, raising the need to add new ones. Thus, feed addition typically starts around day 2 to day 4. The appropriate addition strategy should be selected to supply enough nutrients to cells and allow them to both grow and produce the product of interest. Overfeeding should be avoided as it would lead to critical sub-optimal performances. Thus, an appropriate balance is required. The starting time for feed addition, the type and number of feeds as well as the daily quantities are typically the parameters that need to be studied to develop an appropriate feeding strategy (see Note 25). 3. Multiple process parameters can be critical to ensure a robust and efficient production process. Among them, the glucose concentration is paramount. Indeed, a shortage of glucose is likely to quickly lead to cell death. This means that specific control over those parameters might be required. 4. During the culture, foam is likely to accumulate due to the sparging. Controlling the level of foam is of great importance to avoid clogging of the exhaust filter (see Note 26). 5. The fed-batch run is terminated either at a particular timepoint, or when a minimal cell viability (typically between 50 and 75%) is reached.

3.8 Harvest and Clarification

4

1. When the aim of the bioreactor run is to obtain the (product containing) culture media, a final step to separate cells and culture liquid is required. Depending on the scale and equipment availability, this is usually done by centrifugation or depth filtration or a combination of these. In addition, proper inactivation and disposal of biological material in accordance with local regulations is essential.

Notes 1. Considerations for the choice and optimal operation of a bioreactor are numerous and complex and well beyond the scope of this publication. The aim is to provide a starting point for testing and optimization. Detailed description of operation parameters can be found in publications dedicated to this subject [10] as well as textbooks. 2. Metabolite analyzers of different complexity are available from several suppliers. As a minimum requirement, a method for the

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

11

measurement of dissolved glucose levels should be at hand. This can for example be performed with an over-the-counter glucose meter. Lactate, ammonia, and other metabolites such as glutamate and glutamine could be of interest to be measured to further optimize the feeding strategy based on these compounds (to keep their concentration low or the supply enough in case they are important for the culture). 3. Tubing will be required to connect bottles or other containers with feeds, antifoam, and glucose as well as to transfer the cells for the inoculation. Usually weldable tubing (thermoplastic elastomer tubing) is not suitable for pumping operations which should be done using silicone or bioprene. Silicon tubing however cannot be welded so adaptors are required to connect to the weldable tubing. Bioprene could be welded but is expensive, especially since lines to connect the bioreactor to the vessels containing, e.g., feeds passing through the pump can be quite long. Use of another weldable tubing will reduce global cost. Especially single-use and/or larger bioreactors that cannot easily be moved require significant planning before operations in order to establish all connections in a sterile and efficient manner. 4. Tubing welders allow to quickly establish sterile connections to the bioreactors used for addition and removal of media and cells. Their use is applicable to connect vessel which required to be connected after bioreactor preparation, either because they could not be sterilized with the bioreactor (sensitive to heat), because the line will be used for another purpose first or because the liquid to add needs to be connected at a defined time (inoculum, material sensitive to light or heat). Alternatively, tubing assemblies with sterile single-use connectors are available from a number of suppliers. Otherwise and especially when glass bioreactors are used, connections can be established prior to addition of the media under non-sterile conditions followed by autoclaving. Containers with heat sensitive components like media can then be attached inside a lamina flow hood. Connection of tubing assemblies under the lamina flow hood could be done at the time of bioreactor preparation. 5. Not all media and feeds are compatible. In particular, several manufacturers provide two separate feeds that precipitate if mixed undiluted due to the pH required to solubilize the ingredients at high concentrations. Thus, care has to be taken to avoid mixing of incompatible components for example if only a single feed line is used. 6. Often, the amount of glucose added is based on the difference between the remaining glucose level in the bioreactor

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(determined by daily measurement) and a pre-defined setpoint in the range of the glucose concentration in the basal media (typically between 3 and 10.5 g/L). However, high glucose concentration could negatively impact culture performances and appropriate setpoints should be defined based on culture needs [24]. 7. Since many of the agents are based on an emulsion of silicone beads, sterile filtration is generally not possible. Some can be sterilized by autoclavation, while others are only applicable for bioreactor use when purchased in a pre-sterilized (gammairradiated) format. 8. Depending on available equipment and process understanding, possibilities for automatization and detailed control are almost endless. Where limitations exist, many functions of a controller can be substituted by using external equipment like pumps and scales or placing the bioreactor inside of an incubator. Control of the DO levels by defined sparging of gas however should be considered as an essential functionality of any control equipment. 9. While often used, control of the culture pH by base addition is not always required and omitting it reduces stress the cells experience from increased osmolality [25]. 10. During all handling steps involving suspension cells, great care needs to be taken to properly homogenize the liquid culture so that the counted and transferred materials are aligned. 11. It is advisable to passage the cells several times after thawing, not only to obtain sufficient material, but also to give them time to fully recover. Furthermore, it is essential to maintain cells in logarithmic growth as both too high and too low cell densities can negatively influence stability of the cell line and performance in subsequent steps. 12. At least a two-point calibration of the pH probes should be performed using reference solutions with values similar to what would be expected in the bioreactors (usually 7.0  1.0). As in most systems the calibration is specific to a particular combination of pH probe and controller, care should be taken to perform this step for the equipment pair being used during the run and to label devices accordingly. 13. A two-point calibration of the DO probe will need to be performed in the bioreactor when the media is equilibrated. 14. When using paper autoclaving bags, double bagging is advised and probes should quickly be moved to a sterile zone after removal from the autoclave as wet paper is permeable and might promote contamination.

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines

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15. Appropriate recording of usage of both DO and pH probes is highly recommended as each sterilization could impact their performances. 16. During the first check of the pH, the probe signal value could be off, this is because the probe is in air and not in media. 17. In case of magnetically driven systems, be careful with regard to the speed and the strength of the magnet as decoupling can occur. 18. The topic of gas transfer in bioreactors has been extensively studied and is highly complex, in particular when it comes to adjustment between scales. The values provided in Table 1 can only serve as a starting point and will certainly need adaption for a particular cell line and equipment. 19. DO setpoints can be reached by letting cells consume oxygen present in the media or by sparging nitrogen in the media. A constant air flow through the sparger usually helps the control of DO percentage at the beginning of the process. 20. If extended storage of media before addition is required, pay attention to the significant light sensitivity of many cell culture media and limit exposure to temperatures above 2–8  C as much as possible. Depending on the volume of media and mixing time of the system, equilibration time could last from 1 h to multiple hours. An equilibration period of more than 1 day could also help to detect a potential contamination before the inoculation. 21. When using CO2 (typically 5–8%) during the equilibration step, this needs to be taken into account for the calibration of the DO probe, resulting in an initial calibration value of 92–95%. 22. Careful manipulation of the inoculum is critical as cells are quite sensitive to shear stress. Also, during the transfer cells are not well oxygenated and the temperature is difficult to maintain. This step should be performed as quickly as possible. If the transfer is performed using a peristaltic pump, the liquid flow must be low in order not to damage cells. 23. The initial seeding cell density as well as split ratio have a substantial impact on the performance of the culture and have received increased attention especially in the context of process intensification [26]. This topic is furthermore complicated by the fact that equipment availability will have a strong impact on potential initial cell densities and state of the inoculum (amount of fresh media, DO), especially at larger scales. As with many setpoints, performing a round of experiments in a less resource intensive format like shake flask to assess general trends is highly advisable.

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24. This is dependent on the devices used, the scale (how big an impact in terms of culture volume the sampling might have), and sampling frequency. In particular for low volume bioreactors, the loss of volume due to sampling can have an impact on overall performance. 25. Mainly two feeding modes are used: Continuous feeding, which means constantly adding a small amount of feed (s) once the feed addition has been initiated using a low flow pump (this is usually performed using a connected scale to be able to control the amount added in 24 h). Alternatively, the feeds are added in bolus mode, meaning that feeds are added at a pre-defined interval (usually once per day once the feed addition has been initiated) as a short burst. This requires less automation as the pump could be activated by hand and stopped when the target weight is reached. Continuous feeding is smoother for the culture but requires more infrastructure to be precise. Importantly, it is generally not possible to test this approach in a less automated format like shake flask. 26. Addition of antifoam once to completely remove the foam is more efficient than putting multiple small doses. If foam is not controlled, clogging of the filter could occur.

Acknowledgments We would like to thank David Hacker for his input and support in writing this chapter and Eppendorf AG for providing images used in the creation of the figures. References 1. Chu L, Robinson DK (2001) Industrial choices for protein production by large-scale cell culture. Curr Opin Biotechnol 12 (2):180–187 2. Li F, Vijayasankaran N, Shen AY, Kiss R, Amanullah A (2010) Cell culture processes for monoclonal antibody production. MAbs 2 (5):466–479 3. Zhu J (2012) Mammalian cell protein expression for biopharmaceutical production. Biotechnol Adv 30(5):1158–1170 4. Wurm F (2004) Production of recombinant protein therapeutics in cultivated mammalian cells. Nat Biotechnol 22:1393–1398 5. Kunert R, Reinhart D (2016) Advances in recombinant antibody manufacturing. Appl Microbiol Biotechnol 100:3451–3461

6. Castan A, Schlutz P, Wenger T, Fischer S (2018) Chapter 7—Cell line development. Biopharmaceuticals processing. ISBN 9780081006238 7. Herna´ndez Rodrı´guez T, Frahm B (2020) Design, optimization, and adaptive control of cell culture seed trains. Methods Mol Biol 2095:251–267 8. van der Valk J, Brunner D, De Smet K, Fex Svenningsen A, Honegger P, Knudsen LE, Lindl T, Noraberg J, Price A, Scarino ML, Gstraunthaler G (2010) Optimization of chemically defined cell culture media—replacing fetal bovine serum in mammalian in vitro methods. Toxicol In Vitro 24(4):1053–1063 9. Shukla AA, Gottschalk U (2012) Single-use disposable technologies for biopharmaceutical

Benchtop Bioreactors in Mammalian Cell Culture: Overview and Guidelines manufacturing. Trends Biotechnol 31 (3):147–154 10. Junne S, Neubauer P (2018) How scalable and suitable are single-use bioreactors? Curr Opin Biotechnol 53:240–247 11. Xu S, Hoshan L, Jiang R, Gupta B, Brodean E, O’Neill K, Seamans TC, Bowers J, Chen H (2017) A practical approach in bioreactor scale-up and process transfer using a combination of constant P/V and vvm as the criterion. Biotechnol Prog 33(4):1146–1159 12. Sandadi S, Pedersen H, Bowers JS, Rendeiro D (2011) A comprehensive comparison of mixing, mass transfer, Chinese hamster ovary cell growth, and antibody production using Rushton turbine and marine impellers. Bioprocess Biosyst Eng. 34(7):819–832 13. Reinhart D, Damjanovic L, Castan A, Ernst W, Kunert R (2018) Differential gene expression of a feed-spiked super-producing CHO cell line. J Biotechnol 285:23–37 14. Pan X, Streefland M, Dalm C, Wijffels RH, Martens DE (2016) Selection of chemically defined media for CHO cell fed-batch culture processes. Cytotechnology 69(1):39–56 15. Reinhart D, Damjanovic L, Kaisermayer C, Kunert R (2015) Benchmarking of commercially available CHO cell culture media for antibody production. Appl Microbiol Biotechnol 99(11):4645–4657 16. Hua J, Erickson LE, Yiin TY, Glasgow LA (1993) A review of the effects of shear and interfacial phenomena on cell viability. Crit Rev Biotechnol 13(4):305–328 17. Pelton R (2002) A review of antifoam mechanisms in fermentation. J Ind Microbiol Biotechnol 29(4):149–154 18. Velugula-Yellela SR, Williams A, Trunfio N, Hsu CJ, Chavez B, Yoon S, Agarabi C (2018) Impact of media and antifoam selection on monoclonal antibody production and quality using a high throughput micro-bioreactor system. Biotechnol Prog 34(1):262–270

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19. Demuth C, Varonier J, Jossen V, Eibl R, Eibl D (2016) Novel probes for pH and dissolved oxygen measurements in cultivations from millilitre to benchtop scale. Appl Microbiol Biotechnol 100(9):3853–3863 20. Maruthamuthu MK, Rudge SR, Ardekani AM, Ladisch MR, Verma MS (2020) Process analytical technologies and data analytics for the manufacture of monoclonal antibodies. Trends Biotechnol 38(10):1169–1186 21. Xu J, Tang P, Yongky A, Drew B, Borys MC, Liu S, Li ZJ (2019) Systematic development of temperature shift strategies for Chinese hamster ovary cells based on short duration cultures and kinetic modeling. mAbs 11(1):191–204 22. Odeleye AO, Marsh DT, Osborne MD, Lye GJ, Micheletti M (2014) On the fluid dynamics of a laboratory scale single-use stirred bioreactor. Chem Eng Sci 111(100):299–312 23. Michl J, Park KC, Swietach P (2019) Evidencebased guidelines for controlling pH in mammalian live-cell culture systems. Commun Biol 2:144 24. Goldrick S, Lee K, Spencer C, Holmes W, Kuiper M, Turner R, Farid SS (2018) On-line control of glucose concentration in highyielding mammalian cell cultures enabled through oxygen transfer rate measurements. Biotechnol J 13(4):e1700607 25. Hoshan L, Jiang R, Moroney J, Bui A, Zhang X, Hang TC, Xu S (2018) Effective bioreactor pH control using only sparging gases. Biotechnol Prog 35(1):e2743 26. Xu S, Gavin J, Jiang R, Chen H (2017) Bioreactor productivity and media cost comparison for different intensified cell culture processes. Biotechnol Prog 33(4):867–878 27. Matanguihan R et al (2001) Solution to the high dissolved CO2 problem in high-density perfusion culture of mammalian cells. In: Lindner-Olsson E, Chatzissavidou N, Lu¨llau E (eds) Animal cell technology: from target to market. ESACT proceedings, vol 1. Springer, Dordrecht

Methods in Molecular Biology (2022) 2436: 17–25 DOI 10.1007/7651_2021_415 © Springer Science+Business Media, LLC 2021 Published online: 10 August 2021

Volumetric Mass Transfer Coefficient Measurement in a Stirred Tank Reactor Aysegul Inam, Ezgi Rojda Taymaz, Mehmet Emin Uslu, Baris Binay, and Irem Deniz Abstract A bioreactor is a controlled vessel which provides biological conversions into bioactive components using cells or enzymes. In the aerobic processes, it is important to know oxygen requirements of the cells which may vary during fermentation as a result of microbial activity, aging, substrate depletion and product formation, etc. Here we describe the measurement of volumetric mass transfer coefficient (kLa) in a stirred tank reactor using dynamic method based on unsteady state which is also one of the significant parameter especially in scaling-up. The equipment in the measurement according to dynamic method has low cost compared to steady-state methodology. This method is reliable in the determination of kLa when the gas residence time and probe measuring the oxygen concentration of response time are in specific requirements. Key words Aerobic processes, Dissolved oxygen concentration, Dynamic method, Stirred tank reactor, Volumetric mass transfer coefficient

1

Introduction Gas transfer is defined as the process in which gas moves from one phase to another. This phenomenon is used in mass transfer and oxygen transfer for the continuity of biological processes during bioproduction in bioreactors. The transition of oxygen from gas phase to liquid phase is crucial for aerobic process especially for: enzyme production, biomass related product formation, wastewater treatment, and animal cell cultures [1]. Aerobic microorganisms in fermentation broth produce the energy required for their survival and growth by using dissolved oxygen molecules. Oxygen, like all gases in the atmosphere, dissolve in water to a certain degree [2]. Dissolved oxygen (DO) concentration is highly depended on temperature, pressure, and salinity level of the water. Maximum DO concentration at 1 atm and room temperature is 8.26 mg/L for pure water [3]. In bioprocesses where aerobic microorganisms are involved, it is very important to maintain the desired level of DO concentration in the fermentation broth [4]. Several organisms need very high

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levels of oxygen for biomass-related product formation. This requires continuous measurement of DO concentration in bioreactors, and a control system that will react quickly when there is a deviation from the values designed according to the needs of microorganisms. Because the number of microorganisms in the fermenter and their metabolic rate changed over time, the oxygen requirement of the system also does not remain constant during fermentation [5, 6]. 1.1 The Measurement of kLa

DO concentration can vary during fermentation as a result of microbial activity. Physical and chemical methods have been proposed for the measurement of the kLa in stirred tank reactors (STR). In bioprocesses, steady-state and unsteady-state methods are mostly preferred. These methods have superior advantages for several cases according to their application.

1.2 The UnsteadyState (Dynamic) Method

The unsteady-state (also known as dynamic) technique is the most used method for measuring DO concentration. It is based on monitoring the decrease in DO concentration with an oxygen electrode by cutting off the air (oxygen) that feeds the bioreactor, and the increase in oxygen concentration by re-feeding the air (oxygen). Basically, the DO level is foamed with nitrogen or reduced to zero by adding sodium sulfide. Then, the increase in DO concentration as a function of time is followed [7]. The unsteady-state method consists of two stages: consumption and absorption. Aeration is stopped and DO decreases due to cell respiration during consumption stage. In absorption stage, aeration is resumed, and the DO increases until a steady-state is reached. It is often difficult to obtain accurate kLa using the unsteady-state method. Because, first of all the assumptions about the extent of gas-phase mixing must be chosen correctly. Secondly, the electrode response time must be fast [7, 8]. However, it is commonly used because it gives more accurate result. During the transfer of oxygen from air to water, the actual resistance occurs in the liquid film layer at the interface [2, 3]. Inside the bioreactor where oxygen is not consumed, when resistance in the gas phase is neglected, the time-dependent change of DO concentration is given by q O x ¼ kL aðC ∗  C L Þ ¼ OTR

ð1Þ

where kL is the oxygen transfer coefficient (cm/h), a is the gas liquid interfacial area (cm2/cm3), kLa is the volumetric mass transfer coefficient (h1), C∗ is saturated dissolved oxygen concentration (mg/L), CL is the actual dissolved oxygen concentration in the broth (mg/L), and the qOx is the rate of oxygen transfer (mg O2 L1 h1).

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The system in the STR is not at a steady-state in the course of re-oxygenation. In this step, the rate of DO concentration change is equal to oxygen transfer rate from gas into liquid. If the rate of DO concentration change with time is equal to zero and final DO concentration as steady value is equal to CL (C∗ ¼ CL), an expression could be obtained [9]. 1.3 The Steady-State Method

2

Steady-state methods are based on a chemical or biochemical reaction that occurs in the liquid, which acts as an oxygen absorber. Steady-state method is based on the oxygen flow rate and the difference between the input O2 concentration and the output O2 concentration. This approach allows the kLa to change over time with a reduction in oxygen concentration. In other words, this method uses the global oxygen balance in the reactor’s gas phase. Some notable errors may occur due to the small difference in O2 concentration between the inlet and outlet gas streams [9, 10]. kLa, is a measure of the use of oxygen in gas–liquid systems. kLa value is very important in the synthesis of products and biomass, since metabolically obtaining energy is related to oxygen produced by ventilation systems. There are many variables that are effective in kLa value, also known as aeration efficiency. These parameters are mainly; turbulence in the ventilation pool, i.e., mixing, air pressure, air flow rate, temperature, properties of the fluid (density, viscosity, etc.) and the availability of foaming inhibitors [11].

Materials The bioreactor type is chosen as stirred tank reactor with two baffles for this protocol. The capacity of the STR is 3 L with a working volume of 2 L (see Note 1).

2.1

Bioreactor

1. Sparger: Orifice type. 2. Impeller: 6-blade Rushton turbine. 3. Air pump: Use with silicone pipes. 4. Dissolved oxygen probe: Polarographic type. 5. Temperature detector. 6. Sampling port. 7. Control unit of bioreactor: Follow the temperature, dissolved oxygen concentration, aeration (gas flow rate), and agitation (stirrer speed). 8. Nitrogen tank: Use and arrange with pressure regulator.

2.2

Microorganism

Get the stock culture of Escherichia coli as a model organism for aerobic production in bioreactor (see Note 2).

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Growth Media

Prepare the Luria-Bertani (LB) growth media in broth and agar forms for E. coli with the following components for 1 L: 10 g tryptone, 10 g NaCl, 5 g yeast extract and distilled water. Add agar agar (2%) with these components for agar media (see Note 3).

2.4 Spectrophotometer

Use in following the cell concentration of E. coli at the wavelength of 600 nm.

2.5

Measure the time and also obtain the DO concentration values corresponding to those times.

3

Chronometer

Methods

3.1 Installation of the Bioreactor

1. Use the orifice shape of sparger which is responsible in entering of air bubbles into the bioreactor through air pump (see Note 4). 2. Locate the Rushton turbine port in the STR (see Note 5). 3. Install the sampling port, the temperature detector and polarographic type of DO probe inside the bioreactor. 4. Autoclave the bioreactor with all its parts placed inside for sterilization of 20 min in 121  C (see Note 6).

3.2 Preparation of the Growth Media

1. Prepare agar medium for sustaining the stock culture in petri dishes or slant tubes. Weigh the components of tryptone, NaCl and yeast agar plus agar agar according to required volumes and mix them in the distilled water. Boiled the solution for homogenization. 2. Prepare the components of broth media for scale up from agar petri dishes to end inoculation volume of bioreactor. Start with amounts of 5 mL of tubes and keep going with 10, 20 and 40 mL of volumes, respectively. Weigh the all components except agar agar and mix them in the distilled water in the required quantities (see Note 7). 3. Prepare separately the growth medium for bioreactor in 2 L of volume. 4. Measure the pH values of growth media and adjust the pH to range between 7.5 and 8 (see Note 8). 5. Autoclave the all growth media and equipment for sterilization of 20 min in 121  C. 6. For agar medium: Shortly after sterilization, pour the broth medium in aseptic conditions into sterile petri dishes/tubes while at medium temperature of averagely 60  C and allow it to solidify by cooling.

Volumetric Mass Transfer Coefficient Measurement

3.3 Preparation of Microorganism

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1. Inoculate the E. coli cells on agar media by inoculation loop in aseptic conditions, firstly. Culture the cells at 37  C. 2. After colonies grown on petri, take a single colony from petri and transfer into 5 mL of broth media. Scale up until 40 mL of broth media in 250 mL of Erlenmeyer with following the culture grown (see Notes 9 and 10). 3. Use the 40 mL of culture to inoculate bioreactor.

3.4 Operation of the STR

1. After the sterilization process, transfer the sterile growth medium into bioreactor in aseptic conditions. 2. Inoculate 2% (v/v) of the cells corresponding to 40 mL into the bioreactor in aseptic conditions. 3. Connect the air pump to head of sparger port with silicone pipe. 4. Operate the STR under suitable conditions at 150 rpm and 1 vvm (37  C) with following from the control unit of bioreactor (see Notes 11 and 12).

3.5 Measurement of the Oxygen Transfer Coefficient: kLa

1. Check continuously the optical density of cells for following the growth. 2. After the cells reach to logarithmic growth phase, be ready for determination of kLa through dynamic method in bioreactor (see Note 13). 3. For de-oxygenation of the broth at time, connect the nitrogen tank (see Note 14). 4. Sparge nitrogen gas into the bioreactor for displacing with oxygen. 5. Use DO probe and monitor the changes in DO concentration (see Notes 15 and 16). 6. In this period, C drops (Fig. 1). 7. After observing the value of DO as 0% saturation (or near %0), cut the nitrogen supply (see Notes 17 and 18). 8. After C drops, aerate the fermentation broth through pumping at specific operating conditions like a known constant air flow rate (see Note 18). 9. As a function of time, C increases. Monitor this change in DO concentration with following the air inflow start (Fig. 1). 10. Measure C values in different times in the bioreactor and record these several values with respect to time using chronometer for the plot (see Note 19). 11. Assume that the re-oxygenation of the fermentation broth is fast compared to cell growth, so that reaching of level of DO to a steady-state value will be shortly. C* which is the steady-state value shows an equilibrium between supply and consumption of the oxygen (see Note 20).

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Fig. 1 The plot of C values at different time intervals. C1, oxygen concentration measured at t1 time during reoxygenation; C2, oxygen concentration at t2 time during reoxygenation

Fig. 2 Determination of kLa value based on the linearized plot of the logarithmic values of DO concentrations. The slope of the obtained plot is directly proportional to the kLa value

12. When the necessary requirements are fulfilled such as substitution and integration in the equation, estimate accurately the kLa value with plotting the logarithmic values of (C*  C1)/ (C*  C2) versus time interval (t2  t1) (Fig. 2). 13. Take the slope of this straight line plot as kLa value.

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Notes 1. Types of bioreactor based on geometry and other design parameters could be bubble column, air-lift, fixed-bed column, and stirred tank reactors. Dynamic method could not be correctly applicable in high bioreactors with height more than 1 m due to gas residence time in de-oxygenation step during nitrogen hold-up. 2. Any microorganism other than E. coli can also be used to determine the kLa measurement. However in this case, be aware of choosing the right growth medium and physical conditions that the microorganism requires. 3. Depending on types of microorganisms, other growth media that provides macro and micro elements for cells can also be selected for measurement of the kLa. 4. The sparger type of bioreactor could be porous or nozzle beside orifice type. 5. Microorganisms transform substrates into biological products with the advantages of mixing and consequently heat-mass transfer mechanisms. To provide mixing, 6-blade Ruston turbine impeller is used for bacterial production in this protocol. Marine, paddle, anchor or helical ribbon types of impellers could also be chosen in terms of microorganism types and production purposes. 6. Be aware that the STR and LB medium are autoclaved separately to ensure the loss of volume in the STR is minimized as a result of evaporation. 7. When the growth medium is to be used, prepare it fresh or prepare it not more than 3 days before the experiment and store it in the refrigerator at +4  C until the measurement. 8. In the pH arrangement, diluted hydrochloric acid and sodium hydroxide could be used if the pH of growth media is far away from the range of 7.5–8. 9. The mixing condition for cells in broth culture is 150 rpm on the shaker. The culture temperature is 37  C. 10. The viability of cells and whether there is any contamination must be checked especially in the beginning of growth phase of cells. 11. Make sure that the calibration of DO probe has been done. Also check that all junction of bioreactor are closed except aeration for sustaining batch operation. 12. The fermentation broth is assumed to be well mixed for determining the kLa from mass balance equations.

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13. The dynamic gas out method provides the determination of the kLa value during a fermentation even at different phases of the process. 14. Make sure the nitrogen tank is sufficiently full. 15. In the dynamic method for evaluation of kLa, oxygen probe must has fast response time for accuracy. Response time constant is corresponding to a time when the probe reaches 63% of its final value when it undergoes a step change according to concentration. Theoretically, the probe response time is much smaller than the mass transfer response time: 1/kLa. But in practice this situation would be change. For approaching the theory, various method in literature has been improved for calculation of the kLa for this reason. It is really important criteria that the response time constant must be smaller than 1/kLa (see ref. 9). 16. The electrode of the DO concentrations in the bioreactor must be applied in sterile conditions as well as fast response. Also, this electrode should be able to measure the concentration instantly. Generally, the electrodes which have these specific properties are preferred. 17. The level of DO concentration does not fall to Ccrit level for keeping the constant the volumetric oxygen uptake rate. So, the oxygen uptake rate of the cells does not change with the oxygen level. 18. In the determination of volumetric mass transfer coefficient of biological broth according to this protocol, the dynamic method is actually corresponding to method of “gas out–gas in” due to non-aeration and reaeration steps, respectively. The gas out step must be carried out in a short time. 19. Multiplicity of C value increases the determination of kLa accuracy. Ensure to record C values in every 5–120 s depending on the microorganism. 20. In the re-aeration step, a specific limitation could occur. In a short period, aeration could not catch steady-state. If the gas residence time is close to time constant of oxygen mass transfer coefficient, this situation can cause inaccuracy. References 1. Po¨pel HJ (1983) Aeration and gas transfer. Delft University of Technology, Delft, The Netherlands, p 168 2. Bailey JE, Ollis DF 2nd (1986) Biochemical engineering fundamentals. McGraw-Hill International Editions, New York 3. Cengel YA, Ghajar AJ 5th (1997) Heat and mass transfer: fundamentals & applications.

McGraw-Hill International Editions, New York 4. Treybal R 3rd (1981) Mass transfer operations. McGraw-Hill International Editions, New York 5. Tu¨rker M 1st (2005) Biyoreaksiyon Mu¨hendislig˘i/Biyolojik Proseslerin Kinetig˘i ve Modellenmesi, Su Vakfı Yayınları

Volumetric Mass Transfer Coefficient Measurement 6. Vilac¸a PR, Badino AC Jr, Facciotti MCR, Schmidell W (2000) Determination of power consumption and volumetric oxygen transfer coefficient in bioreactors. Bioprocess Eng 22:261–265 7. Andre´ G, Moo-Young M, Robinson CW (1981) Improved method for the dynamic measurement of mass transfer coefficient for application to solid-substrate fermentation. Biotechnol Bioeng 213:1611–1622 8. Atkinson B, Mavituna F (1987) Biochemical engineering fundamentals. McGraw-Hill, New York

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9. Doran PM (1995) Bioprocess engineering principles. Academic Press, London 10. Nienow AW (2015) Chapter 5: Mass transfer and mixing across the scale in animal cell culture. In: Nienow AW, Al-Rubeai M (eds) Animal cell culture. Springer International Publishing, Switzerland 11. Orhan R, Dursun G (2013) As¸ag˘ı Dog˘ru Birlikte Akıs¸lı Temas Reakto¨ru¨nde Bacillus Amy€ loliquefaciens ile α-Amilaz Uretiminin € I˙ncelenmesi. Fırat Universitesi Kimya Mu¨hendislig˘i Bo¨lu¨mu¨, Elazıg˘

Methods in Molecular Biology (2022) 2436: 27–38 DOI 10.1007/7651_2021_397 © Springer Science+Business Media, LLC 2021 Published online: 27 April 2021

Fabrication Protocol for Thermoplastic Microfluidic Devices: Nanoliter Volume Bioreactors for Cell Culturing Elif Gencturk, Senol Mutlu, and Kutlu O. Ulgen Abstract Microfluidic devices consist of microchannels etched or embossed into substrates made of polymer, glass or silicon. Intricate connections of the microchannels to reactors with some smart mechanical structures such as traps or curvatures fulfil the desired functionalities such as mixing, separation, flow control or setting the environment for biochemical reactions. Here, we describe the fabrication methods of a thermoplastic microbioreactor step by step. First, material selection is made, then, production methods are determined with the equipment that can be easily procured in a laboratory. COP with its outstanding characteristics among many polymers was chosen. Two types of microbioreactors, with and without electrodes, are designed with AutoCAD and L-edit softwares. Photolithography and electrochemical wet etching are used for master mold preparation. Thermal evaporator is employed for pure chromium and gold deposition on COP substrate and etchants are used to form the interdigitated electrodes. Once the master mold produced, hot embossing is used to obtain the designed shape on drilled and planarized COP. Cover COP, with or without electrodes, is bonded to the hot embossed COP via thermo-compression and thermoplastic microfluidic device is realized. Tubings are connected to the device and a bridge between the macro and micro world is established. Yeast or mammalian cells labeled or tagged with GFP/RFP on specific gene products are loaded into the microfluidic device, and real time data on cell dimensions and fluorescence intensity are collected using inverted fluorescence microscope, and finally image processing is used to analyze the acquired data. Key words COP, Deposition, Etching, Hot embossing, In-house fabricated microfluidic device, Photolithography, Thermo-compression Bonding

1

Introduction Unprecedented success of the microelectronics industry on the integration of micro- and nano-sized devices at very high densities using fabrication methods, such as lithography, thin film deposition and etching, paved the way for the development of microfluidic platforms [1]. The driving force for these technologies has always been to miniaturize desktop sized biochemical analysis systems. The main advantage of miniaturization in these systems is the reduction in the sample and reagent volume. Microfluidic devices also provide faster heat transfer, shorter process times and better automation. Biological and medical applications in cell culture, drug screening, point-of-care (POC) systems adopts microfluidics

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technology due to the integration of several steps into single automated platforms [1–5], making them powerful platforms for single cell studies. Cultivation, separation/isolation, detection and analysis of single cells can be conducted properly within microfluidic devices at high throughput rates, high reproducibility and high automation, with easy and low-cost operation [6–8]. There is an increasing demand for these microfluidic technologies in the global market. According to Market and Market reports, the microfluidic market share was $15.7 billion in 2020, and this share is expected to reach $44 billion in 2025. This market has been segmented into hospitals and diagnostic centers, academic and research institutes, and pharmaceutical and biotechnology companies. The hospitals and diagnostic centers are the areas with the highest market share in this field. The microfluidic technologies continue to grow and develop with university and industry collaborations [9]. The choice of material for microfluidic devices is important in studying cells. In recent years, alternative materials to glass and silicon have been researched among elastomeric and thermoplastic materials. Thermoplastic materials composed of linear and branched molecules are highly preferred due their easy surface modification and durability against temperature as well as pressure changes, and they also do not suffer from any structural breakdown. However, it is not easy to satisfy the material requirements of the specific biological applications. Optical properties, thermostability, chemical stability, and gas permeability are the key arguments of the microfluidic device fabrications [2, 10]. Thermoplastic chips must be biocompatible for cells to survive and transparent to monitor them. For these reasons, polycarbonate, cyclo olefin polymer, poly(methyl methacrylate) and polystyrene stand out compared to other polymers. These polymers are commonly used in industrial manufacturing and possess excellent optical qualifications. They allow rapid prototyping [11, 12]. Fabrication methods of the thermoplastic devices are relatively simple. Fabrication tools are low-cost and easy to use. Wet etching, conventional machining, photolithography, hot embossing, injection molding, laser ablation and 3D printing are some examples of production methods. Selection of fabrication method depends on various factors, such as availability of technology and equipment, cost, speed, and capability [13, 14]. In this chapter, the manufacture of thermoplastic microfluidic devices with and without integrated electrodes is explained for simple systems requiring no active components such as micropumps, micro-valves, and sensors. Photolithography, etching, deposition, hot embossing and thermocompression bonding methods are used for the desired device fabrication and every step is explained in detail.

Fabrication Protocol for Thermoplastic Microfluidic Devices: Nanoliter. . .

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Materials Long-lasting thermoplastic materials are preferred for the fabrication of the microfluidic devices presented here. In our previous studies, polystyrene (PS), poly(methyl methacrylate) (PMMA) and cyclo olefin polymer (COP) were compared to each other and COP was selected as a base material due to its low auto-fluorescence and high rigidity [4]. Detailed information about thermoplastics can be found in the article Genc¸tu¨rk et al. [2]. COP under the brand name Zeonor with product code 10-0672-0349-1.0-05 was purchased from Microfluidic ChipShop Company (Jena, Germany). The chemicals and reagents for medium preparation and cleaning were bought from Sigma-Aldrich (Taufkirchen, Germany). Polyethylene tubing with BB31695-PE/p product code and 5 min epoxy are were purchased from Scientific Commodities Inc. SCI (Lake Havasu City, AZ 86406, USA) and BISON (Rotterdam, The Netherlands), respectively.

3

Methods All solutions that require water are made using distilled water. Preparation and storage of all reagents and chemicals are done at room temperature under clean laminar flow hood. All of the thermoplastic chip fabrication steps are accomplished in clean room. Waste materials were collected in biological waste bags and removed from the laboratory following the necessary procedures. Fabrication steps of the device is summarized in the following figure (Fig. 1).

3.1 Thermoplastic Chip Design and Preparing Its Mold

The main purpose of the design in microfluidic devices for cultivation is to trap and keep cells in predefined areas, where real time monitoring of cells of interest can easily be accomplished. Thus, c-shaped regions that provide cell trapping in the channels of the chip are designed (Fig. 2a), and the high-quality images (brightfield and fluorescence) of the trapped cells are taken throughout the experiment.

3.1.1 Photolithography

1. Draw the microfluidic devices using AutoCAD software (Fig. 2a). 2. Use COMSOL software and determine flow dynamics within the microfluidic device. Determine the flow rate required for cell trapping in the c-shaped regions. 3. Transfer finalized design to acetate paper to be used in the photolithography process with the help of an external high resolution printer with 3000 or higher dot-per-inch (dpi).

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Fig. 1 Fabrication steps of the microfluidic device

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Fig. 2 (a) AutoCAD drawing of designed microfluidic device. (b) Fabricated microfluidic device (reproduced from ref. 15 with permission from Biomedical Microdevices). (c) Fabricated microfluidic device with electrodes. (Reproduced from ref. 16 with permission from Biomicrofluidics)

4. Use 1 mm thick stainless steel substrate for the photolithography process. Here, the aim is to transfer the design on the acetate paper to the stainless steel substrate. 5. Clean stainless steel substrate with baby shampoo, acetone, isopropyl alcohol (IPA) and distilled water, respectively. Dry it with nitrogen gun to leave no residue on steel substrate. 6. Place cleaned steel substrate on the spin coater and coat PR-1828 photoresist on it to obtain an average photoresist layer thickness of 5 μm using 2000 rotation-per-minute (rpm). 7. Implement soft baking process for 2 min at 95  C on hot plate to the photoresist coated steel substrate. 8. Superimpose acetate paper with design on the coated steel substrate and apply UV light (365 nm wavelength, 20 mW/ cm2 power density) for 3 min. 9. Develop the exposed photoresist on the steel substrate by dipping it into the developer solution (MF319 solution). 10. Rinse with distilled water and dry the steel substrate to reveal the design made of photoresist. 11. Check the patterns of the design under the microscope and hard bake photoresist layer on the hot plate for 3 h at 110  C.

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3.1.2 Electrochemical Wet Etching

In order to have a positive stainless steel mold, electrochemical wet etching process is used [17]. Once the master mold is created, it can be used in the fabrication of thermoplastic chips with or without electrodes (Fig. 2b, c). 1. Prepare NaCl: Distilled water solution at a ratio of 1:4 (w/w). 2. Place photoresist coated steel substrate and a blank steel plate in a container facing each other and pour NaCl solution in it. 3. Use blank steel plate as cathode and use photoresist coated steel substrate as anode. 4. Apply DC current of 60 A between the steel plates for 20 s, resulting in an etch depth of around 12 μm. Check the patterns of the design under the microscope. If the desired depth is achieved on the steel substrate, remove photoresist patterns in acetone, IPA and distilled water. Rinse and dry the steel mold.

3.2 Fabrication of Electrodes 3.2.1 Creating an Electrode Design

Electrodes integrated to microfluidic devices can be used to study the effects of electric fields on cells or as actuators to control liquid flow or sort cells on chip [16]. 1. Use L-edit software to design the electrodes. 2. Draw interdigitated electrodes considering the cell size and fixed substrate size of 7.5  2.5 cm2. 3. Transfer finalized drawing to the acetate paper and obtain electrode mask.

3.2.2 Chromium and Gold Deposition on Blank COP Substrate

1. Use thermal evaporator with high vacuum capability in this step. 2. Place pure chromium (99.99%) and gold (99.99%) in the specific chambers within the evaporator. 3. Check the lifetime of the evaporator’s crystal, which measures the sample thickness before deposition process. 4. Place blank COP sample, washed with baby shampoo, acetone, IPA and distilled water, respectively and dried with nitrogen, on the round compartment of evaporator. 5. Close shutter and the chamber of the evaporator and start the vacuum. 6. Leave the system pumping for 24 h and attain 10 Note 1) vacuum level.

6

Torr (see

7. Begin the rotation of the sample and open cooling water of the equipment (see Note 2). 8. First, deposit chromium on the blank COP sample of thickness around 8–10 nm using 50 A electrical current (see Note 3). This is achieved after stable reading is established on the crystal

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monitor, shutter is opened, enough time is waited to reach to the desired thickness and shutter is closed. 9. When the chromium deposition was completed, sample rotation is stopped. The system is left for 2 h for cooling before gold deposition. 10. Gold deposition starts at 100 A electrical current. The thickness is monitored using crystal. Typically 50 nm thickness is achieved using around 100 mg gold pellets. 11. After gold deposition, turn off the electrical current supply and start the ventilation to disrupt the vacuum. 12. Take Cr/Au deposited COP samples from the device and stop cooling water. 3.2.3 Cr/Au Etching to Form Electrodes on Coated COP Sample

1. Clean Cr/Au deposited COP sample with nitrogen and place on the spin coater. 2. Immobilize the sample via vacuum and coat photoresist (PR 1828) on the Cr/Au deposited COP sample at 500 rpm for 2 s and 4000 rpm for 30 s. 3. Soft bake the photoresist coated COP sample for 1 min at 90  C. 4. Place the sample on the table of UV lamp and superimpose electrode mask on it. 5. Expose to UV light at 365 nm wavelength and 20 mW/cm2 power density for 3 min. 6. Submerge the sample into the developer solution (MF319) and shake in the direction of short edge to facilitate the opening of the electrode fingers for 3 min. 7. When UV light exposed part of the photoresist disappear, clean the sample via distilled water and dry with nitrogen. 8. Analyze the sample under microscope to check whether the electrode fingers are opened or not. 9. Dip the COP sample into gold etchant (Potassium iodide and iodine solution, Alfa Aesar, 44584) for 1 min to remove unprotected gold regions. This should reveal clear interdigitated electrodes (fingers). 10. After gold patterns are realized, remove photoresist via acetone, IPA and distilled water, respectively. Finally, etch unprotected chromium regions inside chromium etchant (Ceric ammonium nitrate solution, Nichrome etchant, Alfa Aesar, 44585) for 20 s. Rinse it inside distilled water. 11. Dry the sample by nitrogen and analyze under microscope. 12. Cr/Au electrodes are formed on the COP sample.

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3.3 Drilling and Planarization

1. Superimpose a blank COP sample on the stainless steel mold and mark inlets and outlet holes. 2. Drill inlets and outlet holes from the marked locations on the blank COP substrate. Alternatively, a CNC router can be programmed to drill inlets and outlets on the COP substrate. 3. Put the drilled COP sample into ultrasonic bath for 15 min and dry with nitrogen. 4. Clean the burrs around the holes with a craft knife to have smooth surface. 5. Set the temperatures of the top and bottom plates of the hydraulic press machine (Carver) to 130  C. 6. Clean the drilled COP substrate and 2 mm thick glass slides with acetone, IPA and distilled water and dry with nitrogen, respectively. 7. In order to wipe out the water trace, employ dust-free napkin and use nitrogen to remove the residues on the glass slides and COP sample. 8. After cleaning, stack all the pieces in the order of glass, drilledCOP and glass. Wrap aluminum foil around and place them in between the hot plates of hydraulic press machine (see Notes 4 and 5). 9. Bring hot plates of the hydraulic press machine close to each other (see Note 6). Apply 35 bar pressure for 10 min. 10. Leave the system for cooling to 60  C for 1.5 h under pressure and take pieces out of the machine.

3.4

Hot Embossing

1. Set the temperatures of the top and bottom plates of the hydraulic press machine to 130  C (see Note 7). 2. Clean the stainless steel mold, planarized COP and a glass pieces with baby shampoo, acetone, IPA and distilled water, respectively. 3. Use nitrogen to dry all parts, employ dust-free napkin to remove the traces on the surfaces and use nitrogen again to clean out residues. 4. Superimpose the planarized and cleaned COP on the mold, and sandwich them between glass slides. 5. Place all of the pieces on the hot plate of the hydraulic press machine, and use a piece of aluminum foil to wrap them. 6. Bring the hot plates of the press machine close together, and warm up the pieces (the stainless steel mold, COP and glass) for 10 min.

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35

7. Apply 35 bar pressure for 10 min after the temperature is settled. 8. Leave the system for cooling to 60  C, under pressure for 1.5 h. 3.5 ThermoCompression Bonding

The last step of the microfluidic device fabrication is thermocompression bonding. A cover COP substrate with or without electrodes are bonded to the COP substrate with microchannel grooves and drilled inlet and outlet holes. 1. Set hydraulic press machine to 125  C. 2. Clean two glass slides, hot-embossed-COP substrate and another COP substrate (with or without integrated electrodes) similar to previous steps (see Note 8). 3. Align hot-embossed-COP substrate and the other COP substrates. Stack these substrates between the two glass slides. 4. Wrap around the stack with aluminum foil and place in the hydraulic press machine. 5. Apply warming up process for 10 min without pressure. 6. Apply 25 bar pressure for 45 min to proceed the bonding step. 7. Set the temperature controller of the hydraulic press machine to 10  C for cooling under pressure. 8. Cooling process takes 3 h. Separate glass pieces from the bonded substrates carefully without damaging. 9. Device fabrication is successfully completed (Fig. 2b, c) (see Note 9).

3.6 Establishing the Experiment

1. Prepare all the necessary media (cell medium and nutrient medium) and transfer them into separate syringes (5 mL).

3.6.1 Microfluidic Device Without Electrodes

2. Power on the syringe pumps and place the syringes on them. 3. Power on the inverted fluorescence microscope. 4. Place the fabricated microfluidic device on the stage of inverted fluorescence microscope. 5. Connect the syringe pumps to the microfluidic device via tubings. 6. Prime the microfluidic device with relevant medium (see Note 10). 7. Send cells through the middle inlet of the chip. 8. When chambers are filled with cells, stop the cell feeding and start to send fresh medium through the side inlets for 3 h. 9. Feed medicated medium for another 3 h (see Note 11). 10. Stop medicated medium and send fresh medium for 10 h.

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11. Finish the experiment after 10 h. (Total experiment time is 16 h for yeast cells. For other cell types the duration of experiment is longer than 24 h depending on the doubling time.) 12. Take cell images within the chambers with a determined time interval during the experiment (see Note 12). 3.6.2 Microfluidic Device with Electrodes

1. Repeat the first five steps mentioned in Subheading 3.6.1. 2. Connect function generator (Agilent 33220A) to the microfluidic device and turn its power on. 3. Connect a digital multimeter (Keysight U1252B) for instantaneous measurement of the current within the microfluidic device. 4. Prime the microfluidic device with the relevant medium. 5. Start cell feeding through the middle inlet and trap them in the c-shaped region. 6. Once cells are trapped, stop the cell feeding and start to send fresh medium through the side inlets. 7. Apply electric field with predetermined parameters for 6 h and finish the experiment. 8. Take images of the cells within the chambers of the microfluidic device, placed on the stage of the inverted fluorescence microscope, at preset time intervals throughout the experiment.

4

Notes 1. At this pressure value, water vapors, that might be on the walls of the vacuum chamber or COP sample within the device, were removed. 2. Cooling water was used to prevent damage to the vacuum feedthroughs used to make electrical contact to heating coils. 3. The reason of chromium deposition was to establish adhesion of gold film to the COP substrate. 4. The reason for using aluminum foils was to prevent any molten material from sticking to the plates, which may occur between the hot plates. 5. During processing like hot embossing and bonding, COP substrates are put in between glasses that are used to provide smooth surfaces for pressing. 6. The aim of this step was to warm up the pieces placed in the hydraulic press machine and it takes 10 min. 7. Glass transition temperature of the polymer COP is 136  C. While hot embossing was selected close to the glass transition

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temperature, a lower temperature is preferred for the thermocompression bonding process. 8. Cover COP substrate can be with or without electrodes depending on the application. 9. Subheadings 3.1–3.5 are common in the fabrication of microfluidic devices with and without integrated electrodes. 10. YNB medium is used for yeast cells (Saccharomyces cerevisiae) and RPMI complete medium is used for THP-1 cells, leukemia cell line. 11. Medicated medium can be drugs or inhibitors depending on the study. 12. Time interval for Saccharomyces cerevisiae cells is 10 min while it is 1 h for THP-1 cells.

Acknowledgments This work was supported by Bogazici University Research Fund through project 13641D. References 1. Silva ACQ, Vilela C, Santos HA et al (2020) Recent trends on the development of systems for cancer diagnosis and treatment by microfluidic technology. Appl Mater Today 18:100450 2. Gencturk E, Mutlu S, Ulgen KO (2017) Advances in microfluidic devices made from thermoplastics used in cell biology and analyses. Biomicrofluidics 11(5):051502 3. Luo G, Du L, Wang Y, Wang K (2019) Recent developments in microfluidic device-based preparation, functionalization, and manipulation of nano- and micro-materials. Particuology 45:1–19 4. Puza S, Gencturk E, Odabasi IE et al (2017) Fabrication of cyclo olefin polymer microfluidic devices for trapping and culturing of yeast cells. Biomed Microdevices 19:40 5. Odabasi IE, Gencturk E, Puza S et al (2018) A low cost PS based microfluidic platform to investigate cell cycle towards developing a therapeutic strategy for cancer. Biomed Microdevices 20:57 6. Chen P, Chen D, Li S et al (2019) Microfluidics towards single cell resolution protein analysis. TrAC Trends Anal Chem 117:2–12 7. Dabighi A, Toghraie D (2020) A new microfluidic device for separating circulating tumor cells based on their physical properties by using

electrophoresis and dielectrophoresis forces within an electrical field. Comput Methods Prog Biomed 185:105147 8. Pavesi A, Adriani G, Rasponi M et al (2015) Controlled electromechanical cell stimulation on-a-chip. Sci Rep 5:1–12 9. Markets and Markets (2020) Microfluidics market by product (devices, components (chips, sensors, pump, valves, and needles), application (IVD [POC, clinical, veterinary], research, manufacturing, therapeutics), end user and region – global forecast to 2025. Accessed 25 Feb 2021 10. Voicu D, Lestari G, Wang Y et al (2017) Thermoplastic microfluidic devices for targeted chemical and biological applications. RSC Adv 7:2884–2889 11. Liu K, Fan ZH (2011) Thermoplastic microfluidic devices and their applications in protein and DNA analysis. Analyst 136:1288–1297 12. Matellan C, Del Rı´o Herna´ndez AE (2018) Cost-effective rapid prototyping and assembly of poly(methyl methacrylate) microfluidic devices. Sci Rep 8:1–13 13. Tsao CW (2016) Polymer microfluidics: simple, low-cost fabrication process bridging academic lab research to commercialized production. Micromachines (Basel) 7(12):225

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14. Fiorini GS, Chiu DT (2005) Disposable microfluidic devices: fabrication, function, and application. Biotechniques 38:429–446 15. Gencturk E, Yurdakul E, Celik AY et al (2020) Cell trapping microfluidic chip made of Cyclo olefin polymer enabling two concurrent cell biology experiments with long term durability. Biomed Microdevices 22(1):20 16. Gencturk E, Ulgen KO, Mutlu S (2020) Thermoplastic microfluidic bioreactors with

integrated electrodes to study tumor treating fields on yeast cells. Biomicrofluidics 14 (3):034104 17. Gokdel YD, Mutlu S, Yalcinkaya AD (2010) Self-terminating electrochemical etching of stainless steel for the fabrication of micromirrors. J Micromech Microeng 20 (20):95009–95006

Methods in Molecular Biology (2022) 2436: 39–53 DOI 10.1007/7651_2021_396 © Springer Science+Business Media, LLC 2021 Published online: 07 May 2021

Expansion of Human Pluripotent Stem Cells in Stirred Tank Bioreactors Marites T. Woon, Puspa R. Pandey, and Inbar Friedrich Ben-Nun Abstract Bioreactor technolology enables the expansion of mammalian cells, which can be translated to processes compatible with Current Good Manufacturing Practice (cGMP) regulations. Cells are introduced to the bioreactor vessel, wherein key parameters such as temperature, pH, and oxygen levels are tightly controlled to facilitate growth over time. Here, we describe the microcarrier-based expansion of human pluripotent stem cells in a 3 L stirred tank bioreactor. Key words Bioreactor, Cell therapy, Human pluripotent stem cell expansion, Microcarrier, Scalable

1

Introduction Stem cell technology has rapidly advanced since its development and continues to revolutionize personalized medicine, drug development and disease modeling [1]. Human pluripotent stem cells (hPSCs) can differentiate, giving rise to clinically relevant cell types such as cardiomyocytes [2] and neurons [3]. Holding remarkable promise toward the development of curative therapies, considerable attention has been given in the large-scale expansion of these invaluable cells [4–9]. Expansion of hPSCs required for cell-based therapies using bioreactors addresses key limitations of conventional 2D culture methods. The culture of these cells onto glass or plastic surfaces is a process ubiquitious across academic and instustry-based laboratories. However, continued understanding of cell biology and method improvements support the application of 3D culture methods instead [10]. 3D culture methods more accurately represent the cellular microenvironment found in vivo, and cells cultured in these systems respond accordingly [11]. Drug development studies have demonstrated distinct differences in the responses of 2D vs 3D cultured cells [12, 13], underscoring the need to recapitulate complex cellular microenvironments to derive accurate responses toward therapeutic candidates. Another critical limitation of 2D culture systems is scalability. For example, approximately 1
108

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– 1
109 hPSC-derived cardiomyoyctes will be required to replace the considerable number of cells lost in a myocardial infarction [14]. To this end, bioreactors provide a robust, efficient and scalable method to generate an unlimited supply of cells critical in many clinical indications. Here, we describe the microcarrier-based expansion of hPSCs in a 3 L bioreactor, resulting in high-fold expansion of high quality cells within 2 weeks of culture [9]. The cell inoculum—derived from cells cultured in 2D—are introduced into the bioreactor with extracellular matrix-coated plastic microcarriers. Compared to suspension cultures which use an aggregatebased method for expansion [7, 15], microcarriers provide a large surface area on which cells attach and grow [9, 16].

2

Materials Perform the procedures using aseptic technique inside a Biological Safety Cabinet unless otherwise noted. Limit the exposure of the L7™ hPSC culture system to light. Store reagents according to manufacturer’s instructions.

2.1 L7™ TFO2 hPSC Complete Medium

1. Thaw a 10 mL bottle of L7™ hPSC medium supplement in 37  C. Add the entire contents to 1 L of L7™ TFO2 hPSC basal medium (see Note 1). Invert the bottle twice to mix and store in 4  C until use. 2. 7 L bags of L7™ TFO2 hPSC basal medium (see Note 2).

2.2 Passaging Solutions

1. L7™ hPSC passaging solution (see Note 3).

2.3 Buffers and Reagents

1. Cell culture grade water.

2. F3 hPSC passaging solution (see Note 4).

2. Dulbecco’s Phosphate-Buffered Saline with calcium and magnesium (DPBS+/+); pH 7–7.6. Store at 15–30  C. 3. Dulbecco’s Phosphate-Buffered Saline without calcium and magnesium (DPBS/); pH 7–7.6. Store at 15–30  C. 4. 250 μg/mL L7™ hPSC matrix. Add 4 mL of cell culture grade water to 1 mg of lyophilized matrix (see Note 5). 5. 10 mM Y-27632 Dihydrochloride (Rho-associated protein kinase (ROCK) inhibitor). Add 624.4 μL DMSO to 2 mg of ROCK inhibitor. Mix by pipetting (see Note 6).

2.4

Tubing

1. Assemble the dip tube/perfusion line (see Fig. 1), media feed line (see Fig. 2) and harvest line extension assembly (see Fig. 3). Autoclave using the dry cycle (see Note 7).

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Fig. 1 Assemble the dip tube/perfusion line. This allows for the continuous perfusion of cell culture medium, while retaining the cells in the bioreactor vessel during the expansion process

Fig. 2 Assemble the media feed line. This allows for the introduction of cell culture medium during the expansion process 2.5

Bioreactor

1. T-75 flask. 2. 1-Layer cell stack. 3. 125–212 μM polystyrene microcarriers.

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Fig. 3 Assemble the harvest line extension. This allows for the harvest or collection of expanded cells from the bioreactor vessel

4. 3 L stirred tank bioreactor vessel. 5. Bio-welder. 6. pH probe (see Note 8). 7. DO probe. 8. Temperature probe. 9. Motor. 10. Heating jacket. 11. Hemostat. 12. Parafilm. 13. Conical tubes. 14. Syringes. 15. Extension sets. 16. Junction box. 17. Bioprocessing system. 18. Analytical scale. 19. Waste bag. 20. Mesh filter. 21. Intravenous pole. 22. Peristaltic pump. 23. Cell counter (hemocytometer or automated).

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Methods Perform the procedures using aseptic technique inside a Biological Safety Cabinet unless otherwise noted and limit the exposure of the L7™ hPSC culture system to light. To maintain a sterile environment within the bioreactor vessel and cell culture media bags, unclamp the tubings only when introducing reagents, media or cell inoculum or drawing samples. Immediately seal the line and re-clamp the tubing when finished.

3.1 Extracellular Matrix Coating of T-75 and 1-Layer Cell Stack

1. Re-suspend 300 μL of the L7™ hPSC matrix stock solution in 12 mL of DPBS+/+ and add to the T-75 flask. To coat a 1-layer cell stack, resuspend 2.55 mL of the L7™ hPSC matrix stock solution in 125 mL of DPBS+/+. 2. Allow the coating solution to incubate in a 37  C incubator for at least 1 h (see Note 9).

3.2 hPSC Culture on T-75 Flask

1. Thaw cryopreserved hPSC cell line of choice in a 37  C water bath. 2. Transfer the cell solution into a 50 mL conical tube and add 10 mL of L7™ TFO2 complete medium dropwise. 3. Centrifuge at 200
g for 5 min. 4. Aspirate the supernatant and re-suspend the cell pellet in 15 mL of L7™ TFO2 complete medium supplemented with 10 μM ROCK inhibitor. 5. Before adding the hPSCs to the T-75 flask, aspirate the extracellular matrix coating solution. 6. Replace the medium by aspirating the old medium and adding new 15 mL of L7™ TFO2 complete medium (see Note 10). 7. Culture the cells in an incubator at 37  C and 5% CO2 until the cells reach 70–80% confluence (see Note 11).

3.3 hPSC Culture on 2D Seed Train

1. Aspirate the old medium from the T-75 flask and wash cells with 15 mL of DPBS/. 2. Incubate with 15 mL of pre-warmed L7™ hPSC passaging solution in 37  C for 10 min (see Note 12). 3. Aspirate the L7™ hPSC passaging solution and add 15 mL of L7™ TFO2 complete medium to harvest the cells into a 50 mL conical tube. 4. Centrifuge at 200
g for 5 min. 5. Remove the supernatant and re-suspend the cells in 10 mL of L7™ TFO2 complete medium. 6. Count the number of cells (see Note 13) to determine the volume required to seed 0.02–0.03
106 cells/cm2.

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7. Aspirate the extracellular matrix solution from the 1-layer cell stack before adding the hPSCs. 8. Culture the cells in an incubator at 37  C and 5% CO2 until the cells reach 70–80% confluence (see Note 11) by aspirating the old culture medium and replenishing with 150 mL of new culture medium (see Note 10). 9. To harvest the cell inoculum, aspirate the old medium from the 1-layer cell stack. 10. Wash the cells with 75 mL DPBS/ (see Note 14). 11. Aspirate and add 75 mL of pre-warmed L7™ hPSC passaging solution. Incubate at 37  C for approximately 15 min (see Note 12). 12. Tap the 1-layer cell stack to detach the cells. 13. Transfer the cells into a 250 mL conical tube and add 75 mL L7™ TFO2 complete medium into the tube. 14. Centrifuge the cell suspension at 200
g for 5 min, aspirate the supernatant and re-suspend the cell pellet with 30 mL of L7™ TFO2 complete medium. 15. Count the number of cells (see Note 13) to determine the volume required to inoculate a 3 L bioreactor with 120
106 cells harvested with L7™ hPSC passaging solution (see Note 15). 3.4 Microcarrier Coating

1. In a 1 L bottle, add 20 g of microcarriers, 300 mL DPBS+/+ and 18 mL of L7™ hPSC matrix. 2. Incubate at 37  C for 2–4 h with intermittent shaking to mix. 3. Allow the coated microcarriers to settle at the bottom of the bottle before aspirating the DPBS+/+. 4. Add 300 mL of L7™ TFO2 hPSC basal medium without the L7™ hPSC medium supplement (incomplete) and wrap the bottle with aluminum foil. Incubate overnight while shaking at room temperature. 5. Add 700 mL of incomplete L7™ TFO2 hPSC medium and store in 4  C until use (see Note 16). 6. When ready to introduce to the bioreactor vessel, replace the bottle closure. Use a bottle closure pre-mounted with tubing. 7. Weld the pre-mounted tubing of the bottle closure to a 2 L bag of L7™ TFO2 hPSC basal medium and introduce the coated microcarriers to the medium (see Note 17).

3.5

Bioreactor Setup

1. Carefully unwrap the bioreactor in the Biological Safety Cabinet and inspect the vessel to ensure it is not damaged prior to use.

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2. Insert the sterile pH probe into the appropriate port. 3. Insert the sterile perfusion dip tube into the appropriate port (see Note 18). 4. Screw the pH probe and perfusion dip tube in the vessel tightly to ensure they are held in place. 5. Wrap parafilm around the ports to close any gaps and maintain a sterile environment. 6. To the Harvest port on the vessel, remove the male MPC connection. Attach the harvest line extension assembly (see Fig. 3) by removing the MPC coupling body part and inserting the male type connection on the assembly line to the female connection on the vessel. 7. Attach extension sets onto two luer lock female type connections on the vessel ports labeled LA2 (see Note 19) and Sample (see Note 20). 8. Close the Roberts clamp on the vessel and close the hemostat at the far end of the tubing. 9. Carefully remove the vessel from the Biological Safety Cabinet (see Note 21). 10. Weld the Media-In line to the LA1 line. 11. Tare the scale, which is connected to the DeltaV control platform. 12. Place the vessel on the scale. 13. Insert the DO probe to the appropriate port (see Note 22) and connect it to the junction box (see Note 23). 14. Uncap the pH probe and connect it to the Finesse controller. 15. Insert the temperature probe in the appropriate port and connect to the Finesse controller. 16. Attach the motor to the center of the vessel headplate (see Note 24). 17. Affix the heating jacket around the vessel (see Note 25). 18. Connect the gas line from the Mass Flow Controller labeled headspace to the gas inlet line. 19. Record the weight of the vessel with added probes and accessories. Tare the scale (see Note 26). 20. On the DeltaV control platform, confirm that the vessel weight is 0 g. Enter a specific Batch ID and click ‘Batch Start.’ 21. Weld the 3 L L7™ TFO2 hPSC basal medium with coated microcarriers to the LA2 port tubing and allow contents to enter the vessel by gravity. 22. Verify the appropriate process parameters on the DeltaV control platform: 37  C temperature, 100 RPM agitation and 2 L/ min air flow rate.

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23. Allow the vessel to become saturated with air (see Note 27). 24. Weld a syringe-extension set to the LA(S) tubing and draw 5 mL of sample from the vessel. 25. Measure the pH (see Note 28). 26. Adjust the pH value in the DeltaV control platform if the online and offline values differ by >0.05. 27. Verify the critical control parameters and leave the system overnight (see Note 29). Introduction of cell inoculum is performed the following day. 28. Prior to introducing the cell inoculum, transfer 30 mL of L7™ hPSC medium supplement into a 50 mL syringe and attach an extension set. 29. Weld the syringe-extension set to the LA(S) port and add the L7™ hPSC medium supplement into the vessel. 30. After 1 h, weld a syringe-extension set to the LA(S) tubing and draw 5 mL of sample from the vessel. 31. Measure the pH (see Note 28). 32. Adjust the pH value in the DeltaV control platform if the online and offline values differ by >0.05. 3.6 Cell Inoculum Introduction

1. Transfer the cell inoculum harvested from the 2D seed train into an appropriately sized syringe (see Note 30). 2. Connect the syringe to a luer lock extension tubing. 3. Weld the syringe-extension set to tubing corresponding to the LA(S) port on the vessel. Gently introduce the cell inoculum by pushing the syringe piston (see Note 31). 4. To the same LA(S) port line, add 3 mL of 10 mM ROCK inhibitor. 5. Mix the cell inoculum for 30 min to 1 h and draw a 5 mL sample from the vessel to measure the pH offline (see Note 28). 6. Adjust the pH value in the DeltaV control platform if the online and offline values differ by >0.05. 7. Set up the appropriate program in the DeltaV control platform and verify the critical control parameters (see Note 32). This begins the cell growth and expansion process in a 3 L bioreactor vessel.

3.7 Cell Culture Medium Perfusion Setup

1. Weld the C-flex tubing of the media feed line (see Fig. 2) to the LA (1) tubing. 2. Weld the C-flex tubing of the 7 L bag of complete L7™ hPSC (see Note 33) medium to the far end of the media feed line.

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3. Place the Pharmed tubing portion of the media feed line (see Fig. 2) into the pump controller (pump #3) located on the G3 model tower. 4. Connect a 20 L empty bag with a 1/800
1/400 C-flex tubing by welding it to the C-flex tubing of the perfusion out line. This is the waste bag. 5. Place the waste bag on a scale located on the floor and tare the scale. 6. Place the Pharmed tubing portion of the perfusion out line (see Fig. 1) into the pump controller (pump #4) located on the G3 model tower. 7. Unclamp the tubings and prime the waste line by pressing the button next to pump #4. Media will travel from the vessel to the waste bag. To prime the feed line, press the button next to pump #3 until media can be observed entering the vessel (see Note 34). 8. Initiate perfusion by setting the value of pump #3 to cascade on ‘vessel weight output’ and the value of pump #4 to 2.08 g/min on the DeltaV control platform. 3.8 Cell Count during Expansion

1. Aliquot 15 mL of F3 hPSC passaging solution into two 50 mL conical tubes and allow to warm up in the 37  C incubator for 10 min. 2. Weld a syringe-extension set to the bioreactor and draw 15 mL of cell solution. Take two 15 mL samples. 3. Deposit the cell solution into 50 mL conical tubes and allow the microcarriers to settle (see Note 35). 4. Aspirate the media carefully and resuspend the microcarriers in 7.5 mL F3 hPSC passaging solution. 5. Invert the conical tube twice to mix and incubate in the 37  C incubator for 15–20 min (see Note 36). 6. Gently pipette the cell solution. 7. Prepare a single cell suspension by straining the microcarriercell solution through a 70 μM cell strainer into another 50 mL conical tube. 8. Add 7.5 mL of pre-warmed incomplete or complete L7™ TFO2 hPSC medium. 9. Centrifuge the cell only solution at 200
g for 5 min. 10. Re-suspend the cell pellet in 1 mL or 15 mL (see Note 37) L7™ TFO2 hPSC medium. 11. Perform cell count (see Note 13) throughout the expansion process (see Fig. 4).

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A

B Cell count from samples

Cell count from full harvest

Live cells/mL

2.50E+06 2.00E+06 1.50E+06 1.00E+06 5.00E+05 0.00E+00

0

3

6 Days in culture

9

12

Fig. 4 Expansion of hPSC in a 3 L bioreactor. (a) Representative cell density plot over time. (b) Representative image of cell-microcarrier clusters on day 11 of the expansion process as indicated by the yellow arrows; 10
magnification 3.9

Cell Harvest

1. Pre-warm 1.5 L bag of F3 hPSC passaging solution to 37  C. 2. Pre-warm 1.5 L bag of L7™ TFO2 hPSC basal medium to 37  C. 3. Record the weight of the bioreactor vessel. 4. Using two different syringes, transfer 30 mL of L7™ hPSC medium supplement and 3 mL of 10 mM ROCK inhibitor solution into the 5 L media bag containing 1.5 L of L7™ TFO2 hPSC basal medium. 5. Weld the C-flex tubing on the 1.5 L bag of L7™ TFO2 hPSC basal medium to the C-flex tubing of the mesh filter bag (see Note 38). 6. Weld the other end of the mesh filter bag with the C-flex tubing of the harvest line, which is attached to the Harvest port of the bioreactor vessel. 7. Attach an empty and sterile 5 L plastic bag to the Perfusion line via the C-flex tubing ends. This bag will contain the cell culture supernatant derived from the vessel. 8. When the appropriate tubings are welded to the vessel, remove the cell culture medium from the bioreactor vessel through the Perfusion line into the waste bag (see Note 39) while agitating. 9. If cell-microcarrier aggregates on the mesh filter are observed, then gently tap the vessel or rotate the perfusion dipstick. 10. After 1.5 L of volume has been removed from the vessel, remove the heating jacket and lower the agitation speed by 10 rpm (see Note 40). 11. Halt the agitation when ~10% of the original vessel volume remains.

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12. Connect the C-flex tubing of a 1.5 L bag of pre-warmed F3 hPSC passaging solution to the Sampling port tubing. Allow the F3 hPSC passaging solution to enter the bioreactor vessel by gravity (see Note 41). 13. When the vessel is filled with 1.5 L of F3 hPSC passaging solution, put on the heating jacket and center the bioreactor vessel on the analytical scale. 14. During the incubation step, turn on the heating jacket at 37  C and begin agitation at 90 rpm for 30 min. 15. Weld a syringe-extension set to the bioreactor and draw 5 mL of sample. Transfer the solution onto a 10 cm cell culture dish and confirm the detachment of cells from the microcarriers using a microscope. 16. Begin the filtration process when more than 90% of cells are detached from the microcarriers as single cells (see Note 42). 17. While maintaining 90 rpm agitation speed, begin the filtration process by using the pump (200 mL/min) via the harvest line into the mesh filter. 18. When the top of the impeller is visible, decrease the agitation speed to 30 rpm. Tilt the vessel to retrieve as much of the cells from the vessel as possible (see Note 43). 19. Gently shake the bag to mix the cell suspension. 20. Weld a syringe-extension set to the bioreactor and draw three 5 mL samples. 21. Count the number of cells and determine viability (see Note 13). 22. Downstream processes of interest such as directed differentiation and cryopreservation [9] can now be performed on the expanded cells (see Note 44).

4

Notes 1. Incomplete medium refers to the L7™ TFO2 hPSC basal medium without the L7™ hPSC medium supplement. 2. Other commercially available cell culture medium such as mTeSR1 [6] have been used in the expansion of stem cells in bioreactors. During the expansion process in a 3 L bioreactor vessel, the Lonza L7™ TFO2 hPSC basal medium packaged in 7 L media bags were used. 3. The L7™ hPSC passaging solution is a non-enzymatic solution that results in cell clumps. 4. The F3 hPSC passaging solution is a non-enzymatic solution that results in single cells and may require the addition of

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10 μM ROCK inhibitor to facilitate attachment to the cell culture vessel. 5. The L7™ hPSC matrix stock solution can be aliquoted in 1.5 mL Eppendorf tubes and stored at 20  C until use. Avoid multiple freeze-thaw cycles. 6. Aliquot the stock solution of ROCK inhibitor and store at 20  C until use. Avoid multiple freeze–thaw cycles. 7. Following the autoclave cycle, allow the sterilized materials to cool before use. Assemble the required tubings with the bioreactor vessel in a Biological Safety Cabinet. 8. Autoclave the pH probe using the liquid cycle. Allow the probe to cool after autoclaving. 9. Cell culture vessels may be coated with the extracellular matrix coating solution for at least an hour and up to 48 h in 37  C. Store in 4  C if not used immediately. Use the coated vessels within 2 weeks. 10. The L7™ hPSC culture system allows for an every-other-day medium change. 11. Ensure to passage or harvest cells at 70–80% confluence. Higher cell confluence can lead to lower cell viability and impact attachment to the coated microcarriers. 12. Every 5 min, examine the colonies under the microscope. Over time, the colonies will form holes indicating that the cells are ready to be harvested. 13. Cell counts can be performed using a hemocytometer or an automated cell counter. 14. Tilt the 1-layer cell stack when adding the DPBS/ to wash, but not to dislodge the attached cells. 15. Lower cell densities may be used. The cell density of cell inoculum and rate of expansion are also dependent on the cell line used in the process. 16. The coated microcarriers may be stored in 4  C for up to 7 days. Longer storage times have not been tested. 17. When introducing the coated microcarriers to the bag of L7™ TFO2 hPSC basal medium, periodically shake the bottle to avoid settlement of microcarriers at the bottom of the bottle. 18. Insert the mesh end of the perfusion dip tube carefully. Do not bend the mesh against the bottom of the vessel lest it breaks. 19. The LA2 port corresponds to the port from which aliquots of cell solutions can be drawn. 20. The Sample port is used as an alternative to the LA2 port. 21. Prior to removing the vessel from the Biological Safety Cabinet, ensure that the ports are closed and the tubings are

Expansion of Human Pluripotent Stem Cells in Stirred Tank Bioreactors

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securely attached. A sterile environment inside the vessel must be maintained. 22. Tightly press the DO probe against the membrane at the bottom of the vessel. 23. Connecting the DO probe to the junction box (J-box) allows the careful monitoring of DO levels during the expansion process. 24. The motor may impinge on the DO probe. Therefore, minor adjustments may be required. 25. Ensure that the electric wire connected to the heating jacket is connected to the Finesse controller. The heating jacket must not touch the scale to accurately report the vessel weight. 26. Following this point, the weight recorded on the scale corresponds to the liquid contents inside the vessel. 27. Saturation is achieved after 4–7 h when the DeltaV control platform shows that the DO levels have plateaued. 28. Offline pH measurement can be done using the BioProfile FLEX Analyzer (Nova Biomedical) according to manufacturer’s instructions. 29. Input the set points for temperature, pH, DO, gas, and agitation speed in the DeltaV control platform. 30. In lieu of obtaining cell inoculum from a 2D seed train, cryopreserved cells or a 3D seed train can also be used [9]. 31. Pushing air may be required to ensure the entire cell suspension is introduced into the vessel. 32. The values entered in the DeltaV control platform are dependent on the experiment’s parameters. 33. The 7 L bag of complete L7™ TFO2 hPSC medium was prepared by adding 70 mL of L7™ hPSC medium supplement to the L7™ TFO2 hPSC basal medium. This will be used during the expansion process. Elevate the bag on an intravenous pole, which creates a pressure gradient allowing the medium to enter the vessel. Cover the bag to protect the medium from light. 34. Carefully inspect for any leaks in the tubings. 35. 1 mL of the supernatant can be used to check the number of cells not attached on the microcarriers. This value should be negligible as it is expected that the cells adhered and grew on the coated microcarriers. 36. Periodically invert the tube to mix. This ensures the homogeneous dissociation of cells from the microcarriers. Larger aggregates tend to take longer than 20 min to dissociate from

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the microcarriers. Therefore, periodically check under the microscope to determine whether the cells have detached. 37. For cell counts performed between the first to tenth day of culture in the bioreactor, re-suspend the cell pellet in a smaller volume of media. However, for cells cultured more than 10 days in the bioreactor, re-suspend the cell pellet in a larger volume of media. 38. The 65 μm pore size of the mesh filter allows the separation of microcarriers from the cell solution during harvest. 39. Use a peristaltic pump (100–200 mL/min) or gravity to introduce the F3 hPSC passaging solution to the bioreactor vessel and to remove the cell culture medium from the vessel to the waste bag. 40. For every 1/3 decrease in volume, lower the agitation speed by 10 rpm. When the impeller is visible, discontinue agitation. 41. The bag of F3 hPSC passaging solution can also be connected to the harvest line. Use a peristaltic pump (100–200 mL/min) or gravity to fill the vessel with 1.5 L of F3 hPSC passaging solution. 42. The cell-microcarriers can be incubated with the F3 hPSC passaging solution for another 5–10 min to ensure release of cells from the microcarriers. Longer incubation of the F3 hPSC passaging solution will not adversely impact the health of the cells. Take another 5 mL sample to confirm. 43. At this juncture, single cells will pass through the mesh filter to the media bag and the microcarriers will be deposited to the other side instead. Therefore, the media bag will contain 3 L of single cell suspension composed of single cells in 1.5 L of F3 hPSC passaging solution and 1.5 L of L7™ TFO2 hPSC complete medium. 44. The expanded cells can be characterized using qualitative and quantitative methods to ascertain pluripotency such as immunofluorescence staining and flow cytometry, respectively. References 1. Shi Y, Inoue H, Wu JC, Yamanaka S (2017) Induced pluripotent stem cell technology: a decade of progress. Nat Rev Drug Discov 16:115–130 2. Protze SI, Lee JH, Keller GM (2019) Human pluripotent stem cell-derived cardiovascular cells: from developmental biology to therapeutic applications. Cell Stem Cell 25:311–327 3. Tao Y, Zhang SC (2016) Neural subtype specification from human pluripotent stem cells. Cell Stem Cell 19:573–586

4. Shafa M, Yang F, Fellner T, Rao MS, Baghbaderani BA (2018) Human-induced pluripotent stem cells manufactured using a current good manufacturing practice-compliant process differentiate into clinically relevant cells from three germ layers. Front Med 5. https://doi. org/10.3389/fmed.2018.00069 5. Li X, Ma R, Gu Q, Liang L, Wang L, Zhang Y, Wang X, Liu X, Li Z, Fang J, Wu J, Wang Y, Li W, Hu B, Wang L, Zhou Q, Hao J (2018) A fully defined static suspension culture system

Expansion of Human Pluripotent Stem Cells in Stirred Tank Bioreactors for large-scale human embryonic stem cell production. Cell Death Dis 9. https://doi.org/ 10.1038/s41419-018-0863-8 6. Nogueira DES, Rodrigues CAV, Carvalho MS, Miranda CC, Hashimura Y, Jung S, Lee B, Cabral JMS (2019) Strategies for the expansion of human induced pluripotent stem cells as aggregates in single-use Vertical-Wheel™ bioreactors. J Biol Eng 13. https://doi.org/10. 1186/s13036-019-0204-1 7. Shafa M, Panchalingam KM, Walsh T, Richardson T, Baghbaderani BA (2019) Computational fluid dynamics modeling, a novel, and effective approach for developing scalable cell therapy manufacturing processes. Biotechnol Bioeng 116:3228–3241 8. Pan T, Chen Y, Zhuang Y, Yang F, Xu Y, Tao J, You K, Wang N, Wu Y, Lin X, Wu F, Liu Y, Li Y, Wang G, Li Y, Wang G, Li Y-X (2019) Synergistic modulation of signaling pathways to expand and maintain the bipotency of human hepatoblasts. Stem Cell Res Ther 10. https://doi.org/10.1186/s13287-019-1463y 9. Pandey PR, Tomney A, Woon MT, Uth N, Shafighi F, Ngabo I, Vallabhaneni H, Levinson Y, Abraham E, Ben-Nun IF (2020) End-to-end platform for human pluripotent stem cell manufacturing. Int J Mol Sci 21. https://doi.org/10.3390/ijms21010089 10. Jensen C, Teng Y (2020) Is it time to start transitioning from 2D to 3D cell culture? Front Mol Biosci 7. https://doi.org/10. 3389/fmolb.2020.00033

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11. Edmondson R, Broglie JJ, Adcock AF, Yang L (2014) Three-dimensional cell culture systems and their applications in drug discovery and cell-based biosensors. Assay Drug Dev Technol 12:207–218 12. Langhans SA (2018) Three-dimensional in vitro cell culture models in drug discovery and drug repositioning. Front Pharmacol 9. https://doi.org/10.3389/fphar.2018. 00006 13. Lv D, Hu Z, Lu L, Lu H, Xu X (2017) Threedimensional cell culture: a powerful tool in tumor research and drug discovery. Oncol Lett 14:6999–7010 14. Mummery CL, Zhang J, Ng ES, Elliott DA, Elefanty AG, Kamp TJ (2012) Differentiation of human embryonic stem cells and induced pluripotent stem cells to cardiomyocytes: a methods overview. Circ Res 111:344–358 15. Schwedhelm I, Zdzieblo D, Appelt-Menzel A, Berger C, Schmitz T, Schuldt B, Franke A, Mu¨ller FJ, Pless O, Schwarz T, Wiedemann P, Walles H, Hansmann J (2019) Automated realtime monitoring of human pluripotent stem cell aggregation in stirred tank reactors. Sci Rep 9. https://doi.org/10.1038/s41598019-48814-w 16. Badenes SM, Fernandes TG, Cordeiro CSM, Boucher S, Kuninger D, Vemuri MC, Diogo MM, Cabral JMS (2016) Defined essential 80 medium and vitronectin efficiently support scalable xeno-free expansion of human induced pluripotent stem cells in stirred microcarrier culture systems. PLoS One 11. https://doi. org/10.1371/journal.pone.0151264

Methods in Molecular Biology (2022) 2436: 55–66 DOI 10.1007/7651_2021_412 © Springer Science+Business Media, LLC 2021 Published online: 10 August 2021

High-Efficiency Differentiation of Human Pluripotent Stem Cells to Hematopoietic Stem/Progenitor Cells in Random Positioning Machine Bioreactors Xiaohua Lei, Chiyuan Ma, Yujing Cao, Yue Xiong, Jian V. Zhang, and Enkui Duan Abstract Human pluripotent stem cells (PSCs) are known to differentiate into almost all the blood lineage cells in vitro and hold a great promise for studying human early hematopoietic development and have a huge potential in the treatment of hematological disorders. Although several methods of hematopoietic stem/ progenitor cell (HSPC) differentiation have been developed, the HSPC yields achieved using these strategies are not yet available for clinical application. Recently, bioreactor-based devices and biochemical factors synergistically have been used to induce hematopoietic differentiation and showed a potential role in hematopoiesis. This chapter describes a protocol for using a random positioning machine bioreactor to culture human PSCs and the large-scale production of HPCs. Techniques for characterizing the differentiated cells and assessing the efficiency of hematopoietic differentiation in the bioreactor with immunostaining and flow cytometry are also presented. Key words Differentiation, Hematopoietic stem/progenitor cells, Human pluripotent stem cell, Random positioning machine

1

Introduction Human pluripotent stem cells (hPSCs) include human embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs), which have the capacity of self-renewal and can differentiate into almost all blood lineage cells, thus offering an invaluable model for dissecting early human hematopoietic development and the in vitro production of hematopoietic stem/progenitor cells (HSPCs) and functional blood cells for therapies of various hematologic disorders [1–3]. Some in vitro directed hematopoietic differentiation protocols from hPSCs have been developed, yet to date it remains a great challenge to generate HSPCs with robust multilineage engraftment potential and infusion dosage levels of functional blood cells from hPSCs [4]. Kaufman et al. reported a strategy for the first time to generate hematopoietic progenitors derived from human embryonic stem cells by coculturing with the murine S17

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stromal cells, which were able to myeloid colony-forming [1]. Moreover, Coculture of hPSCs with OP9 bone marrow stromal cells or with mAGM-S3 can also generation of multilineage HSPCs [3, 5, 6]. The differentiation efficiency of embryoid bodies (EBs)-based model is low due to the heterogeneity of cell types and the lack of access to nutrients of internal cells [7]. Additionally, coculture with mouse feeder cells leads to human cells coming into contact with cells of foreign species, which may be detrimental to subsequent therapeutic applications. Hematopoietic differentiation from hPSCs in chemically defined systems were also developed [6, 8], but often involves culturing in specialized medium and many steps of the differentiation, which make the cost of HPSCs very expensive and it is difficult to produce the large-scale of HPSCs. To promote hematopoietic differentiation of hPSCs, the proper use of growth factors and cytokines to benefit the production of HSPCs were investigated. Kennedy et al. reported that inhibition of Nodal/Activin pathway during hematopoiesis from human ESCs in a chemically defined medium containing the SB-431542, can induce the development of definitive HSPCs and blocked the primitive hematopoiesis process [9]. The canonical Wnt pathway is also known to play a vital role in the induction of definitive HPSCs derived from human ESCs [10]. Treatment of human ESCs with GSK-3 inhibitor CHIR99021 by activation of canonical Wnt signaling can promote definitive hematopoiesis and inhibited the number of primitive HSPCs [11]. In contrast, treating human ESCs with the Wnt-antagonist IWP2 augmented the number of primitive HSPCs. In keeping with this study, Wang et al. reported that R-spondin2 plays a key role in early hematopoietic differentiation of hPSCs that increased the generation of APLNR+ mesoderm cells by activating TGF beta signaling [12]. In another approach, ectopic expression of transcription factors promotes the hematopoietic commitment of hPSCs by increasing the expression of mesoderm, hemogenic endothelial and the genes associated with hematopoietic development. Ran et al. demonstrated that expression of endogenous RUNX1a promotes hematopoietic lineage commitment from hPSCs and enhanced definitive hematopoiesis [13]. It has also been shown that ectopic expression of HOXA9 increased hematopoietic commitment from human ESCs; however, HOXA9 was not sufficient to confer in vivo long-term engraftment potential [14]. Interesting, a recent study reported that suppression of MSX2 enhances hematopoietic differentiation of hPSCs via inhibition of TGF beta signaling [15]. Recent work has indicated the importance of the local physical environment (such as blood flow, wall shear stress) in regulating HE specification and HSPC production [16–18]. To mimic the physical microenvironment, some bioengineering techniques that promote hematopoiesis from hPSCs have been applied [19, 20]. Additionally, the combination of bioreactors and

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chemical factors might further promote hematopoiesis [21]. In the following section, we describe a protocol for using the random positioning machine (RPM) bioreactor to culture hPSCs and the large-scale production of HSPCs in a chemically defined system. We identify the characteristics of the cells and analyze the efficiency of hematopoietic differentiation in the RPM bioreactor by immunohistochemistry and flow cytometry.

2 2.1

Materials Cell Lines

2.2 Cell Culture Medium and Reagents

Human ESC line H1 is obtained from Wicell Research Institute, Inc., Madison, WI. 1. Medium for undifferentiated human ESCs expansion: TeSR™-E8™ medium (Cat# 05990, STEMCELL Technologies), basal medium store at 4  C, 25 supplement store at 20  C. 2. Matrigel for undifferentiated human ESCs expansion: Matrigel Matrix HESC-qualified (BioCoat, 354277); Matrigel for human ESCs differentiation: Corning Matrigel Growth Factor Reduced Basement Membrane Matrix (Corning, Cat#354230). The final dilution of Matrigel should be 1:50. 3. IF9S basal medium for human ESC differentiation: The formulation of IF9S is list in Table 1 according to previously described [22]. Store at 4  C. 4. Mesoderm induction medium: IF9S basal medium with 50 ng/mL BMP4 (PeproTech), 15 ng/mL ACTIVIN A (PeproTech) and 1.5 μM CHIR99021 (STEMCELL™ Technologies) . Stable for 1 week at 4  C. 5. Hemogenic endothelium progenitor cell differentiation medium: IF9S basal medium with 10 μM SB431542 (R&D Systems), 50 ng/mL VEGF (R&D Systems), 50 ng/mL bFGF (PeproTech), and 50 ng/mL SCF (R&D Systems). Stable for 1 week at 4  C. 6. Hematopoietic progenitor cells medium: IF9S basal medium with 50 ng/mL VEGF (R&D Systems), 50 ng/mL bFGF (PeproTech), 50 ng/mL SCF (R&D Systems), 10 ng/mL IL-3 (R&D Systems), 50 ng/mL IL-6 (R&D Systems), and 20 ng/mL TPO (Sino Biological). Stable for 1 week at 4  C.

2.3 Bioreactor Equipment

The random positioning machine (SM-31J) for cell culture is designed by the National Center of Space Science, Chinese Academy of Sciences, which is a microgravity simulated device. The device included two parts: (1) biaxial drive rotator, which can support 3D rotation of cell culture vessel (Fig. 1a), and (2) the electronic control system, aims to supply power and to adjust the rotation speed of rotator (Fig. 1b).

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Table 1 Formulation for IF9S medium

Medium component (stock concentration) Source

Volume added (250 mL final volume)

Final concentration

IMDM

Iscove’s modified Dulbecco’s medium (IMDM), no phenol red (Gibco, cat# 21056023)

117.25 mL

–

F12

Ham’s F-12 nutrient mix, GlutaMAX supplement 117.25 mL (Gibco, cat# 31765-027)

–

PVA (5%)

Polyvinyl alcohol (Sigma-Aldrich, cat# P8136-250G)

50 μL

10 mg/L

Lipids (100)

Chemically defined lipid concentrates (Gibco, cat# 11905031)

250 μL

0.1% (vol%)

ITS-X (100)

Insulin–transferrin–selenium–ethanolamine (Gibco, cat# 51500-056)

5 mL

2% (vol%)

αMTG (1.3% in IMDM)

Monothioglycerol (Sigma-Aldrich, cat# M6145-25ML)

750 μL

40 μ/L

AA2P (5 mg/mL)

Sigma-Aldrich, cat. no. A8960

3.2 mL

64 mg/L

GlutaMAX™ (100)

GlutaMAX-1 supplement (Gibco, cat# 35050-061)

2.5 mL

1% (vol%)

NEAA (100)

MEM nonessential amino acids solution(100) (Gibico, cat# 11140-035)

2.5 mL

1% (vol%)

Pen-strep (5000 U/mL)

Gibco, cat# 15070-063

1.25 mL

0.5% (vol%)

Fig. 1 A random positioning machine (RPM). (a) Biaxial drive rotator. (b) Electronic control system. (c) The T12.5 flasks in biaxial drive rotator were placed into CO2 incubator for cell cultivation

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1. Blocking solution: 5% bovine serum albumin (BSA, Cat# A1933, Sigma) in PBS or 5% donkey serum in PBS. Store at 20  C. 2. Primary antibody: PE Mouse Anti-Human CD31 (Cat# 560983, BD), FITC Mouse Anti-Human CD34 (Cat# 560942, BD), APC Mouse Anti-Human CD43 (Cat# 560198, BD), and Goat Anti-Human Brachyury Polyclonal Antibody (Cat# AF2085, R&D systems). 3. Secondary antibodies: Alexa Fluor® 568 Donkey Anti-Goat secondary antibody (Cat# A-11057, ThermoFisher Scientific).

2.5

3

Other Material

T12.5 cell culture Flasks (TCF-012-025, JET BIOFIL), 4% PFA (Cat# P-6148, Sigma) in 100 mL PBS, 5% Normal Donkey Serum (Cat#ab7475, abcam) in PBS, 0.1–1 μg/mL of Hoechst 33342 (Cat# B2261, Sigma) in PBS, Nikon Ti microscope (Nikon, Japan).

Methods

3.1 Feeder-Free Expansion of Human PSCs

Before differentiation of human ESCs, undifferentiated feederdependent human ESCs are cultured in feeder-free and chemically defined conditions with the use of TeSR™-E8™ medium (STEMCELL Technologies) on Matrigel Matrix HESC-qualified (BioCoat) coated plates as the protocols described in the manual. We can skip this step if human ESCs are maintained in TeSR™-E8™ medium. The following instructions are used in our experiments. 1. Before human ESC thawing or passing, coat a new 6-well plate with Matrigel Matrix solution (see Subheading 2.2), and preincubate at room temperature (25  C) for at least 1 h before use (see Note 1). 2. Remove the Matrigel Matrix solution using a pipette or by aspiration. 3. Add 2 mL TeSR™-E8™ medium to 6-well plate. 4. Put the small cell aggregate mixture (500–1000 aggregate/ wells) onto the 6-well plate containing TeSR™-E8™ and observe the state of cell aggregate under the microscope to ensure the desired cell density. 5. Place the 6-well plate in a 37  C CO2 incubator. Move the plate in a back-and-forth and side-to-side motions to evenly distribute the small cell aggregates. 6. Change medium daily with TeSR™-E8™ and monitor cell growth daily until the cell next passaging or cell differentiation.

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3.2 Mesoderm Induction from hPSCs

1. Before mesoderm induction, coat each T12.5 flasks with 1.5 mL diluted corning Matrigel Growth Factor Reduced Basement Membrane Matrix (see Subheading 2.2) at room temperature for 30 min and then incubated the flasks in 37  C for another 30 min. 2. At the time of cell passaging in Subheading 3.1, digest the hPSCs colonies into small cell aggregates by ReLeSRTM (STEMCELL Technologies) at 37  C for about 3 min. According to the manual described (see Note 2). 3. Stop digestion with TeSR™-E8™, pat the cell dish gently to make sure the small cell aggregates fall off from the bottom of the dish. 4. Harvest the cell aggregates to 15 mL centrifuge tube and the cell aggregate suspension is pipetted up and down gently about 3–5 times to adjust the size of aggregates to 100–200 cells/ aggregate (see Note 3). 5. Centrifuge the harvested aggregates at 500 rpm for 5 min at room temperature, remove the supernatant of 15 mL centrifuge tube and resuspend the small cell aggregates in 2 mL of TeSR™-E8™. 6. Remove the liquid of the Corning coated 12.5 flasks and add 2 mL TeSR™-E8™ into the flask. 7. Transfer the cell aggregates onto Corning Matrigel-coated T12.5 flask at a density of about 100 aggregates/cm2 in TeSR™-E8™ (see Note 4). 8. Place the flasks in CO2 incubator (37  C, 5% CO2 and 95% humidity) and shake the flasks in several quick sides to side, forward to back motions to uniformly distribute the aggregates. 9. Check the growth status of the cell colonies on the second day, if the size of colonies exceeds 300 μm that should be ready to be replaced into differentiation medium. 10. Aspirate medium from the T12.5 flask and add 5 mL of mesoderm induction medium (see Subheading 2.2), place the flasks in an incubator for culturing 2 days. 11. After 2 days of culture in mesoderm induction medium, we have found that almost all of the colonies exhibit loose colony appearance and elongated cell morphology (Fig. 2a) and give rise to differentiated Brachyury+ cells (Fig. 2b).

3.3 Generation of Hemogenic Endothelium Progenitor

After 2 days of mesoderm induction, the cells are further cultured by changing with hemogenic endothelium progenitor cell differentiation medium (see Subheading 2.2). Some of the T12.5 flasks with mesoderm-induced cells are put into the biaxial drive rotator. The

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Fig. 2 Mesoderm induction from human embryonic stem cells (hESCs). (a) Representative morphology of differentiated cells after 2 days culture. Scale bars, 100 μm. (b) Representative immunostaining images of day 2 cells for brachyury. Scale bars, 50 μm

T12.5 flasks and the biaxial drive rotator are placed into CO2 incubator for cell cultivation (Fig. 1c). 1. Aspirate the mesoderm induction medium and add 30 mL hemogenic endothelium progenitor cell differentiation medium into the T12.5 flasks of RPM group and add 5 mL medium into the T12.5 flasks of control group (see Note 5). 2. Put the T12.5 flasks of the RPM group into the biaxial drive rotator. 3. Place the biaxial drive rotator with T12.5 flasks and the T12.5 flasks of the control group into the CO2 incubator at 37  C with 5% CO2 and 95% humidity for cell inoculation and cultivation. 4. Turn on the power of the electronic control system and set it in random models of speed (0.1–10 rpm). 5. Change the medium with a half-volume fresh hemogenic endothelium progenitor cell differentiation medium every other day (see Note 6). 6. After 3 days of culture in hemogenic endothelium progenitor cell differentiation medium, the cell with endothelial morphology and tube-like structures emerged in the RPM group (Fig. 3a). Some differentiated cells contained round cells with a “grape-like” clusters (Fig. 3a, arrow). The differentiated cells are characterized by surface marker CD31 and CD34. 3.4 Generation of Hematopoietic Progenitor Cells

For the further induction of hematopoietic progenitor cells, the cells are growth in hematopoietic progenitor cells medium for another 3–5 days. 1. Turn off the power the electronic control system and carefully remove the T12.5 flasks from the biaxial drive rotator. 2. Aspirate the hemogenic endothelium progenitor cell differentiation medium and add about 30 mL hematopoietic progenitor cells medium into the T12.5 flasks of RPM group.

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Fig. 3 Hemogenic endothelium induction from human embryonic stem cells (hESCs) in RPM. (a) Representative morphology of differentiated cells after further 3 days culture. Several clusters of grape-like appear at day 5 (Arrows). Scale bars, 50 μm. (b) Representative immunostaining images of day 5 cells for CD34 and CD31. Scale bars, 50 μm. (c) Representative flow cytometry results of surface markers CD31and CD34 at day 5

3. Put the T12.5 flasks into the biaxial drive rotator and place them back into CO2 incubator for further 3 days of cell cultivation. 4. Change the medium with a half-volume fresh hematopoietic progenitor cells medium every other day. 5. After 3 days of culture in hematopoietic progenitor cells medium, many grape-like clusters are forming (Fig. 4a) and many round cells are floating in medium (Fig. 4b). The differentiated hematopoietic progenitor cells are characterized by surface marker CD34 and CD43 (Fig. 4c, d). 3.5 Characterization of Differentiated Cells Derived from hESCs in Bioreactor 3.5.1 Immunofluorescence Staining

1. At the culture of day 2, 5, and 8, the differentiated cells in flasks under different culture conditions are collected and prepared for immunostaining. 2. Remove the medium of the flasks and add PBS to the cells for washing 1–2 times. 3. Aspirate the PBS and then fix with 4% paraformaldehyde solution at room temperature for 20–30 min. 4. Wash the cells one time with PBS and the bottom of the flasks is divided into small pieces (1 cm  1 cm size) by sharp blades. 5. Put each small piece containing cells into the 24-well cell culture dish and add PBS to the well. 6. Permeabilize the cells for 20 min with 0.2% Triton X-100 and 1% donkey serum at room temperature (see Note 7). 7. Wash the cells two times with PBS and add the 5% donkey serum blocking solution into the well for 1 h blocking at 37  C. 8. Prepare the primary antibody in 5% donkey serum at dilutions recommended by the manufacturer.

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Fig. 4 Hematopoietic progenitor cells induction from human embryonic stem cells (hESCs) in RPM. Bright-field images of representative cellular morphology of hematopoietic progenitor cells with grape-like clusters (a) and with floating round cells at culture of day 8 (b). Scale bars, 50 μm. (c) Representative immunostaining images of differentiating cells in RPM at day 8 for CD34 and CD43. Scale bars, 50 μm. (d) Representative flow cytometry results of surface markers CD34 and CD43 at day 9 from normal gravity (NG) and simulated microgravity (SMG)

9. Incubate the cell in primary antibody solution for overnight at 4  C (or 1 h at room temperature with fluorescent-Dye conjugated antibodies). 10. Wash the cells three times with PBS in order to reduce the fluorescent background. 11. Prepare the mixed solution containing the second antibody (1:200 dilutions) and Hoechst 33342 (0.1–1 μg/mL) in 5% donkey serum PBS. 12. Incubate the cells in mixed solution for 1 h at room temperature.

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13. Wash the cells twice for 5 min in PBS. 14. Put one drop of mounting medium, add coverslip and seal with nail polish. 15. Observe the slides and acquire the image under the Confocal microscope. 3.5.2 Flow Cytometry

At the culture of day 5 and 8, the differentiated cells in flasks under different culture conditions are harvested and prepared for immunostaining. 1. Dissociate the cells into single cells with 0.05% trypsin and 0.1% EDTA. 2. Add trypsin inhibitor solution and centrifuge for 5 min, 500  g at 4  C and discard the supernatant. 3. Wash cells with cell washing buffer (PBS containing 5% BSA). Count the cells and resuspend in 5% BSA of PBS at 1  106 cells/mL. 4. Transfer 50 μL cell suspension to test tubes. 5. Add the diluted fluorescent conjugated, purified primary antibodies (CD31, CD34, and CD43) into test tubes at dilutions recommended by the manufacturer. Controls include: negative (no stain added), isotype control (with similarly labeled, nonspecific primary antibody). 6. Incubate on ice for 30 min in the dark. 7. Wash each sample three times with 2 mL cell washing buffer, spin down at 500  g for 5 min at 4  C. 8. Add 500 μL PBS to each pellet and resuspend cell sample. 9. Example the cell samples using FACS Calibur flow cytometer (BD). 10. Analyze the data with FlowJo software, version 10.0.7.

4

Notes 1. If not used immediately, the coated culture plate must be sealed with parafilm to prevent evaporation of the Matrigel® solution and the plates can be stored at 2–8  C for up to 1 week after coating. 2. When cell dissociation about 2 min in the incubator, you should check the cells state to make sure the colonies are loose and fall off from the bottom of the dish or flask. Do not overdigest the cells to generate a single-cell suspension. 3. Gently resuspended the cell pellet with a pipet to avoid generating single-cell suspension. Cell aggregate size can be

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adjusted by altering the number of times the cell aggregate mixture is pipetted up and down. Take care to maintain the cells as aggregates. 4. Seeding the cell aggregates at an optimal density to the flask (about 100 aggregates/cm2). Do not seed too dense or too sparse. 5. The flask should be filled with a cell culture medium and make sure the bubble is pulled out in the RPM group. 6. The power of the electronic control system should be turn off when prepare to change the cell medium. 7. This step can be omitted when we stain the cells with CD31, CD34 or CD43 antibodies.

Acknowledgments This work has been funded by the Chinese Manned Space Station Application Project (YYWT-0901-EXP-15, to XL), CAS Key Technology Talent Program (to XL), and the NSFC Grant (31471287, to XL). References 1. Kaufman DS, Hanson ET, Lewis RL, Auerbach R, Thomson JA (2001) Hematopoietic colony-forming cells derived from human embryonic stem cells. Proc Natl Acad Sci U S A 98(19):10716–10721 2. Chadwick K, Wang L, Li L, Menendez P, Murdoch B, Rouleau A, Bhatia M (2003) Cytokines and BMP-4 promote hematopoietic differentiation of human embryonic stem cells. Blood 102(3):906–915 3. Vodyanik MA, Bork JA, Thomson JA, Slukvin II (2005) Human embryonic stem cell-derived CD34+ cells: efficient production in the coculture with OP9 stromal cells and analysis of lymphohematopoietic potential. Blood 105 (2):617–626 4. Vo LT, Daley GQ (2015) De novo generation of HSCs from somatic and pluripotent stem cell sources. Blood 125(17):2641–2648 5. Choi KD, Vodyanik M, Slukvin II (2011) Hematopoietic differentiation and production of mature myeloid cells from human pluripotent stem cells. Nat Protoc 6(3):296–313 6. Wang H, Liu C, Liu X, Wang M, Wu D, Gao J, Su P, Nakahata T, Zhou W, Xu Y et al (2018) MEIS1 regulates Hemogenic endothelial generation, Megakaryopoiesis, and Thrombopoiesis in human pluripotent stem cells by

targeting TAL1 and FLI1. Stem Cell Rep 10 (2):447–460 7. Huang X, Gschweng E, Van Handel B, Cheng D, Mikkola HK, Witte ON (2011) Regulated expression of microRNAs-126/ 126* inhibits erythropoiesis from human embryonic stem cells. Blood 117 (7):2157–2165 8. Wang CY, Tang XM, Sun XM, Miao ZC, Lv YX, Yang YL, Zhang HD, Zhang PB, Liu Y, Du LY et al (2012) TGF beta inhibition enhances the generation of hematopoietic progenitors from human ES cell-derived hemogenic endothelial cells using a stepwise strategy. Cell Res 22(1):194–207 9. Kennedy M, Awong G, Sturgeon CM, Ditadi A, LaMotte-Mohs R, Zuniga-Pflucker JC, Keller G (2012) T lymphocyte potential marks the emergence of definitive hematopoietic progenitors in human pluripotent stem cell differentiation cultures. Cell Rep 2 (6):1722–1735 10. Woll PS, Morris JK, Painschab MS, Marcus RK, Kohn AD, Biechele TL, Moon RT, Kaufman DS (2008) Wnt signaling promotes hematoendothelial cell development from human embryonic stem cells. Blood 111(1):122–131

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11. Sturgeon CM, Ditadi A, Awong G, Kennedy M, Keller G (2014) Wnt signaling controls the specification of definitive and primitive hematopoiesis from human pluripotent stem cells. Nat Biotechnol 32(6):554–561 12. Wang Y, Gao J, Wang H, Wang M, Wen Y, Guo J, Su P, Shi L, Zhou W, Zhou J (2019) R-spondin2 promotes hematopoietic differentiation of human pluripotent stem cells by activating TGF beta signaling. Stem Cell Res Ther 10(1):136 13. Ran D, Shia WJ, Lo MC, Fan JB, Knorr DA, Ferrell PI, Ye Z, Yan M, Cheng L, Kaufman DS et al (2013) RUNX1a enhances hematopoietic lineage commitment from human embryonic stem cells and inducible pluripotent stem cells. Blood 121(15):2882–2890 14. Ramos-Mejia V, Navarro-Montero O, Ayllon V, Bueno C, Romero T, Real PJ, Menendez P (2014) HOXA9 promotes hematopoietic commitment of human embryonic stem cells. Blood 124(20):3065–3075 15. Wang H, Wang M, Wang Y, Wen Y, Chen X, Wu D, Su P, Zhou W, Shi L, Zhou J (2020) MSX2 suppression through inhibition of TGFbeta signaling enhances hematopoietic differentiation of human embryonic stem cells. Stem Cell Res Ther 11(1):147 16. North TE, Goessling W, Peeters M, Li PL, Ceol C, Lord AM, Weber GJ, Harris J, Cutting CC, Huang P et al (2009) Hematopoietic stem cell development is dependent on blood flow. Cell 137(4):736–748 17. Diaz MF, Li N, Lee HJ, Adamo L, Evans SM, Willey HE, Arora N, Torisawa YS, Vickers DA,

Morris SA et al (2015) Biomechanical forces promote blood development through prostaglandin E2 and the cAMP-PKA signaling axis. J Exp Med 212(5):665–680 18. Kwan W, Cortes M, Frost I, Esain V, Theodore LN, Liu SY, Budrow N, Goessling W, North TE (2016) The central nervous system regulates embryonic HSPC production via stressresponsive glucocorticoid receptor signaling. Cell Stem Cell 19(3):370–382 19. Bourgine PE, Klein T, Paczulla AM, Shimizu T, Kunz L, Kokkaliaris KD, Coutu DL, Lengerke C, Skoda R, Schroeder T et al (2018) In vitro biomimetic engineering of a human hematopoietic niche with functional properties. Proc Natl Acad Sci U S A 115 (25):E5688–E5695 20. Lundin V, Sugden WW, Theodore LN, Sousa PM, Han A, Chou S, Wrighton PJ, Cox AG, Ingber DE, Goessling W et al (2020) YAP regulates hematopoietic stem cell formation in response to the biomechanical forces of blood flow. Dev Cell 52(4):446–460. e445 21. Yang Y, Liu C, Lei X, Wang H, Su P, Ru Y, Ruan X, Duan E, Feng S, Han M et al (2016) Integrated biophysical and biochemical signals augment Megakaryopoiesis and Thrombopoiesis in a three-dimensional rotary culture system. Stem Cells Transl Med 5(2):175–185 22. Cao X, Yakala GK, van den Hil FE, Cochrane A, Mummery CL, Orlova VV (2019) Differentiation and functional comparison of monocytes and macrophages from hiPSCs with peripheral blood derivatives. Stem Cell Rep 12(6):1282–1297

Methods in Molecular Biology (2022) 2436: 67–81 DOI 10.1007/7651_2021_423 © Springer Science+Business Media, LLC 2021 Published online: 15 September 2021

Integrating Human-Induced Pluripotent Stem Cell Expansion Capability and Cardiomyocyte Differentiation Potential in a Microcarrier Suspension Culture Valerie Ho, Gerine Tong, Alan Lam, Shaul Reuveny, and Steve Oh Abstract Human-induced pluripotent stem cells are known for their high proliferation capacity as well as their ability to differentiate to different lineages (Ban et al., Theranostics 7(7):2067–2077, 2017; Chen et al., Stem Cell Res 15(2):365–375, 2015; Serra et al., Trends Biotechnol 30(6):350–359, 2012). For stem-cell-derived cardiomyocytes to evolve into a scalable therapeutic source, a large quantity of highly pure cardiomyocytes is needed. Thus, lies the challenge of defining an efficient cardiomyocyte differentiation process. This chapter describes a method to evaluate multiple human-induced pluripotent stem cell lines for their cardiac differentiation potentials before evaluating their integrated proliferation and differentiation abilities in microcarrier cultures in a spinner culture format. Key words Cardiomyocytes, Human-induced pluripotent stem cells, Microcarrier

1

Introduction Cardiac-related diseases are the one of the leading causes of death around the world [1]. Since mature cardiomyocytes (CM) are unable to regenerate to restore original functionality, the only way to restore a damaged heart is through heart transplantation [1, 2]. However, demand for such treatment exceeds the supply due to the shortage of donors [1, 2]. To satisfy the demand, stemcell-derived CMs can be considered as a potential solution to replace damaged cardiac tissue. For this reason, an optimized, scalable, efficient system of producing high-purity stem-cellderived CMs utilizing microcarriers (MC) is needed [3]. By taking advantage of human-induced pluripotent stem cells’ high proliferation as well as their ability to differentiate into CMs, the vision for a renewable source of mature CMs may be realized [4–6]. Different cell lines are shown to have different cardiac differentiation potentials as well as growth rates on microcarriers [7]. The first step is cell-line selection for high cardiac differentiation potentials on the monolayer cultures. While the second is selection for high proliferation capacity in an MC spinner culture followed by differentiation

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to CMs [8, 9]. This protocol aims to evaluate cell lines for their cardiac differentiation potential as well as describe an integrated system for their production using MC cultures.

2

Materials Here lists the minimal equipment, medium, chemicals, and reagents required for culture, differentiation, and characterization of hiPSCs toward CMs.

2.1 General Equipment and Consumables

1. Biosafety cabinet Class II, Type A2 (BSC). 2. CO2 incubator. 3. Eppendorf® New Brunswick™ S41i CO2 Incubator Shaker. 4. Corning® Four Position Magnetic Stirrer. 5. Pipettors. 6. Aspiration device. 7. Bench rocker. 8. Low-speed centrifuge (for spinning cells). 9. Laboratory water bath. 10. Phase contrast microscope. 11. NucleoCounter® automated cell counter (Chemometec). 12. Flow cytometer. 13. 15-mL and 50-mL conical tubes. 14. 40-μm Cell Strainer. 15. Tissue culture-treated six-well plates. 16. Tissue culture-treated 24-well plates. 17. 0.2 μm PES filter (for media filtration).

2.2 Media and Consumables for hiPSC Cultures

1. 60 mm tissue culture-treated culture dish. 2. Tissue culture-treated six-well plates. 3. mTeSR™1 (StemCell Technologies). 4. DMEM/F12 with glutamine and HEPES (Gibco). 5. GelTrex™ (Gibco). 6. Dulbecco’s phosphate-buffered saline without Ca2+ and Mg2+, DPBS( ) (Gibco). 7. TrypeLE-Express (Gibco). 8. ReLeSR™ (StemCell Technologies). 9. Y27632 (Rock inhibitor, RI) (StemCell Technologies).

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1. Cytodex® 1 (Cytiva). 2. Costar® ultra-low attachment (ULA) six-well plates (Corning). 3. Corning® 125-mL Disposable Spinner Flask with 70-mm top cap and two angled sidearms, sterile. 4. Sigmacote® (Sigma-Aldrich). 5. Duran® 500-mL laboratory glass bottles, with blue PP screw cap, for preparation of microcarriers.

2.4 Media and Small Molecules for CM Differentiation

1. RPMI-1640 with glucose and L-glutamine (Gibco #11875085). 2. B27 without insulin (Gibco #A1895601). 3. L-Ascorbic acid 2-phosphate (Sigma-Aldrich # A8960). 4. CHIR99021 (CHIR) (Tocris). 5. Endo IWR1 (IWR) (Tocris).

2.5 Antibodies and Reagents 2.6 Preparation of Extracellular Matrices for hPSCs Cultures

See Table 1.

1. To prepare GelTrex™, #MAN0007332 Rev. 3.0.

refer

to

Gibco’s

user

guide

2. Briefly, thaw a 5-mL vial of Geltrex™ LDEV (lactose dehydrogenase elevating virus) free reduced growth factor basement membrane matrix overnight at 4  C. 3. Add 50 mL of cold DMEM/F12 medium to a 50-mL conical tube. Dilute 500 μL of Geltrex™ into the cold DMEM/F12

Table 1 Antibodies and reagents for cardiomyocyte characterization Pluripotency markers Oct3/4 Tra-1-60 Cardiac specific markers Hepatocyte nuclear factor 4 alpha (HNF4a) Mesenchymal fibroblast marker CD44 Cardiac transcription factor, Homeobox protein NKX-2.5 (NKX2-5) Cardiac troponin T (cTnT) Myosin regulatory light chain 2, atrial isoform (MLC2a) Secondary antibodies FITC-conjugated goat anti-mouse IgG FITC-conjugated goat anti-rabbit IgG Reagents Para-formaldehyde 1% BSA Triton X-100

R&D Systems Biolegend

#MAB1759 #330602

Cell signaling Cell signaling Cell signaling

#3113S #3570 #8792

ThermoScientific Synaptic systems

#MA5-12960 #311011

ThermoScientific ThermoScientific

#A11029 #A11034

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medium (1:100) and quickly mix thoroughly by inverting the tube several times. 4. Plate 3 mL per 6 cm plate or 2 mL per well of a six-well plate. Store the coated plates at 4 C prior use. 5. Aspirate diluted Geltrex™ solution and add pre-warm medium before use. 2.7 Preparation of Cell Culture Medium

See Tables 2 and 3.

2.8 Preparation of Small Molecules

See Table 4. 1. Sigmacote a 500-mL glass bottles with Sigmacote® siliconizing reagent (Sigma-Aldrich), prevents the microcarriers from adhering to the glass surface during storage. For coating protocol, refer to Sigma-Aldrich Product information sheet: VNC, MAM 10/16-1.

2.9 Preparation of Microcarriers

Table 2 500 mL hiPSC culture medium (refer product information sheet, Document #10000003789-PIS_03) mTeSR™1 Basal medium

400 mL

mTeSR™1 5 supplement

100 mL

Store medium at 2–8  C up to 4 weeks

Table 3 1 L Cardiomyocytes (CM) differentiation basal medium (RPMI/B27-IN) Medium

Amount added

RPMI-1640 medium (with glucose and L-glutamine)

1L

50 B-27™ supplement, minus insulin

20 mL

L-ascorbic

213 mg

acid 2-phosphate sesquimagnesium salt hydrate

Dissolve completely and filter medium using 0.2 μm PES filter Store medium at 2–8  C up to 2 weeks

Table 4 Preparation of small molecules Small molecule

Stock amount (mg)

Working solution (mM)

Volume of DMSO used (mL)

Y27632 (RI)

10

10

3.122

CHIR99021 (CHIR)

25

12

4.152

Endo IWR1

10

2.5

Mix well and make 25–50 μL aliquots, to be stored at

9.76 

20 C

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2. Cytodex 1 microcarriers (Cytiva). For preparation of Cytodex 1 stock, refer to Cytiva’s related documents “Instructions for use - 18111979 AE” (see Note 1).

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Methods All procedures are carried out in a Biosafety cabinet Class II, Type A2 (BSC), unless otherwise stated. In addition, all cell incubations and culturing are done in a 5% CO2 incubator at 37  C unless otherwise stated. All spinner cultures are agitated at 25–30 rpm (see Note 2) unless otherwise stated.

3.1

hiPSC Culture

1. hiPSCs are grown as 70–90% confluent monolayers on Geltrex®-coated 60-mm dishes in mTeSR™1, as described in the manufacturer’s Product Information Sheet, Document #10000003789-PIS_03. Passaging is performed at a ratio of 1:10 every 6–7 days by ReLeSR™, as described in the manufacturer’s Document #28207 Version 1_4_0.

3.2 Screening for High Cardiac Differentiation Potency hiPSC Lines (Fig. 1)

The optimal concentration of CHIR may vary between cell lines based on their sensitivity to the treatment. Therefore, an initial test of treating cells at various concentrations (0–14 μM) [3, 7] is needed.

3.2.1 hiPSC Seeding for CM Differentiation

1. When hiPSCs (60-mm dish) reach 70–80% confluence, aspirate the spent mTeSR™1 and wash cells once with 4 mL of DPBS ( ). 2. Dissociate the cells into single-cell suspension by TrypeLE™Express, as described in the manufacturer’s user guide Pub. No. MAN0007321 Rev. 2.0 (see Note 3). 3. Count the live cell number using NucleoCounter® automated cell counter, as described in the manufacturer’s Application Note No. 0258 (Rev 1.2) or other cell counting methods (e.g., Trypan blue staining with hemocytometer).

Fig. 1 Screening for high cardiac differentiation potency hiPSC lines

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4. Seed 5  105 cells in each well of the Geltrex®-coated six-well plate. Culture the cells at 37 C in CO2 incubator until 95–100% confluency. 3.2.2 CM Differentiation (Monolayer Method)

The protocol described here is an adaptation of a previously described CHIR-based monolayer method [3, 7] (see Fig. 1). 1. For start of differentiation, grow cultures to 95–100% confluence in each well of a six-well tissue culture. 2. Wash the hiPSCs twice with 5 mL of DPBS( ) before differentiation. 3. To screen the working concentrations of CHIR for CM differentiation, incubate the cells with 5 mL of CM differentiation basal medium (RPMI/B27-IN, see Table 3) with 0–14 μM of CHIR [7] for 24 h (see Notes 4 and 5). 4. Day 1: After 24 h, aspirate the spent media and gently add 5 mL of fresh RPMI/B27-IN, incubate for another 24 h. 5. Day 2: Aspirate the spent media and gently add 5 mL of RPMI/B27-IN containing 2.5 μM of IWR-1, an optimal concentration as previously reported [3, 7], then incubate for 48 h until day 4. 6. Day 4: Aspirate the spent media and gently add 5 mL of fresh RPMI/B27-IN. Thereafter, conduct media change with RPMI/B27-IN every day until day 14 (see Note 6). 7. Day 14: Dissociate the cells into single-cell suspension by TrypeLE™-Express, as described in the manufacturer’s user guide Pub. No. MAN0007321 Rev. 2.0 (see Note 7), for characterization using flow cytometry.

3.2.3 Characterization of hiPSC-Derived CM from Monolayer by Flow Cytometry (Fig. 2)

1. Count the cell number using NucleoCounter® automated cell counter, as described in the manufacturer’s Application Note No. 0258 (Rev 1.2). 2. Prepare a suspension of at least 1  106 cells/mL and fix the cells with 500 μL 4% PFA solution for 15–20 min at room temperature (see Note 8). 3. Remove the PFA by centrifugation (300–400  g for 3 min).

Fig. 2 Preparation of cells for flow cytometry

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4. Resuspend cells in 100 μL of 1% BSA/Triton X-100 buffer containing cardiac-specific primary antibodies (see Table 1). Incubate for 30 min at room temperature. 5. Centrifuge the stained cell suspension at 300–400  g for 3 min, then discard the supernatant. 6. Wash once with 1% BSA/PBS buffer. 7. Resuspend in 100 μL of secondary antibody (1:500) (see Table 1) in 1% BSA/PBS buffer. Incubate for 30 min at room temperature in dark. 8. Centrifuge the stained cell suspension at 300–400  g for 3 min. Discard the supernatant. 9. Resuspend cells in 200 μL of 1% BSA/PBS buffer. Keep cells on ice in dark until ready to analyze by flow cytometry. 10. The hiPSC line(s) with high cardiac differentiation potency (i.e., high expression of NKX2-5, cTnT, and MLC2a, but low in HNF4a and CD44) will be selected for testing of expansion in microcarrier culture (see Note 9). 3.3 Testing of Cell Expansion in Microcarrier Cultures 3.3.1 Preparation of GelTrex-Coated Microcarriers

1. To prepare Geltrex™-coated microcarriers (Cytodex® 1): Transfer 33.7 mL of cold DMEM/F12, 5 mL of Cytodex® 1 (MC) suspension (50 mg of Cytodex® 1) and 1.3 mL of Geltrex™ [8–10] into the 50-mL conical tube (see Note 10). 2. Place the centrifuge tube horizontally onto bench rocker at 4  C for overnight gentle agitation. 3. After agitation, store in 4  C until ready to use. 4. Pre-warm Geltrex™-coated MC (step 3) in 37  C water bath for about 20–30 min. Spin down in a centrifuge at 300–400  g for 5 min. 5. Carefully aspirate supernatant from conical tube (see Note 11). 6. Resuspend the coated MC with 25 mL of mTeSR™1 supplemented with 10 μM Y-27632. 7. Transfer all 25 mL of resuspended MC into a Corning® 125-mL Disposable Spinner Flask and put the flask on a Corning® Four Position Magnetic Stirrer placed in a 37 C CO2 incubator.

3.3.2 Seeding Cells into Spinner Flask

1. Dissociate the selected hiPSCs (with high cardiac differentiation potency chosen in Subheading 3.2) into single-cell suspension by TrypeLE™-Express, as described in the manufacturer’s user guide Pub. No. MAN0007321 Rev. 2.0. Ensure there are no cell clumps (see Note 12). 2. Count the live cell number using NucleoCounter® automated cell counter, as described in the manufacturer’s Application

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Note No. 0258 (Rev 1.2) or other cell counting methods (e.g., Trypan blue staining with hemocytometer). 3. Seed 1  107 cells in 5 mL mTeSR™1 with 10 μM Y-27632 into the 125-mL Spinner Flask containing Geltrex™-coated MC prepared in Subheading 3.3.1. Now total volume of medium in the flask is 30 mL. 4. Place the flask in static condition at 37 C in CO2 incubation for 4–6 h to allow initial cell attachment. 5. After 6 h of static incubation, top up with mTeSR™1 (containing 10 μM Y-27632) into the flask to a total volume of 50 mL. 6. Place the spinner back to the magnetic stirrer in the CO2 incubator, and the culture is then stirred for 6 days [9, 10] (see Note 13). 7. Change 80% of medium daily: Take out the spinner flask and put in static for 5 min to allow the MC aggregates to settle down by gravity. Then remove 80% (40 mL) of the spent medium by a 25-mL pipette carefully without disturbing the MC aggregates. Subsequently replenish 80% (40 mL) of fresh mTeSR™1 into the flask. Place the spinner back to the magnetic stirrer in the CO2 incubator and continue culture at stirring of 25–30 rpm. 3.3.3 Cell Counting and Imaging during Cultivation

1. Take 1 mL of sample from the spinner culture daily for cell counting and imaging: Transfer the spinner flask from the incubator to a BSC, placing the flask on a stir plate set at 25–30 rpm. With the culture in the stir mode, remove the cap from one sidearm of the flask. Take out about 1 mL of the culture using 10-mL serological pipette. Then close the cap and place the spinner flask back to the magnetic stirrer in the CO2 incubator and continue the culture at stirring of 25–30 rpm. 2. For cell counting, take out 2 100 μL aliquots (using blunt 200-μL tips) from the 1 mL sample and transfer to 1.5-mL tubes (see Note 14). Count the cell number as aggregates, using NucleoCounter® automated cell counter, as described in the manufacturer’s Application Note No. 0262 (Rev 1.1) or other cell counting methods (see Note 15). 3. For cell imaging, take out around 500 μL of aliquot (using blunt 1000-mL tips) from the 1 mL sample and transfer to a well of 24-well plate. Visualizing the cells under phase-contrast microscopy. At least 10 pictures should be taken in order to measure the aggregate size using image analysis software, e.g., Image-J (see Note 16).

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Fig. 3 Evaluating cell-line proliferation in a microcarrier spinner culture 3.3.4 Evaluation the Pluripotency of hiPSCs in Spinner Flask Cultures by Flow Cytometry (Fig. 3)

1. To evaluate the pluripotency of spinner-grown hiPSCs after 6 days of expansion: Apart from taking 1 mL of sample for cell counting and aggregates imaging, take 2 mL of sample from the spinner culture, as described in Subheading 3.3.3, step 1. 2. Allow MC aggregates to settle and gently remove cell culture medium without distributing the MC aggregates. 3. Wash the MC aggregates two times with DPBS( ). 4. Add 2 mL of warm TrypLE™Express and incubate at 37  C for 5–7 min on a 75 rpm shaker for cell dissociation to obtain single-cell suspension. 5. After incubation, pipet up and down to break clumps and detach cells from MC, and dilute cell suspension by adding 2 mL of DPBS( ). 6. Separate the single cells from MC by using a 40-μm strainer. 7. Centrifuge the single cells at 300–400  g for 3 min. 8. Wash the cells two times with DPBS( ). 9. Perform flow cytometry as described in Subheading 3.2.3 (see Fig. 2) with pluripotency markers (see Table 1), 10. The expression of pluripotency markers Oct3/4 and Tra-1-60 should both be above 90% [3].

3.4 Cardiac Differentiation in Microcarrier Spinner Cultures (Fig. 4)

Production of cardiomyocytes in an integrated bioprocess of stem cell expansion and differentiation in MC spinner cultures. The cell line with high cardiogenic potential (Subheading 3.2) and high proliferative potential in MC spinner culture (Subheading 3.3) will be used.

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Fig. 4 The cardiac differentiation process in a spinner culture

This is an 18-day protocol, starting from cell expansion to cardiomyocytes differentiation. Through the differentiation process, take 1 mL of sample from the MC spinner culture daily to measure cell number, viability, and aggregate sizes, as described in Subheading 3.3.3. 1. Expand hiPSCs in MC spinner culture as described in Subheading 3.3. 2. After 6 days of expansion, wash the MC suspension twice with 20 mL of DPBS( ) using a serological pipet in the spinner flask (see Note 11). 3. Add 50 mL of RPMI/B27-IN containing the optimal concentration of CHIR determined in Subheading 3.2. Incubate the culture for 24 h in static conditions. 4. Day 1: After 24 h, wash the culture two times with 20 mL of fresh RPMI/B27-IN, using a 25-mL serological pipet (see Note 11), and incubate for another 24 h under agitation on the magnetic stirrer with 25–30 rpm in CO2 incubator. 5. Day 2: Aspirate the spent media and gently add 50 mL of RPMI/B27-IN containing 2.5 μM of IWR-1 [3, 7], then incubate for 48 h in static conditions until day 4. 8. Day 4: Aspirate the spent media and gently add 50 mL of fresh RPMI/B27-IN. Thereafter, conduct 50% media change (i.e., 25 mL) with RPMI/B27-IN daily until day 12. As mentioned earlier, remember to take 1 mL of sample from the MC spinner culture daily to measure cell number, viability, and aggregate sizes (see Note 17). 9. Day 12: Apart from taking 1 mL of sample for cell counting and aggregates imaging, take another 10 mL of sample from the spinner culture for flow cytometry to determine the expression of cTnT cardiac marker (see Table 1). Briefly, dissociate the cells from MC as described in Subheading 3.3.4 (see Note 18) and perform flow cytometry with anti-human cTnT antibody, as

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described in Subheading 3.2.3. cTnT expression should be above 80% [3] for successfully differentiated cardiomyocytes (see Note 17). 10. Take another 2 mL of sample from the spinner culture and transfer a six-well ULA plate, and then determine the percentage of aggregate beating under microscopy (see Note 17). 11. Dissociate rest of cells from the spinner culture as described in Subheading 3.3.4 (see Note 18). Cryopreserve the cells in CryoStor® CS10 Cell Freezing Medium until further characterization, e.g., patch-clamp recording, followed by the instruction of the manufacturer’s Product Information Sheet Document #10000000383 version 01.

4

Notes 1. MCs can take up to a few minutes to fully settle. Once settled, keep track of the amount of supernatant that is removed. Replace with the same volume of DPBS( ) so the total volume does not change during the washing process. Some MC loss during wash is normal however should be minimized. 2. Agitation speed has room for adjustment. Start the speed at 25 rpm and ensure MCs are properly suspended and not settling in the middle of the spinner flask (see Fig. 5). If MCs remain improperly suspended, increase the agitation speed up to 30 rpm. 3. Cells should be detached easily after treatment with TrypeLE™-Express, if many cells remain adherent after pipetting, the plate can be incubated in 37  C for another 1 or 2 min. Gentle pipetting is sufficient to remove cells from the culture surface; rigorous pipetting will result in more cell death.

Fig. 5 Illustration of proper suspension of microcarriers

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4. It is normal to observe significant cell death after 24 h. However, if there are little to no cells attached to the well, the cells had not reached appropriate confluence (95–100%) before differentiation. Seed cells again and wait the cells to reach 95–100% confluence before CHIR treatment. 5. Record the time when CHIR is added as the medium must be changed at exactly 24 h later. 6. Cells with transparent and web-like morphology will appear, and cell death will continue to be observed during this period as well. 7. Cardiomyocytes require longer incubation times before successfully detaching. Using a preferred single-cell dissociation reagent incubate for longer time compared to normal monolayer cultures. After 15 min of incubation, attempt to remove cells from cell attachment surface by pipetting. However, if cells remain attached, place plate back into incubator to incubate for another 15 min. Stop incubation once cells easily detach after pipetting. 8. Cardiomyocytes can be clumpy despite dissociation, upon noticing clumps after fixing cells, consider filtering cells before running flow cytometry to avoid damaging the machine and producing false positives. 9. As a guideline, successfully differentiated hiPSCs lines have a low expression of HNF4a (below 40%) and CD44 (below 40%) and high expression in NKX2-5 (60–80%), cTnT (80–83%), and MLC2a (65–70%) [3]. 10. Work by adding the largest volume reagent (DMEM/F12) first, followed by the Cytodex® 1 MCs then Geltrex™. To prepare Geltrex™, refer to Gibco’s user guide #MAN0007332 Rev. 3.0. Geltrex™-coated MCs are always prepared at least a day before use and can be stored in 4  C until ready for use. 11. Use a serological pipet to remove supernatant instead of aspirating, Cytodex® 1 compacts weakly and will be easily removed if aspirating. 12. In cases where cell clumps are seen but when resuspension is not working, filter cells through a 40-μm cell strainer. This is to ensure a complete single-cell suspension. 13. When observing cells over the course of 6 days, ensure cells are not over-confluent on MCs. Robust and fast-growing cell lines can be evaluated earlier after 5 days [3]. 14. It is recommended to cut tips when pipetting MC clumps to ensure clumps are not stuck. When obtaining cell counts, it is recommended to use a cut tip to properly dissociate cells.

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15. Several methods can be used for cell counting in MC cultures. The simplest method is to take sample from the culture, spin down, aspirate the medium from the sample, wash the pellet with DPBS( ), aspirate, and add TrypeLE™-Express. Allow cells to incubate in TrypeLE™-Express until they dissociate from the MC surface (generally 10 min at room temperature), then pipette to obtain a single-cell suspension. Cells can then count with a hemocytometer using Trypan blue. Microcarriers generally do not get under the cover slip, so they will not interfere with the count. 16. Aggregate sizes should be increasing over the course of 6-day cell expansion (see Fig. 6), ideally reaching 0.3–0.42 mm2 in size after 6 days [3]. Aggregates will not be uniform in size, measure a few aggregates of different sizes to properly evaluate these cultures. Do note that cell aggregate sizes serve as a guideline, and sizes may be larger if they are not due to overconfluency of cells but rather more MCs encapsulated into an aggregate. In lieu, cell counts on day 6 of cell expansion should be tenfold or more than the original seeded amount [3] (see Fig. 6). Refer to Fig. 7 that illustrates an example of successfully attached and proliferating cell lines on MCs and the aggregate size present after 6 days of culture. This is in comparison to a cell line that failed to attach and proliferate on MCs even after 6 days of culture. 17. To evaluate successful cardiomyocyte differentiation, aggregate sizes should be increasing after Day 1 of differentiation (see Fig. 8). In addition, cell counts should show a steady increase after day 3 of differentiation (see Fig. 9). CM yields are calculated by multiplying percentage of cells positive for cTnT with

Fig. 6 (a) Cell counts of a well-expanding hiPSC cell line on Cytodex 1 microcarriers [8]. (b) Aggregate sizes of a well-expanding hiPSC cell line on Cytodex 1 microcarriers [3]

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Fig. 7 (a) Example of hiPSCs properly attaching and proliferating into large clumps. (b) Example of hPSCs improperly attaching and not proliferating after 6 days of expansion on Cytodex 1 microcarriers in a spinner culture system

Fig. 8 Changes in aggregate size over the course of cardiac differentiation on Cytodex 1 microcarriers [3]

Fig. 9 Cell counts over the course of cardiac differentiation on Cytodex 1 microcarriers [11]

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the total live cell count obtained on day 12, good CM yields fall around 1.9  106/mL [3, 9]. Percentage of cells beating falls around 70% [12]. 18. Attempt to break MC clumps and dissociate cells by pipetting after 5 min incubation is complete. If MC clumps remain, place back into 37  C for longer incubation. It can take up to 2 h for cardiomyocytes to properly detach and dissociate from MC clumps depending on the original hiPSC line used. If too much pipetting is required to remove the cells or if clumps remain stubborn, incubate it for more time at 37  C on a shaker at 75 rpm. References 1. Jafarkhani M, Salehi Z et al (2018) Strategies for directing cells into building functional hearts and parts. Biomater Sci 6 (7):1664–1690. https://doi.org/10.1039/ c7bm01176h 2. Garbern JC, Lee RT (2013) Cardiac stem cell therapy and the promise of heart regeneration. Cell Stem Cell 12(6):689–698. https://doi. org/10.1016/j.stem.2013.05.008 3. Laco F, Lam ATL et al (2020) Selection of human induced pluripotent stem cells lines optimization of cardiomyocytes differentiation in an integrated suspension microcarrier bioreactor. Stem Cell Res Ther 11:188. https://doi. org/10.1186/s13287-020-01618-6 4. Ban K, Bae S et al (2017) Current strategies and challenges for purification of cardiomyocytes derived from human pluripotent stem cells. Theranostics 7(7):2067–2077. https:// doi.org/10.7150/thno.19427 5. Chen VC, Ye J et al (2015) Development of a scalable suspension culture for cardiac differentiation from human pluripotent stem cells. Stem Cell Res 15(2):365–375. https://doi. org/10.1016/j.scr.2015.08.002 6. Serra M, Brito C et al (2012) Process engineering of human pluripotent stem cells for clinical application. Trends Biotechnol 30 (6):350–359. https://doi.org/10.1016/j. tibtech.2012.03.003 7. Laco F, Woo TL et al (2018) Unraveling the inconsistencies of cardiac differentiation

efficiency induced by the GSK3beta inhibitor CHIR99021 in human pluripotent stem cells. Stem Cell Rep 10(6):1851–1866. https://doi. org/10.1016/j.stemcr.2018.03.023 8. Ting S, Chen A et al (2014) An intermittent rocking platform for integrated expansion and differentiation of human pluripotent stem cells to cardiomyocytes in suspended microcarrier cultures. Stem Cell Res 13(2):202–213. https://doi.org/10.1016/j.scr.2014.06.00 9. Ting S, Lam A (2018) Meticulous optimization of cardiomyocyte yields in a 3-stage continuous integrated agitation bioprocess. Stem Cell Res 31:161–173. https://doi.org/10. 1016/j.scr.2018.07.020 10. Chen AK, Chen X et al (2011) Critical microcarrier properties affecting the expansion of undifferentiated human embryonic stem cells. Stem Cell Res 7(2):97–111. https://doi.org/ 10.1016/j.scr.2011.04.007 11. Lam AT, Chen AK (2014) Conjoint propagation and differentiation of human embryonic stem cells to cardiomyocytes in a defined microcarrier spinner culture. Stem Cell Res Ther 5(5):110. https://doi.org/10.1186/ scrt498 12. Lecina M, Ting S et al (2010) Scalable platform for human embryonic stem cell differentiation to cardiomyocytes in suspended microcarrier cultures. Tissue Eng 16(6):1609–1619. https://doi.org/10.1089/ten.TEC.2010. 0104

Methods in Molecular Biology (2022) 2436: 83–111 DOI 10.1007/7651_2021_426 © Springer Science+Business Media, LLC 2021 Published online: 06 October 2021

Chemically Defined, Xeno-Free Expansion of Human Mesenchymal Stem Cells (hMSCs) on Benchtop-Scale Using a Stirred Single-Use Bioreactor Misha Teale , Valentin Jossen , Dieter Eibl , and Regine Eibl Abstract In recent years, the use of hMSCs, which may be isolated from adipose tissue among others, for the treatment of diseases has increased significantly. The cell quantities required for such therapeutic approaches, between 1012 and 1013, have thus far been predominantly produced using commercially available multi-tray systems, such as the Cell Factory (Thermo Fisher Scientific) or HYPERStack (Corning), which can be purchased with up to 40 layers. However, the handling of these planar multilayer systems is difficult, and process monitoring opportunities remain limited. Here, automated stirred single-use bioreactors provide a viable alternative to the time-consuming multiplication of cells using such planar systems, while still managing to achieve the desired clinically relevant quantities. In these stirred single-use systems, adherent cells are predominantly cultivated in suspension up to pilot scale using carrier materials, also referred to as microcarriers (MCs). This chapter describes the steps which need to be realized to guarantee successful hMSC expansion within a stirred single-use bioreactor (Eppendorf’s BioBLU® 0.3c) operated using MCs under serum- and xeno-free conditions at benchtop scale. The cultivations were performed using an immortalized human adipose-derived mesenchymal stem cell (hASC) line, hence referred to as hASC52telo, and a new chemically defined, xeno-free medium, hence referred to as the UrSuppe formulation. Spinner flask cultivations were performed under comparable process conditions. Key words BioBLU® 0.3c bioreactor, Cell expansion, hTERT immortalized ASC5Telo cells, Microcarrier, Single-use, UrSuppe

1

Introduction Human mesenchymal stem cells are plastic-adherent cells with multipotent differentiation capacity in vitro [1]. They are characterized not only by their high safety [2] but also by their remarkable therapeutic function [3–5], which includes their paracrine activity, ability to transfer mitochondria and organelles to other cells, and their application in achieving therapeutic molecule transfer using exosomes secreted during cultivation. Thus, it is not at all surprising that currently more than 1000 clinical trials are registered on www.ClinicalTrials.gov for the treatment of autoimmune and metabolic diseases, orthopedic diseases, as well as

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infectious diseases such as HIV and, more recently, COVID-19. However, in order to produce clinically relevant cell quantities for existing autologous or allogeneic therapeutic approaches, reproducible and controllable cultivation conditions are essential. In this context, single-use bioreactors have become increasingly popular in recent years [6]. The fundamental principle behind single-use bioreactors is that they make use of a plastic vessel or bag as a cultivation container, instead of re-usable stainless-steel or glass. This gives these bioreactors a major advantage over their reusable counterparts, as they can be put into operation immediately. The cultivation container is purchased pre-assembled and sterile, eliminating the need for cleaning and sterilization, while further reducing the risk of contamination during the production process [7, 8]. Moreover, the single-use cultivation containers, in which stirrers, spargers, and measuring probes may be implemented, may then be decontaminated, and discarded directly following application, significantly reducing the time between cultivations. This also reduces the validation work required for good manufacturing practice (GMP) production, resulting in more runs realized per year and increased process output. Earlier shortcomings of the technology, such as leakage of the systems (with cubic meter range working volumes) or the migration of leachables (bisphenol A or bis(2,4-di-tert-butylphenyl) phosphate) in critical concentrations [9, 10] have also since been brought under control through the use of improved materials and the appropriate detection tests [11–14]. Oosterhuis [15] and Jossen et al. [16] describe the different types of commercially available single-use bioreactors, which are mainly stirred, wave-mixed, and orbitally shaken with a maximum working volume of 6 m3, in their respective book chapters. Stirred systems, however, are considered to be the best studied and most commonly used of all the single-use bioreactor types, with such systems successfully implemented for both the bench-top and pilotscale expansion of hMSCs. 1.1 Expansion of hMSCs in Stirred Single-Use Bioreactors

As hMSCs can be used for both autologous (patient-specific) and allogeneic (off-the-shelf) cell therapies, the choice of approach has a strong influence on the required production scale and thus on the choice of the stirred single-use bioreactor system. Stirred, singleuse bioreactors in the benchtop range are usually sufficient for the production of cell quantities required for autologous therapies. In such systems, cells are generally expanded either as cell aggregates (also known as spheroids) or on MCs. The use of spheroid-based cultures means that the cells are in direct contact to one another over a variety of cell junctions, thus enabling cell–cell interaction. However, due to the heterogeneous nature of such spheroid-based cultures, this method is more commonly used for the study of complex 3D structures or for cell differentiation purposes in tissue engineering, than for mass expansion. The main motivation for

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using this approach is to avoid implementing exogenous support material, such as scaffolds or MCs, during the production process. Furthermore, the absence of the exogenous support material means that cells behave in a similar manner to native tissue [17, 18]. Spheroid formation takes place when cells self-attach and begin producing their own extracellular matrix (ECM), which not only acts as a scaffold for the cells, but also mimics cell–cell and cell–matrix signaling networks, enabling them to survive in suspension. As various studies have shown, such 3D structures can lead to increased secretion of growth and anti-inflammatory factors [19– 22]. Media composition also plays a crucial role in efficient spheroid formation, as successful cell aggregation requires supplementation of adhesion molecules (e.g., integrins, E-cadherin) to facilitate cell– cell attachment. Bartosh et al. [19] showed that proliferationrelated genes are downregulated in hMSCs after aggregation. Therefore, the maximum cell density in spheroid-based cultures is limited to the specific spheroid size and number within a bioreactor at the point of harvest. In addition, large spheroids are subject to diffusion-related limitation (e.g., oxygen, nutrients), which is a major drawback for high cell density cultures. Several studies have shown that spheroids larger than 200–300 μm tend to induce apoptosis or even unwanted spontaneous differentiation, due to nutrient or oxygen limitation in the core of the spheroids [23– 27]. In this regard, the use of MCs presents an advantageous alternative for the mass expansion of hMSCs. Many different MCs, which are usually spherical in shape, have been developed and tested over the years for the expansion of hMSCs. These different MC types vary greatly with regard to size (90–380 μm), core material (e.g., polystyrene, cellulose, dextran, gelatine), and surface coating (e.g., collagen, laminin, fibronectin). An overview of such commercially available MCs, including their physical characteristics, may be found in several reviews [28– 30]. The core material and surface coating affect not only MC sedimentation and cell growth, but also the stirrer speed required to keep MCs in suspension and ensure adequate mass transfer. Rafiq et al. [31] and Leber et al. [32] studied different MC types in stirred single-use bioreactors (benchtop scale) for hMSCs under predefined stirrer speeds (Njs ¼ Ns1). Both observed significant differences in cell adhesion, cell growth, glucose consumption, and metabolite formation depending on MC type. They found that bone marrow-derived hMSCs grew best on collagen- and recombinant protein-coated MCs, such as ProNectin® F from Solohill® and Synthemax™ II from Corning®. This is not surprising, as these MCs are coated with ECM proteins, which contain the arginine-glycine-aspartate (RGD) sequence, known to promote the attachment and growth of demanding cells. In general, cell adhesion to MCs follows a Poisson distribution, with cell– MC ratios of one, two, or three resulting in theoretical probabilities

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of unoccupied MCs of 0.365, 0.135, and 0.05, respectively. Consequently, from a theoretical perspective, cell densities for inoculation are in the range of 3–5 cells per MC. Following the cell attachment phase, which is performed statically or with periodic stirring, each MC should have at least one cell on its surface. In reality, this is not always the case. Studies have shown that uneven distribution of cells on MCs led to increased MC aggregation, reducing the efficiency of the expansion process [33, 34]. However, the success of MC-based cultivations is not only dependent on suitable MC choice but also on the cell culture medium formulation (discussed in further detail in Subheading 1.2). Various authors have shown that the stirred, single-use bioreactors of the Eppendorf BioBLU® series are suitable for MC-based cultivation of hMSCs on a benchtop scale, with maximum cell densities of up to 0.4  106 described in literature for the BioBLU® 0.3c (Fig. 1a) and BioBLU® 5c [35, 36]. The BioBLU® 0.3c, which was also used in the present study, is well suited for the suspension of MCs due to its geometric dimensions (Fig. 1b). These dimensions, namely the stirrer diameter (d), liquid height (HL), vessel diameter (D), and vessel height (H), may be used to calculate specific ratios which facilitate the comparison

Fig. 1 BioBLU® 0.3c bioreactor. (a) Picture of the BioBLU® 0.3c vessel. (b) Technical drawing with the main geometrical dimensions. D ¼ 65 mm, d ¼ 33 mm, HL ¼ 78 mm, H ¼ 117 mm

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between various bioreactor scales. For the BioBLU® 0.3c, these ratios are 0.5, 1.2, and 1.8 for d/D, HL/D, and H/D, respectively. More information regarding the dimensions of the various commercially available BioBLU® bioreactor types may be found on the manufacturer’s website. Furthermore, due to the low stirrer installation height, high relative velocities are obtained near the bottom of the reactor, which allows the suspension of MCs at low power inputs and thus at acceptable hydrodynamic loads for the cells. As previously mentioned, allogenic therapies require large quantities of cells to treat multiple patients using a single universal donor. This necessitates the use of pilot or production scale stirred single-use bioreactors to achieve therapeutically relevant cell densities in an economic and ecological manner. Due to the limitations faced by spheroid-based cultures and the associated lack of process reproducibility, this technique is of diminished importance here. In contrast, the improved reproducibility when cultivating with MCs, means their application, especially with regard to the production of hMSCs for subsequent use in allogeneic therapies, is very popular. Nevertheless, the same aspects (e.g., cell attachment, medium composition), as described for the benchtop systems, apply at this scale also. In their studies, Schirmaier et al. [37], Jossen et al. [6, 38], and Lawson et al. [39] achieved maximum cell densities of up to 0.7  106 cells mL1 when cultivating in stirred pilot-scale bioreactors (Biostat STR® 50 L and Mobius® CellReady 50 L) and using cell culture medium supplemented with 10% human platelet lysate and 5% fetal bovine serum (FBS), respectively. While it has been shown that maximum cell densities (0.04–0.4  106 cells mL1) and expansion factors achieved in stirred bioreactors using xeno- and serum-free cell culture media are still lower than those achieved in serum-containing medium, a wide variety of new serum-free media have recently come to market, paving the way for higher hMSC densities to be achieved on production scale in future. 1.2 Chemically Defined, Serum-Free Expansion of hMSCs

The most conventional cell culture media used for the production of hMSCs are based on defined basal media, such as Dulbecco’s Modified Eagle Medium (DMEM), Roswell Park Memorial Institute (RPMI) 1640, or Minimum Essential Medium (αMEM). These basal media are subsequently supplemented with expensive additives such as (a) proteins for cell adhesion, (b) lipids for cellular anabolism, (c) growth factors and hormones to promote cell growth, and in most cases, (d) an additional 10–20% (vol/vol) FBS. While the disadvantages of serum addition (e.g., high batchto-batch variability, possible contamination with prions, viruses, or zoonotic agents [40–44]) have been known for years, serumcontaining media are still used in many published academic studies to this day. Meanwhile chemically defined, serum-free media not only offer the advantage that all their components are known but also that their quantities are stoichiometrically defined, facilitating

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Table 1 Comparison of serum containing medium and chemically defined, serum-free medium Serum-containing medium Advantages

Chemically defined, serum-free medium

Protection of cells from shear stress No lot-to-lot variability of composition Inoculation with lower cell densities possible Far lower risk of contamination Less laborious purification process Easier validation and registration of product

Disadvantages Chemically not defined and serum present Lot-to-lot variability of composition Potential sources of contamination (i.e., endotoxins, mycoplasma, prions) More complicated purification process of product More difficult product validation and registration process

Cells grown in serum free medium more sensitive to hydrodynamic stresses Choice and quantity of supplement (e.g., growth factors) strongly depend on the cell type and process design

Examples

UrSuppe (Cardiocentro Ticino) Stemline® XF (Merck) StemPro™ MSC SFM XenoFree (Gibco) NutriStem MSC XF (biological industries)

Any basal medium for mammalian cell cultivation such as DMEM, α-MEM, or RPMI 1640, supplemented to a total serum content of 5–20%

their adaptation to meet specific cell or process requirements. A more comprehensive comparison between serum-containing and chemically defined serum-free media can be found in Table 1. Medium adaptability is important with regard to hMSC cultivation, as studies have shown that, depending on the mesenchymal tissue they were isolated from, the cells have very different requirements regarding cell culture medium composition [45]. This observation indicates that the development of various tissue-specific hMSC media, as opposed to a universal approach, will be necessary in future. Moreover, such aspects must be taken into consideration to ensure not only that the target cell densities required for clinical application are met but also that the desired cell quality is maintained. The implementation of chemically defined, serum-free media also comes with its own challenges, however. Due to the absence of serum as a shear stress protector, cells in dynamically mixed systems, such as stirred bioreactors, are more sensitive to hydrodynamic stress. This means that prior process characterization of the bioreactor systems, with respect to the cell culture medium and microcarrier selection, is essential and that choosing the appropriate chemically defined serum-free medium or chemically defined basal medium supplement should not be underestimated, especially when working with hMSCs. Special attention should also be paid to cell attachment efficiency, which may be lower without the addition of serum. Personal experience has shown that chemically defined cell culture medium composition has a strong influence on

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suitable MC choice and should therefore always be investigated in relation to one another, while Muoio et al. [46] were able to show that parallel development of MC and chemically defined, serumfree hMSC expansion medium resulted in enhanced cell attachment, improving the expansion process. Hence, the development of new chemically defined and serum-free hMSC media will likely result in the creation of novel MCs specifically tailored to be used in tandem. To this end, selecting the appropriate MC type for a particular expansion approach and optimizing the attachment phase in the bioreactor is as important to the development of the hMSC cultivation process as the separation of the cells from the MCs during harvest. As shown in Fig. 2a, the stem cell expansions were realized using cells from a working cell bank (WCB), which had been previously adapted to the culture medium used in the subsequent experiments. Inoculum preparation for all spinner flask (see Subheading 3.2) and bioreactor cultivations (see Subheading 3.3) was performed using T75-Flasks (see Subheading 3.1), which yielded an average of 8  106 cells per flask with a viability of 97.5% after 3 days, corresponding to an approximate doubling time of 21 h. Following harvest, the cells of four flasks were pooled to produce the inoculum. Here, we would like to point out that, with regard to

1.3 General Procedure for and Results of the Serumand Xeno-Free Expansion of Human Mesenchymal Stem Cells from Adipose Tissue in the BioBLU®

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Cell expansions (Spin (Spinner flasks or BioBLU)

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pur Cell purification

Spinner flask: Fluid dynamic characterization and determination of the suspension criteria, optimization of cell attachment and harvest efficiency

BioBLU: Fluid dynamic characterization and determination of the suspension criteria

Cell freezing and storage

(A)

(B)

Fig. 2 A general overview of the steps to be taken for the successful expansion of hMSC cells: which includes (a) the gradatim approach to the cultivation procedure as described in this book chapter and (b) the prerequisite experiments, which need to be performed prior to the expansion cultivations

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the expansion of primary hMSCs, direct inoculation of the bioreactor with the cells thawed from the WCB is strongly recommended to ensure that the population doubling level (PDL) is kept below 15. The stem cell expansions, which were accompanied by sampling, to determine cell density, viability and distribution on the MCs, substrate and metabolite concentrations (see Subheading 3.4), as well as flow cytometry analysis, and a partial media exchange (50%) on the fourth day of cultivation, were concluded with a final cell harvest. The harvest was followed by product processing and freezing to enable the long-term storage of the expanded cells, prior to their application. Cell application is, however, not the focus of the procedures described in this chapter. The stem cell expansions were preceded by, as already described by Kaiser et al. (2013) [47] and Jossen et al. (2014) [38] and shown in Fig. 2b, (a) screening to determine the optimal medium-MC combination, (b) elucidation of the fluid dynamic characteristics (tip speed, Reynolds number, specific power input, local normal and shear stresses, as well as Kolmogorov length), and (c) the identification of the cultivation vessel dependent suspension criteria (Ns1, Ns1u) under the defined process conditions (working volume, agitation rate, inoculation density, and MC concentration). Here an Ns1 of approximately 60 rpm could be determined for the spinner flask and 130 rpm for the BioBLU® 0.3c. According to Jossen (2020) [6], the NS1u criterion (or the lower limit of Ns1, where several microcarriers are located on the floor of the cultivation vessel, but none are at rest) is the most suitable for predicting the agitation rate which guarantees the maximum achievable live cell density and retention of stem cell properties during hMSC expansion. For more information regarding experimental realization, the interested reader is referred to the five part instructional video “Expansion of human adipose tissue-derived mesenchymal stem cells in stirred single-use bioreactors” (see https://youtu.be/ PZv7sx4gY9Y). Regarding the screening for optimal medium-MC combinations, results showed that ProNectin® F MCs (Pall® SoloHill®) performed best in combination with the UrSuppe medium formulation (Cardiocentro Ticino). This combination was subsequently used to optimize cell attachment to the MCs at spinner scale, conduct fluid dynamic studies of both the spinner flasks and BioBLU® 0.3c, including sedimentation studies to determine optimal agitation rates, and experiments to achieve at least a 90% harvest efficiency. The experimentally determined Ns1u criteria were met at agitation rates of 49 rpm and between 70–80 rpm for the spinner flask and BioBLU® 0.3c, respectively. The ideal point for cell harvest was determined to lie between days 5 and 6. To further increase the number of harvested cells for clinical application, the operation at the maximum working volume of the BioBLU® 0.3c vessel and a subsequent scale-up to the

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instrumented BioBLU® 5c stirred bioreactor are recommended. The growth of cells in the spinner flasks can be described with an accuracy of approximately 80% using the model developed by Jossen [6] and can therefore also be applied to cultivations undertaken in the BioBLU® 5c. As exemplified by the results of Fig. 3a, the recorded cell growth is almost identical in both cultivation systems, with the BioBLU® 0.3c achieving a maximum viable cell density of

Fig. 3 Exemplary, as well as modeled results, of the hASC expansion in 125-mL spinner flasks (n ¼ 3) and the BioBLU® 0.3c: (a) the viable cell density per mL (χ S) and viable cell density per cm2 (χ A) of the cells during expansion in the BioBLU® 0.3c (left) and spinner flasks (right). The double-headed arrow represents a 50% media exchange 4 d post-inoculation. (b) Changes in glucose (Glc), lactate (Lac) and ammonium (Amn) concentrations during the 7 days cultivation process. (c) Marker expression rates as a percentage of the total singlet cell population isolated from both cultivation vessel types and analyzed by flow cytometry and (d) images taken by microscope after staining with 40 ,6-diamidino-2-phenylindole (DAPI) to determine cell attachment and distribution, following a 7-days expansion period

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6.0  105 cells mL1 (1.7  105 cells cm2) after 6 d, while the spinner flasks required 7 d to achieve a density of 5.3  0.5  105 cells mL1 (1.5  105 cells cm2). Due to the difference in maximum cell densities achieved and the working volumes of the cultivation vessels, 70% more viable cells could be produced in the BioBLU® 0.3c, compared to a spinner flask. The slight deviation in the CD105 marker expression rates between cell populations sampled from the two cultivation vessels following the 7 d cultivation period, highlight the importance of harvesting prior to reaching the stationary phase (between 5 and 6 days) to assure cell quality. Harvesting the cells 24 h prior to reaching the stationary phase would result in viable cell densities of 3.9  105 cells mL1 for the BioBLU® 0.3c and 3.4  105 cells mL1 for the spinner flask, superseding or matching values reported for hASCs cultivated in similar stirred bioreactor systems, namely the 1.3 L Bioflo® (0.6  105 cells mL1) and 2 L UniVessel® SUB (4.1  105 cells mL1), using chemically defined xeno- and serum-free media [48, 49]. It is worth mentioning that subsequent studies should also include investigations to determine the differentiation potency of the harvested hASCs, as described by Panella et al. [50].

2

Materials During preparation of the cultivation systems, inoculum, media, and microcarriers, be sure to use at least research grade material from a reputable supplier. Diligently follow all biosafety regulations, especially when discarding waste which has come into contact with biological material. The materials listed below were necessary for the expansion of the hASCs in the single-use spinner flasks and BioBLU® 0.3c. In addition, the following standard cell culture equipment and materials were routinely used: biosafety cabinet class II, centrifuge, water bath, magnetic stirrer platform, CO2 incubators, pH meter, sterile pipettes, and pipette tips. Furthermore, the following devices were required for routine analytics: microscope (see Note 1), cell counter (see Note 2), media component analyzer (see Note 3), and flow cytometer (see Note 4).

2.1 hASC Inoculum Production in T75Flasks

1. The hASC cell line of interest (see Note 5). 2. The corresponding chemically defined, xeno-free cultivation medium prepared according to the manufacturer’s instructions and stored at 4  C in the dark until use (see Note 6). 3. T75-Flasks with filtered venting caps. 4. A CO2 incubator (see Note 7). 5. Synthemax™ II-SC working solution (see Note 8). 6. The enzymatic dissociation reagent and buffer of choice (see Note 9).

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1. A sufficient quantity of hASC prepared during the inoculum production (see Subheading 3.1). 2. The corresponding chemically defined, xeno-free cultivation medium prepared according to the manufacturer’s instructions and stored at 4  C in the dark until use (see Note 6). 3. 125 mL spinner flasks (Corning®). 4. A CO2 incubator containing a magnetic stirrer platform (see Note 7). 5. ProNectin® F MCs (see Note 10). 6. The enzymatic dissociation reagent and buffer of choice (see Note 9).

2.3 hASC Expansion in the BioBLU®

1. A sufficient quantity of hASC cells prepared during the inoculum production (see Subheading 3.1). 2. The corresponding chemically defined, xeno-free cultivation medium prepared according to the manufacturer’s instructions and stored at 4  C in the dark until use (see Note 6). 3. BioBLU® 0.3c bioreactor vessels, the corresponding DASGIP® Parallel Bioreactor System (Eppendorf), and conventional compatible pH probes. 4. ProNectin® F MCs (see Note 10). 5. The enzymatic dissociation reagent and buffer of choice (see Note 9).

2.4 Process Analytics

1. Sample drawn from the cultivation systems (see Subheadings 3.2 and 3.3). 2. The enzymatic dissociation reagent and buffer of choice (see Note 9). 3. A CO2 incubator with integrated orbital shaker (see Note 7).

3

Methods The following protocols, developed for the expansion of the hASC line hASC52Telo, include the preliminary preparation of the media and MCs, inoculum production, preparation of the cultivation systems, inoculation, sampling, analytics, and cell harvest. With minor modifications, these protocols may be adapted for the expansion of other patient adipose tissue-derived mesenchymal stem cells.

3.1 T75-Flask-Based Inoculum Production

1. Prepare 20 mL of cell culture medium (see Note 6) for the inoculum production in a T75-Flask by pre-warming it to 37  C. 2. Coat the T75-Flasks with 15 mL of Synthemax™ II-SC working solution (see Note 8).

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3. Following the coating procedure, remove the remaining Synthemax™ II-SC working solution from the T75-Flask and add the 20 mL of pre-warmed cell culture medium. 4. Expand the hASC cells (see Note 5) in T75-Flasks to produce enough cells to inoculate the desired cultivation systems. (a) Thaw the hASC cells at least 7 days prior to their expansion in the stirred systems. Remove one cryovial containing approximately 1 mL cell suspension with a density of at least 1  106 cells mL1 from the cryogenic storage tank and immediately thaw the vial in a pre-warmed water bath (37  C). Prevent the lid from touching the surface of the water bath to prevent contamination. As soon as only small ice crystals are left, wipe the vial down with a tissue soaked in ethanol. The entire contents of the cryovial should be transferred to the T75-Flask, containing the pre-warmed media, under sterile conditions to achieve a final cell density of 50,000 cells mL1 (or approximately 13,333 cells cm2). (b) Incubate the cells under static conditions at 37  C, 5% CO2, and 80% relative humidity for 6 h (see Note 7). (c) Following a 6 h incubation period, inspect the T75-Flask under the microscope (see Note 1) and determine cell attachment to the substrate. Once confirmed, replace the supernatant with fresh pre-warmed cell culture media. (d) Monitor the T75-Flask by microscope every 24 h until the cells achieve 80–90% confluency (see Note 1). At this point, passage the cells (see Notes 9, 11, and 12) into 2–5 freshly coated T75-Flasks (see Note 8), depending on the desired inoculum quantity. Inoculate each flask containing 20 mL of pre-warmed culture medium (see Note 6), at a density of 10,000 cells cm2. (e) Again, monitor the T75-Flasks every 24 h by microscope until the cells have achieved 80–90% confluency (see Note 1). Subsequently, passage the cells (see Notes 9 and 11), determine their cell density and quality (see Note 12), then transfer the required cell quantity to the cultivation systems of choice. 3.2 hASC Expansion in Corning’s Spinner Flasks

1. The following procedure describes the xeno- and serum-free expansion of hASCs in chemically defined medium and 125 mL spinner flasks (100 mL working volume) using a partial media exchange approach. Approximately 24 h prior to the inoculation of the spinner flasks, begin with preparations of the culture medium, MCs, and spinner system. (a) Pre-warm at least 150 mL of cell culture medium (see Note 6) to 37  C for the expansion procedure per spinner flask.

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(b) Subsequently, transfer at least 1 g (¼ 360 cm2) of ProNectin® F MCs (see Note 10) per spinner flask to a 100 mL Schott Flask (Duran) and suspend them in sterile Dulbecco’s phosphate-buffered saline (DPBS) to achieve a final MC concentration of 100 g L1. Autoclave the MC suspension at 121  C for 20 min. (c) Allow the suspension to cool to room temperature and the MCs to sediment at the bottom of the flask. Remove and discard the supernatant, while ensuring that no MCs are lost in the process. Replace the discarded supernatant with pre-warmed culture medium (see Note 13). Repeat this step twice to obtain a sterile ProNectin® F MC stock solution. (d) Transfer the packaging containing the spinner flask to the biosafety cabinet. Open the package and assemble the spinner flask under the bench using the appropriate technique to guarantee cultivation vessel sterility. (e) Add 90 mL of pre-warmed culture media (see Note 6) and 10 mL of ProNectin® F MC stock solution to the spinner flask. (f) Transfer the spinner flask back to the incubator containing the stirrer platform and allow the MCs to equilibrate under process conditions, i.e., 37  C, 5% CO2, 80% relative humidity and Ns1u (see Notes 7 and 14), for approximately 24 h prior to inoculation. This step also serves to ensure vessel sterility prior to cultivation. 2. Inoculation of the spinner flask. (a) Calculate the necessary volume of inoculum, following cell density and quality control (see Note 12), to achieve an initial cell density of 15,000 cells cm2 or 5.4  106 cells per spinner flask. (b) Transfer the equilibrated spinner flask to the biosafety cabinet and allow the MCs to sediment. Subsequently remove the top lid and place it stirrer side up within the biosafety cabinet. (c) Remove the volume of culture medium to be replaced with the inoculum and transfer it to a sterile 15-mL centrifuge tube (Corning®). This sample may then be used to determine the initial substrate and metabolite concentration within the spinner flask (see Note 3). Add the inoculum to the spinner flask to achieve the target starting cell density. (d) Transfer the spinner flask back to the incubator containing the stirrer platform and set the agitation rate to Ns1u for 5 min (see Note 14). Subsequently, reduce the speed to 0 rpm and allow the cells to sediment and attach to the MCs for 12 h.

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3. Initial and routine in-process control of the spinner flask during the cultivation process. (a) Following the attachment phase, transfer the spinner flask to the biosafety cabinet, open the spinner flask as previously described, resuspend the MCs using a 10-mL sterile pipette and take a homogenous sample (7 mL). Sample treatment prior to analyses is more closely described in Subheading 3.4. (b) Refasten the lid and transfer the spinner flask back to the incubator containing the stirrer to ensure process conditions can be maintained for the remainder of the cultivation. (c) Subsequent routine sampling (5 mL) should be performed every 24 h post-inoculation, especially prior to and following the medium exchange (see Fig. 3). 4. Perform a 50% media exchange 4 days post-inoculation to prevent the accumulation of inhibitory cell metabolites or the glucose concentration from falling below 5.55 mmol L1 (¼ 1 g L1). (a) Transfer the spinner flask to the biosafety cabinet and allow the MCs to sediment. (b) Open the spinner flask as previously described, then remove 50% of the supernatant (approximately 36.5 mL) and replace with fresh pre-warmed culture media (see Note 6). (c) Following the media exchange, refasten the lid of the spinner flask and return it to the incubator containing the stirrer platform to maintain process conditions (see Note 7). 5. The harvest criterion is met at least 24 h prior to reaching the stationary phase of cell growth (between 5 and 6 days post inoculation). At this point perform a final harvest (see Fig. 3). (a) Transfer the spinner flask to the biosafety cabinet and allow the MCs to sediment. (b) Open the spinner flask as previously described, then remove the supernatant without aspirating MCs and replace it with pre-warmed DPBS. Allow the MCs to sediment, then replace the DPBS with pre-warmed TrypLE Select 1 (see Note 13). (c) Return the spinner flask to the incubator containing the stirrer platform and increase the agitation rate to Ns1u for 15 min (see Notes 7 and 14). (d) Following enzymatic dissociation, transfer the spinner flask back to the biosafety cabinet, open and gently

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resuspend the suspension 10 times using a 10-mL sterile pipette, then filter the suspension through a sterile 70-μm cell strainer (Corning®) into a sterile centrifugation tube. (e) Use an equal amount of fresh pre-warmed culture media (see Note 6) to wash the interior of the spinner flask, filter it through the strainer into the sterile centrifugation tube containing the initial filtrate, then centrifuge at 300  g for 5 min. (f) Remove the supernatant and resuspend the cells in pre-warmed DPBS, then centrifuge the cells at 300  g for 5 min. Repeat this washing step twice. (g) Finally, remove the supernatant and resuspend the cells in the desired amount of either fresh pre-warmed culture media (see Note 6), for direct use, or pre-cooled cryogenic medium, for long-term cell storage at 160  C (see Note 15). 3.3 hASC Expansion in Eppendorf’s Instrumented BioBLU® 0.3c

1. The following procedure describes the xeno- and serum-free expansion of hASCs in chemically defined medium and an instrumented BioBLU® 0.3c stirred bioreactor (150 mL working volume). This method makes use of a partial media exchange approach. Approximately 24 h prior to the inoculation of the BioBLU® 0.3c, begin with preparations of the culture medium, MCs, and bioreactor system. (a) Calibrate the offset of the dissolved oxygen (DO) probe using pure N2 gas, as well as the offset and slope of the pH probe using the appropriate buffers. (b) Secure the pH probe on an open Schott flask containing sterile DPBS in such a way that more than half of the glass section is submerged. Autoclave the pH probe at 121  C for 20 min to ensure sterility prior to installation. As DO is measured optically through a membrane, the probe does not come into contact with the cultivation medium and does therefore not require autoclaving. (c) Pre-warm at least 200 mL of cell culture medium (see Note 6), per BioBLU® 0.3c vessel, to 37  C for the expansion procedure. (d) Thereafter, transfer at least 1.5 g (¼ 540 cm2) of ProNectin® F MCs (see Note 10) per BioBLU® 0.3c to a 100 mL Schott Flask (Duran) and suspend them in sterile DPBS to achieve a final MC concentration of 100 g L1. Autoclave the MC suspension at 121  C for 20 min to ensure sterility. (e) Allow the suspension cool to room temperature and the MCs to sediment to the bottom of the flask. Remove and

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discard the supernatant, while ensuring that no MCS are lost in the process. Replace the discarded supernatant with pre-warmed culture medium (see Note 13). Repeat this step twice to obtain a sterile ProNectin® F MC stock solution. (f) Transfer the packaging containing the bioreactor to the biosafety cabinet. Open the package and assemble the BioBLU® 0.3c under the bench using the appropriate technique to guarantee bioreactor sterility. (g) Remove one of the two port lids and add 135 mL of pre-warmed culture media (see Note 6) and 15 mL of ProNectin® F MC stock solution to the bioreactor. Install the sterilized pH probe, by securing it in place of the port lid. (h) Transfer the bioreactor back to the control unit and proceed to install the exhaust gas cooler, DO probe, temperature probe, pH probe cable, inlet gas piping, and mount the stirrer. Then begin operation of the bioreactor by activating the various control loops, i.e., 37  C, pH 7.2, DO >30%, 0.1 vvm overlay and Ns1u (see Notes 14, 16, and 17 for more information). (i) Allow the MCs to equilibrate under process conditions for approximately 24 h prior to inoculation. This step also serves to ensure system sterility prior to cultivation. 2. Inoculation of the BioBLU® 0.3c. (a) Set the slope of the DO probe by using the saturated culture medium as a reference to calibrate 100% DO. (b) Calculate the necessary volume of inoculum, following cell density and quality control (see Note 12), to achieve an initial cell density of 15,000 cells cm2 or 8.1  106 cells per bioreactor. (c) Deactivate the control loops. Transfer the equilibrated bioreactor to the biosafety cabinet and allow the MCs to sediment. Subsequently remove the remaining port lid and place it right side up on a sterile surface within the biosafety cabinet. (d) Remove the volume of culture medium to be replaced with the inoculum and transfer it to a sterile 15-mL centrifuge tube (Corning®). This sample may then be used to determine the initial substrate and metabolite concentration within the bioreactor (see Note 3). Add the inoculum to the bioreactor to achieve the target starting cell density. (e) Refasten the sterile port lid, transfer the vessel back to the control unit, and reactivate all the necessary control loops. Subsequently, deactivate agitation after 5 min to allow the cells to sediment and attach to the MCs for 12 h.

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3. Initial and routine in process control of the BioBLU® 0.3c during cultivation. (a) Following the attachment phase, transfer the bioreactor to the biosafety cabinet, remove the remaining port lid as previously described, resuspend the MCs using a 10 mL sterile pipette, and take a homogenous 7 mL sample (see Note 18). Subsequent sample treatment prior to the analyses is more closely described in Subheading 3.4. (b) Transfer the bioreactor back to the control unit and reactivate all the control loops to maintain the predefined process conditions, such as the Ns1u criteria for the remainder of the cultivation. (c) Subsequent routine sampling (5 mL) should be performed every 24 h post-inoculation, especially prior to and following the media exchange (see Fig. 3). 4. Perform a 50% media exchange 4 d post-inoculation to prevent the accumulation of inhibitory cell metabolites and the glucose concentration from dropping below 5.55 mmol L1 (¼ 1 g L1). (a) Transfer the bioreactor to the biosafety cabinet and allow the MCs to sediment. (b) Open the remaining port lid as previously described and remove 50% of the supernatant (approximately 61.5 mL) while ensuring that no MCs are aspirated, and replace with fresh pre-warmed culture media (see Note 6 and 13). (c) Following the media exchange, refasten the sterile port lid and return the bioreactor to the control unit and reactivate all the control loops. 5. The harvest criterion is met at least 24 h prior to reaching the stationary phase of cell growth (within 5–6 days postinoculation), at which point a final harvest may be performed. (a) Deactivate all the control loops and allow the MCs to sediment, then transfer the bioreactor to the biosafety cabinet and remove the remaining port lid of the bioreactor as previously described. (b) Proceed to remove the supernatant without aspirating MCs and replace with pre-warmed DPBS. Allow the MCs to sediment, then replace the DPBS with pre-warmed TrypLE Select 1 (see Note 13). (c) Refasten the sterile port lid and return the bioreactor to the control unit. Reattach the necessary instruments and periphery, reactivate all the control loops and set the agitation rate to Ns1u for 15 min (see Note 14).

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(d) Following enzymatic dissociation, deactivate all the control loops and return the bioreactor to the biosafety cabinet. (e) Remove the remaining port lid as previously described and gently resuspend the cell suspension 10 times using a 10 mL sterile pipette, then filter the suspension through a 70 μm cell strainer (Corning®) into a sterile centrifugation tube. (f) Use an equal amount of fresh pre-warmed culture media (see Note 6) to wash the interior of the bioreactor, filter it through the strainer into the sterile centrifugation tube containing the initial filtrate, then centrifuge at 300  g for 5 min. (g) Remove the supernatant and resuspend the cells in pre-warmed DPBS, then centrifuge the cells again at 300  g for 5 min. Repeat this washing step twice. (h) Finally, remove the supernatant and resuspend the cells in the desired amount of either fresh pre-warmed culture media (see Note 6), for direct use, or pre-cooled cryogenic medium, for the long-term cell storage at 160  C (see Note 15). 3.4 Sampling and Quality Control

This section describes the sampling procedure for the two cultivation systems and their corresponding workup, as well as basic quality control measures, which may be performed in an upstream setting. A simplified overview of the routine sample work-up is given in the Fig. 4. 1. Directly following the attachment phase, a 7 mL sample is taken to determine cellular attachment efficiency, cell distribution on the MCs, as well as the prevailing substrate and metabolite concentrations. (a) Transfer the homogenous sample to a 15-mL centrifuge tube (Corning®) for intermediate storage. (b) Remove 1 mL of the homogenous sample and transfer to a 1.5-mL microtube (Eppendorf®) and allow the MCs to sediment in both containers. (c) Transfer the supernatant to another 1.5-mL microtube in such a way as to not aspirate any MCs. This sample may then be analyzed to determine substrate and metabolite concentrations (see Note 3). (d) To the pellet remaining in the 1.5-mL microtube, add 1 mL of formalin and incubate at room temperature for 30 min. Subsequently, the sample may be either used directly to determine cell distribution and MC

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Sample processing

Aliquote 1

Aliquote 2

Aliquote 3

1 mL

2 mL

2 mL

Pellet

Pellet

Pellet

Trypsination

Trypsination

Supernatant

Filtration

DAPI analysis for determination of cell attachment and distribution

Flow cytometry for cell quality control

Cell density and viability analysis

Substrate and metabolite analysis

Fig. 4 A general overview of the sample work-up. The homogenous 5 mL sample is divided into three aliquots, which are subsequently processed separately to determine MC aggregation, substrate, and metabolite concentrations, as well as cell distribution, density, viability, and quality using various analytical approaches such as staining with DAPI

aggregation (see Note 19) or stored at 4  C for up to a week until analysis. (e) Following sedimentation of the MCs in the 15-mL centrifuge tube, gently resuspend the MCs in 1 mL of pre-warmed DPBS and transfer the suspension to a 2-mL microtube (Eppendorf®) using a 1-mL pipette and a wide orifice (1.5 mm) pipette tip. (f) Allow the MCs to sediment, then remove and replace the DPBS with 0.5 mL of pre-warmed TrypLE Select 1. (g) Place the 2 mL microtube horizontally in the incubator with integrated orbital shaking platform at 130 rpm, 25 mm, 37  C for 7 min (see Note 7). (h) Resuspend the suspension 10 times using a 1-mL pipette and standard pipette tip, then determine viable cell density (see Note 2). Attachment efficiency may then be calculated by dividing the result by the cell density of the inoculum. Be sure to account for the concentration factor (approximately 10:1, see Note 13). 2. Standard samplings of 5 mL (see Fig. 4) are performed every 24 h post-inoculation for the duration of the cultivation.

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(a) Transfer the homogenous sample to a 15-mL centrifuge tube for intermediate storage. (b) Gently resuspend the sample using a 10-mL sterile pipette and aliquot the sample into 2  2 mL and 1  1.5 mL microtubes, as shown in Fig. 4. (c) Allow the MCs to sediment in all containers. (d) Transfer the supernatant from one 2-mL microtube to a new 1.5-mL microtube in such a way as to not aspirate any MCs. This supernatant may then be analyzed to determine substrate and metabolite concentrations (see Note 3). (e) To the 1.5-mL microtube, add 1 mL of formalin. This sample, henceforth referred to as aliquot 1, may then be used to determine cell distribution, as well as MC aggregation (see Note 19). (f) Using 1 mL of pre-warmed DPBS for each, resuspend the sedimented MCs in both 2-mL microtubes. (g) Allow the MCs to sediment, then remove and replace the supernatant with 0.5 mL of pre-warmed TrypLE Select 1. (h) Place the 2-mL microtubes horizontally on an orbital shaking platform at 130 rpm, 25 mm, 37  C for 7 min (see Note 7). (i) Following incubation, resuspend the suspension in both 2-mL microtubes 10 times using a 1-mL pipette and standard pipette tip, then use aliquot 3 to determine cell density (see Note 2). Be sure to account for the concentration factor (approximately 10:1, see Note 13). (j) Based on the cell density results above, use a 70-μm cell strainer to filter the volume of aliquot 2 corresponding to a maximum of 1.5  106 cells into a 1.5-mL microtube and separate them from the MCs (aliquot 3 may also be used in combination with aliquot 2, if the latter’s volume proves insufficient). Then proceed with the flow cytometry analysis (see Note 4). 3. Determining cell distribution and MC aggregation by staining with DAPI. (a) Prepare the Perm/Stain working solution by adding DAPI working solution (see Note 20) to the Triton® X-100 working solution (see Note 21) at a ratio of 2.5:1000. In other words, 2.5 μL of DAPI working solution for every 1,000 μL of Triton® X-100 working solution. (b) Following fixation with formalin of the sample in the 1.5mL Eppendorf tube, remove the supernatant, while

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ensuring that no MCs are aspirated in the process, and replace with sterile DPBS. Repeat this washing step twice. (c) Remove the sample supernatant, as described above, then add 1 mL of Perm/Stain solution to the tube in one gentle motion, being sure to resuspend all the MCs in the process. (d) Incubate the sample for 15 min at room temperature. (e) Aspirate the supernatant in the tube and use it to resuspend the MCs in the tube. (f) Incubate for 15 min at room temperature. (g) Ensure that the MCs have sedimented, then wash the permeabilized and stained cell/microcarrier suspension with DPBS twice as mentioned previously. (h) Transfer the permeabilized and stained cell/MC suspension to the uncoated well of a six-well plate using a 1-mL pipette and a wide orifice (1.5 mm) pipette tip. (i) Add an additional 1 mL of DPBS to each well containing MC aggregates. (j) The permeabilized and stained cell/MC suspension can now be viewed under the microscope using the correct fluorescence filter (see Note 1). (k) Stained samples can be stored for up to 7 days if wrapped in aluminum and kept at 4  C. 4. Flow cytometry analysis of the sample to determine hMSC marker expression rates. (a) Following the separation of the cells from the MC, dilute the sample with 1 mL of staining buffer (see Note 22). (b) Centrifuge the sample at 400 g for 5 min. (c) Remove the supernatant, while ensuring that the cell pellet is not aspirated, then resuspend the pellet in 200 μL of staining buffer (see Note 22). Aliquot the sample and add the fluorophore-conjugated antibodies listed in Table 2. (d) Briefly vortex the samples to ensure homogeneity, then incubate the samples at 4  C for 10 min. (e) Dilute each sample by adding 1 mL of staining buffer (see Note 22) and briefly vortex to interrupt the staining procedure. (f) Centrifuge the samples at 400 g for 5 min. (g) Remove the supernatant, while ensuring that the cell pellet is not aspirated, then resuspend each pellet in 100 μL of staining buffer (see Note 22). The samples may now be analyzed using flow cytometry.

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Table 2 Cell suspension and fluorophore-conjugated antibody combinations required for the flow cytometry analyses REA control antibodies Sample ID

Cell suspension

Blank

50 μL

Isotype

47 μL

CD CD

+

47 μL



48 μL

PE

Cluster of differentiation (CD) antibodies

CD105FITC APC PE

CD90FITC

CD73APC

1 μL

1 μL

CD45PE

CD36APC

1 μL

1 μL

1 μL 1 μL 1 μL 1 μL

(h) For result evaluation using the appropriate software (see Note 4), first determine the cell population by gating the events of interest using the Blank sample (see Table 2) and a forward scatter area (x-axis) versus side scatter area (y-axis) plot. Then exclude potential cell aggregates from the cell population by gating the singlet events using a forward scatter area (x-axis) versus forward scatter height (y-axis) plot. (i) Once the singlet population has been adequately determined, use the Isotype sample (see Table 2) to gate for PE+, FITC+, and APC+ events using a PE-, FITC-, or APC-area (x-axis) versus side scatter area (y-axis) plot. Gate in such a way that the field contains only 1% of the stained population. This results in non-specifically stained cells being excluded from the evaluation process. (j) Through application of this gating strategy, the percentage of specifically stained cells in the CD+ and CD samples (see Table 2) may be determined.

4

Notes 1. Cell attachment and confluency were determined using the EVOS™ FL Auto 2 Imaging System and the corresponding software DiamondScope v2.0.2094.0 and EVOS Analysis v1.4.998.659 (Thermo Scientific™ Invitrogen™). 2. Cell density, viability, diameter, and aggregation rate were determined post-trypsination using the NucleoCounter® NC-200™ (Chemometec). This system makes use of the fluorescent dyes, acridine orange, and DAPI, to differentiate between cells with an intact and compromised membrane,

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thus elucidating the density and viability of the cell population. For further information, consult the user manual supplied by the manufacturer. 3. Substrate (glucose, L-alanyl-L-glutamine, L-glutamine, and Lglutamate) and metabolite (lactate and ammonium) concentrations were measured using the Cedex Bio Analyzer (Roche Custom Biotech). The analyzer can be fitted with various test kits, which allows the system to measure the concentration of multiple media components simultaneously. 4. The expression of specific extracellular hMSC markers serves as an indicator of cell quality. Thus, marker expression rates were analyzed at the beginning, during, and upon completion of cultivation. To determine the prevalence of these CDs, fluorophore-conjugated primary antibodies corresponding to the CD or antigen of interest (positive markers: CD73, CD90, CD105, negative markers: CD36, CD45) were used in conjunction with the MACSQuant 10 flow cytometer (Miltenyi Biotec). The evaluation of the results was performed using the FlowLogic software version 7.2.1 (Inivai Technologies Pty. Ltd.). 5. An adipose-derived mesenchymal stem cell line, isolated from a Caucasian female patient, immortalized with human telomerase reverse transcriptase (hTERT) (hASC52Telo or ATCC® SCRC-4000™) and purchased from ATCC®. 6. Chemically defined culture media guarantees reproducible cultivations due to the absence of animal-derived serum, proteins, peptides, or other components. Furthermore, the use of xenofree media is essential for the production of cell-based therapies, as it eliminates the risk of zoonotic diseases, such as bovine spongiform encephalitis, which could potentially contaminate the final product. To this end, the chemically defined, xenoand serum-free medium UrSuppe (Cardiocentro Ticino) was used for the expansion of the hASCs, as recommended by the manufacturer. See the publications by Jossen et al. [51] and Panella et al. [50] for more information. 7. In order to maintain the optimal process conditions for static cell expansion and subsequent sample work-up, both a CO2 incubator with (Minitron, Infors AG) and without (BBD 6220, Heraeus Group) an integrated orbital shaker platform (set to 5% CO2, 130 rpm, 25 mm and 37  C) were used, while for the spinner flask cultivations, an agitation platform (DuraMag, Chemglass Life Sciences) was used in addition to the latter. 8. The Synthemax™ II-SC working solution can be prepared by reconstituting Synthemax™ II-SC Substrate (Corning®) using sterile cell culture grade water (WFI) to produce a 1 mg mL1

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stock solution. The stock solution may be stored at 4  C for up to 6 months if required. The stock solution must be further diluted at a ratio of 1:40 using WFI to achieve the working solution with a final Synthemax™ II-SC concentration of 0.025 mg mL1 prior to application. Add the working solution to the cultivation vessel under sterile conditions and incubate for 2 h at room temperature. For more information, consult the self-coating protocol available on the manufacturer’s website. 9. Enzymatic dissociation of the adherent hASCs can be achieved by removing the cell culture medium supernatant from the cultivation vessel, then washing with and subsequently removing pre-warmed DPBS. Finally, add 10 mL of TrypLE Select 1 (Gibco®) to the T75-Flask, and incubate the cultivation vessel at 37  C for 5 min. For further information regarding the volumes of TrypLE Select 1 required for other cultivation vessel types, please consult the online protocol supplied by the manufacturer. Cell dissociation should be observed by microscope directly following incubation. If this is not the case and a large percentage of cells remain attached to the substrate, incubate the cultivation vessel for an additional 5 min or gently tap the side of the cultivation vessel using the back of a scissor to facilitate the dissociation process. 10. ProNectin® F MCs (Pall® SoloHill®) consist of a cross-linked polystyrene core (diameter of 125–212 μm) and a coating containing multiple copies of the human fibronectin RGD attachment domain. See the publication by Rafiq et al. [30] for more information. Recently, these MCs were transferred to Sartorius AG as part of a divestiture agreement, after which the product line was discontinued. A suitable commercially available coated polystyrene alternative with the same dimensions and density as the ProNectin® F MCs are the Low Concentration Synthemax® II MCs (Corning®). 11. The dissociated cells suspended in TrypLE Select 1 should be transferred into a 50-mL centrifuge tube (Corning®) containing 20 mL of pre-warmed cell culture media (see Note 6), centrifuged at 300  g for 5 min, resuspended in 5 mL fresh pre-warmed culture media (see Note 6) and their cell density determined (see Note 2). 12. The volume of cell suspension required for inoculation following each passaging step may be calculated using the viable cell density determined using an appropriate cell counter (see Note 2). Simultaneously, additional information regarding cell quality may be ascertained using flow cytometry (see Note 4). 13. Gravimetrical measurements may be used to assist in ensuring the concentration of the MC suspension either has remained constant or has been sufficiently concentrated (or diluted)

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during the process step. For example, if the weight of the suspension remains unchanged after removing and replacing the supernatant with fresh cell culture media, DPBS or TrypLE Select 1, and no MCs were lost during aspiration, one may assume that the microcarrier suspension has maintained its initial concentration. 14. The Ns1u criterion describes the impeller speed required to locate the particles at the bottom of the bioreactor, while ensuring that none are at rest. This criterion not only guarantees sufficient mass transfer, but also reduces the shear stress which cells may be subjected to during cultivation [38, 52]. Studies have shown the criterion is met for ProNectin® F MCs at agitation speeds of 49 rpm in 125 mL spinner flasks (Corning®) and between 70 and 80 rpm in the BioBLU® 0.3c (Eppendorf) with marine impeller and down-pumping set-up [47]. 15. The use of an appropriate cryogenic medium for the long-term storage of the cells at 160  C is beneficial to maintaining cell quality during and following the thawing process. To this end, cells were resuspended in Bambanker™ (Nippon Genetics Europe GmbH) to achieve a concentration of 1  106 cells mL1 prior to freezing. 16. To reduce and maintain a pH of 7.2 within the instrumented BioBLU® 0.3c bioreactor, CO2 was added to the overlay of the inlet gas. 17. The process parameters used for the cultivation in the instrumented stirred bioreactor system are summarized in Table 3. 18. The BioBLU® 0.3c is sold with a sampling valve which allows for the sterile sampling of the bioreactor, while it is coupled to the control unit. However, this port was developed for sampling single-cell suspension cultures and is therefore only partially compatible with MC processes. The strong aggregation of the MCs towards the end of the cultivation could potentially result in clogging of the sample valve, adversely impacting sample homogeneity. Therefore, sampling of the bioreactor over the spare probe port, while within a biosafety cabinet is highly recommended. Table 3 Parameter settings for temperature, pH, DO, and CO2 control loops Parameters

Setpoint 

Deadband

P value

I value

Temperature

37 C

0.00

10.0

240.0 s

pH

7.20

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19. Valuable insight into the distribution of the cells on the MCs, cell confluency on single MCs and MC aggregate size can be gleaned by fixing the MC bound cells with 10% neutral buffered formalin solution (Sigma-Aldrich), staining the cells with DAPI, and analyzing the MCs using the EVOS® FL Auto Imaging System (Thermo Fisher Scientific). The staining procedure results in a shift in the excitation (λex) and emission (λem) wavelength from 340 to 364 nm and 488 to 454 nm, respectively, when DAPI selectively binds to double-stranded DNA. This causes the cell nuclei to fluoresce. Thus, when images taken using both the phase contrast and DAPI filter lens at 4 magnification are merged, a single image showing both the orientation of the MCs, as well as the location of individual cell nuclei emerges. These images may then be used to draw conclusions regarding cell distribution and MC aggregation. 20. DAPI working solution is prepared by dissolving DAPI (Roche) in WFI to a final concertation of 1.43 mmol L1. This solution is light and temperature sensitive and should therefore be stored in an opaque container at either 2 to 8  C for up to 6 months or 15 to 25  C for up to 12 months. 21. Prepare a 0.3% Triton® X-100 working solution by diluting 30 μL of Triton® X-100 (Sigma-Aldrich) in 10 mL of sterile DPBS. The working solution may be further improved through sterile filtration (using a 0.22-μm syringe filter) which removes any solid impurities introduced by the detergent, as these may impact the quality of the images taken poststaining. 22. Staining buffer is a mixture consisting of 19 mL autoMACS® Rinsing Solution (Miltenyi Biotec) and 1 mL MACS BSA Stock Solution (Miltenyi Biotec). The buffer contains sodium azide as a preservative, serum to minimize nonspecific binding of antibodies to the cells and ethylenediaminetetraacetic acid to reduce cell aggregation.

Acknowledgments The authors would like to thank Joel R€ath and Riccardo Pianezza, as well as Dr. Tiziano Tallone for their assistance and support with the cell cultivation experiments. Finally, some figures were rendered using the intuitive online application BioRender (see www.bio render.com for more details).

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References 1. Pittenger MF, Mackay AM, Beck SC et al (1999) Multilineage potential of adult human mesenchymal stem cells. Science 284:143–147. https://doi.org/10.1126/sci ence.284.5411.143 2. Lange C, Cakiroglu F, Spiess A-N et al (2007) Accelerated and safe expansion of human mesenchymal stromal cells in animal serum-free medium for transplantation and regenerative medicine. J Cell Physiol 213:18–26. https:// doi.org/10.1002/jcp.21081 3. Han C, Sun X, Liu L et al (2016) Exosomes and their therapeutic potentials of stem cells. Stem Cells Int 2016:1–11. https://doi.org/ 10.1155/2016/7653489 4. Zhou Y, Yamamoto Y, Xiao Z, Ochiya T (2019) The immunomodulatory functions of mesenchymal stromal/stem cells mediated via paracrine activity. J Clin Med 8:1025. https:// doi.org/10.3390/jcm8071025 5. Mohammadalipour A, Dumbali SP, Wenzel PL (2020) Mitochondrial transfer and regulators of mesenchymal stromal cell function and therapeutic efficacy. Front Cell Dev Biol 8:603292. https://doi.org/10.3389/fcell.2020.603292 6. Jossen V (2020) Bioengineering aspects of microcarrier-based hMSC expansions in different single-use bioreactors. Doctoral Thesis, Technische Universit€at Berlin 7. Guldager N Realizing the cost advantages of single-use technology. In: TechTalk https:// www.nne.com/techtalk/Realizing-the-costadvantages-of-single-use-technology/. Accessed 30 May 2021 8. Eibl D, Eibl R (2019) Single-use equipment in biopharmaceutical manufacture: a brief introduction, 2nd edn. Wiley, Hoboken, NJ 9. Bestwick D, Colton R Extractables and leachables from single-use disposables. In: BioProcess Int https://bioprocessintl.com/2009/ extractables-and-leachables-from-single-usedisposables-184117/. Accessed 30 May 2021 10. Soban´tka A, Weiner C (2019) Extractables/ Leachables from single-use equipment: considerations from a (bio)pharmaceutical manufacturer. In: Single-use technology in biopharmaceutical manufacture, 2nd edn. Wiley, Hoboken, NJ, pp 143–158 11. Ding W, Madsen G, Mahajan E et al (2014) Standardized extractables testing protocol for single-use systems in biomanufacturing. Pharm Eng 34:74–85 12. Eibl R, Steiger N, Fritz C et al (2014) Recommendation for leachable studies: standardized cell culture test for the early identification of

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Methods in Molecular Biology (2022) 2436: 113–125 DOI 10.1007/7651_2021_425 © Springer Science+Business Media, LLC 2021 Published online: 15 September 2021

Large-Scale Expansion of Umbilical Cord Mesenchymal Stem Cells with Microcarrier Tablets in Bioreactor Huanye Xu, Zhongxiao Cong, Yuanyuan Zhang, Wei Liu, Xiaojun Yan, and Yanan Du Abstract Mesenchymal stem cells show great potential in tissue engineering and cell-based therapies. This protocol demonstrates the use of 3D TableTrix® microcarrier tablets for large-scale manufacturing of human umbilical cord mesenchymal stem cells (hUCMSCs) in a 5-L stirred tank bioreactor. 3D TableTrix® microcarrier tablets readily disperse into tens of thousands of porous microcarriers to simplify cell seeding, and 3D FloTrix® vivaSPIN bioreactor could automate, monitor, and control the entire culture process. 3D TableTrix® microcarriers could also be fully dissolved upon adding dissolution reagent to gently harvest the expanded cells at a high recovery rate. With this protocol, more than 109 cells could be produced in a 5-L bioreactor. Key words 3D cell culture, Bioreactor, Dissolvable microcarrier, Large-scale expansion, Mesenchymal stem cells

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Introduction Umbilical cord mesenchymal stem cells (UCMSCs) have high capacity of multilineage differentiation and immunomodulatory effects, which show promising prospects in hematopoietic function recovery, tissue injury repair and regeneration [1]. UCMSCs have been greatly applied in tissue engineering and cell-based therapies for stroke, spinal cord injury, neurodegenerative diseases, etc. [2]. Human UCMSCs (hUCMSCs), which could be easily isolated from umbilical cord, with minimal invasiveness to both neonate or mother, get more attention due to their high multiplication capacity, low immunogenicity, as well as little ethical concerns [3]. Nowadays, the demand for UCMSCs is exceeding supply because the required dose for clinical efficacy is high, usually ranging from 40 to 100 million cells [4]. Traditional in vitro monolayer cell culture methods under static conditions are labor intensive and have limited production capacity and efficiency [5]. To produce huge

Huanye Xu and Zhongxiao Cong contributed equally to this work.

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quantity of UCMSCs, multilayer culture flasks with automatic culture systems are introduced, but this still requires large production footprint for incubators. Thus, three-dimension (3D) dynamic cell culture process based on microcarriers and perfusion of medium in automated bioreactor is developed [6]. Here we illustrate a novel culture system developed by Beijing CytoNiche Biotechnology Co., Ltd. for large-scale hUCMSC expansion, with their proprietary dissolvable 3D TableTrix® microcarrier tablets in a low-speed stirred tank 3D FloTrix® vivaSPIN bioreactor. The use of these microcarriers provides anchorage for MSCs during the suspension culture, and the porous nature of these microcarriers dramatically increases surface-to-volume ratio of the whole culture system [7]. In comparison to traditional microcarriers, 3D TableTrix® is fully dissolvable using the complimentary 3D FloTrix® digest solution, which simplifies cell harvesting from microcarriers as no additional separation of microcarriers from cells is required. This gentle harvesting protocol also ensures cell quality. More than 109 cells can be produced in 4 days using one bioreactor and cells retained their MSC phenotypic markers, i.e., >95% expression of CD73, CD90, and CD105, and < 2% for the myeloid lineage cell marker CD14 [8]; B-cell marker CD19 [9], hematopoietic cell markers CD34, CD45; and HLA-DR [10].

2

Materials

2.1 Cell Culture Reagents

1. General MSC culture reagents (i.e., hUCMSCs complete culture medium, phosphate-buffered solution (PBS)) and consumables (T75 culture flasks) are required. 2. hUCMSCs: 2.5
108 cryopreserved hUCMSCs are needed for this culture protocol. Thaw 10 tubes of 2.5
107/tubes cryopreserved hUCMSCs seed cells (passage 4–5) in a 37  C water bath, dilute to 50 mL by slowly adding warm complete hUCMSC culture medium. Centrifuge at 179
g for 5 min. Discard supernatant and resuspend with 50 mL warm complete hUCMSC culture medium (see Note 1).

2.2 3D TableTrix® Microcarrier Tablets

Microcarrier suspension: 3D TableTrix® microcarrier tablets (F01, Beijing CytoNiche Biotechnology Co., Ltd.) are sterile products and are provided as weight-defined tablets (Fig. 1a). Handle these materials under sterile condition. Disperse 500 tablets (equivalent to 10 g, 4,893  288 microcarriers/mg, offering at least 9000 cm2/g surface area) with 500 mL hUCMSC complete medium (20 mg/mL) and gently agitate by shaking the bottle to fully disperse them into individual microcarriers (Fig. 1b). This should take about a minute. After dispersion, most microcarriers are about 200 μm (D10–D90 size distribution between 125 and

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Fig. 1 (a) A photo of bottles of 3D TableTrix® microcarrier tablets. (b) SEM images a porous microcarrier, scale bar ¼ 100 μm. (c) Images of a tablet dispersing in culture medium at 1 s, 5 s, and after agitation

273 μm, Fig. 1c). These microcarriers are porous with pore size of 20.6  5.7 μm (Fig. 1d–f). Microcarrier suspension may be stored at 4 C for 24 h. Warm to 37 C before use. 2.3 3D FloTrix® Digest Solution

1. Concentrated 3D FloTrix® Digest (R005, Beijing CytoNiche Biotechnology Co., Ltd.) solution: Dissolve 2.5 g lyophilized powder in 1.5 L DMEM basic medium. Sterile filter through 0.22-μm filter, store at 4 C for up to 24 h. Warm to 37 C before use. 2. Working 3D FloTrix® Digest solution: Dissolve 10 mg lyophilized powder in 10 mL DMEM basic medium. Sterile filter through 0.22-μm filter, store at 4 C for up to 24 h. Warm to 37 C before use.

2.4 Cell Quality Assessment-Related Reagents

1. Trypan blue solution: 0.4%, liquid, sterile-filtered, suitable for cell culture. 2. Antibodies: PE-conjugated anti-human CD73, FITCconjugated anti-human CD90, PE-conjugated anti-human CD105, FITC-conjugated anti-human CD14, PE-conjugated anti-human CD19, FITC-conjugated anti-human CD34, APC-conjugated anti-human CD45, and PE-conjugated antihuman HLA-DR CD19.

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Other Reagents

1. Sodium hydroxide solution: Dissolve 20 g sodium hydroxide (NaOH) in 5 L deionized (DI) water to prepare 0.1 M NaOH solution. 2. Calcein-AM staining solution: Prepare Calcein-AM working solution according to manufacturer’s instructions. Generally, dilute 2 μL of 1 mM calcein AM stock solution in 1 mL PBS, vortex the resulting solution to ensure thorough mixing. Keep from light and use within 24 h.

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Methods The process is simply summarized in a flowchart in Fig. 2.

3.1 Setting up 3D FloTrix® vivaSPIN Bioreactor on Day 1

1. Calibrate pH sensor (see Note 2). 2. Dismantle all stainless-steel tubes and two 3-blade impellers. Rinse with ultrapure water thoroughly and sonicate for 60 min. 3. Wash the 5 L culture vessel with mild detergent. Rinse with ultrapure water thoroughly for three times. Fill up the vessel with 5 L 0.1 M NaOH solution and soak all stainless-steel tubes and impellers for 12 h (see Note 3). Discard NaOH solution safely. Rinse the vessel, tubes, and impellers with ultrapure water for another three times.

Fig. 2 Flowchart of the protocol, with light blue boxes indicating preparation work involved and dark blue boxes indicating the cell culture process

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Fig. 3 Ports on the top plate of 3D FloTrix® vivaSPIN bioreactor and their respective functions

4. Assemble pH sensor, dissolved oxygen (DO) sensor, all stainless-steel tubes and impellers to the respective ports on the top plate (Fig. 3), according to manufacturer’s instructions. 5. Fill the culture vessel with 2 L PBS and assemble the top plate to the culture vessel and tighten the bolts to ensure the vessel is airtight. 6. Remove the accessories from accessory bags and plug the silicon tubes 1–8 to their respective ports on the top plate (see Note 4). 7. Preform a pressure test to ensure the vessel is airtight (see Notes 5 and 6). 8. Wrap all luer connectors with aluminum foil and tighten all Robert clamps on the silicon tubes 1–6. Leave tube 8 open for pressure relieve during autoclave (Fig. 3, see Note 7). 9. Autoclave the assembled culture vessel at 121  C for 60 min (see Note 8). Remove from the autoclave and cool to room temperature. Remove all hemostatic forceps if used. 10. Install the rotor motor and connect to the controller as according to manufacturer’s instructions. Insert the temperature probe and wrap the heating mat around the culture vessel. Connect compressed air and CO2 to the controller. Fit the silicon tubes 1 and 2 to pump 1 and 2, respectively (Fig. 4, see Note 9). 11. Unclamp the Robert clamps on tubes 5, 6, and 8. On the controller, set temperature to 37 C, agitation speed to 100 rpm/min and inlet air to 22 ccm.

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Fig. 4 Fully assembled 3D FloTrix® vivaSPIN bioreactor

12. Connect a sterile waste bottle to tube 2 with luer connector (see Note 10), unclamp tube 2, and pump out all PBS in the vessel into the waste bottle at 300 mL/min with pump 2. Clamp and disconnect the waste bottle from luer connector (see Note 10). 13. Fill a sterile feed bottle with 2 L complete medium and connect to tube 1 with luer connector, unclamp tube 1 and feed in at 300 mL/min with pump 1. Clamp and disconnect the feed bottle from luer connector. 14. Remove tube 2 from pump 2 and fit tube 4 on pump 2. Unclamp tube 4 and pump at 100 mL/min to aliquot 10 mL of complete medium into the sampling bottle. Clamp tube 4 and transfer sample by connecting a syringe to the luer connector and aspirating the sample from the sampling bottle. 15. Transfer this sample of complete medium to a clean 50-mL centrifuge tube and incubate in a 37 C orbital shaker for 24 h to check for the sterility of the bioreactor (see Note 11). 16. Saturate the complete medium in the vessel with air for 24 h. 17. After 24 h, if no bacteria growth is observed in the vessel and sterility check sample, proceed with inoculation. 3.2 Inoculation on Day 0

1. On Day 0, calibrate the amount of saturated air as 100% for the DO sensor. 2. After calibration, set the DO setting value to 70%, pH to 7.2, temperature to 37 C and agitation to 35 rpm/min. The

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controller will automatically regulate the air and CO2 input as well as the heat mat to maintain the system at the set points. 3. Connect a sterile waste bottle to tube 2 with luer connector, unclamp the Robert clamp and pump out all medium in the vessel into the waste bottle at 300 mL/min with pump 2. Clamp and disconnect the waste bottle from luer connector. 4. Disperse 500 tablets 3D TableTrix® microcarrier tablets (F01, equivalent to 10 g) with 500 mL hUCMSC complete medium (20 mg/mL) in a sterile feed bottle. 5. Thaw 10 tubes of 2.5
107/tubes cryopreserved hUCMSCs seed cells in a 37  C water bath, dilute to 50 mL by slowly adding warm complete hUCMSC culture medium. Centrifuge at 179
g, 5 min. Discard supernatant and resuspend with 50 mL warm complete medium. Transfer to the feed bottle. 6. Top up with 2.45 L warm complete medium to a final volume of 3 L. 7. Connect the feed bottle via luer connector to tube 1. Unclamp the Robert clamp and transfer this microcarrier and cell suspension into the culture vessel via tube 1 on pump 1 at 300 mL/min. Clamp tube 1 and remove the feed bottle. 3.3 Cell Culture, Medium Replenishment, and Growth Monitoring on Days 1–3

1. At 24 h, fill 2 L of warm complete medium in a new sterile feed bottle, connect the bottle to tube 1 via luer connector. Unclamp and feed into the culture vessel by setting pump 1–300 mL/min. The total culture volume is now 5 L. Clamp tube 1 and remove the feed bottle. 2. At 72 h, connect a sterile waste bottle via luer connector on tube 3. Fit tube 3 to pump 2 in the desired direction. Unclamp the Robert clamp and pump out 3 L of spent medium in the vessel to the waste bottle using pump 2 at 300 mL/min. Clamp tube 3 and remove the waste bottle. 3. Fill a sterile feed bottle with 3 L of warm fresh complete medium and connect to tube 1 via luer connector. Unclamp and feed in the fresh medium using pump 1 at 300 mL/min. Clamp tube 1 and remove the feed bottle. 4. Increase agitation speed to 40 rpm. 5. At 24 h, 48 h, 72 h, aliquot 10 mL of cell-microcarrier suspension into the sampling bottle according to step 14, Subheading 3.1. 6. Transfer sample to a clean 15-mL centrifuge tube for cell enumeration by connecting a syringe to the luer connector and aspirating the sample from the sampling bottle. Discard the syringe after each sampling. 7. Aspirate 50 μL of cell-microcarrier suspension to a 96-well plate for cell staining. Leave the rest for cell enumeration.

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Fig. 5 Calcein-AM staining of live cells on microcarriers at 24 h, 48 h, and 72 h

8. To stain cells, let microcarriers settle for 2 min, aspirate supernatant without disturbing the microcarriers and rinse once with PBS. Aspirate PBS and add 100 μL Calcein AM staining solution. Incubate at room temperature for 30 min, observe and image cells under fluorescent microscope with a blue filter (Fig. 5). 9. To enumerate cells, centrifuge cell-microcarrier sample at 179
g, 2 min, discard supernatant. Add 10 mL working 3D FloTrix® Digest solution and incubate in 37  C water bath. Mix thoroughly by pipetting for 30 s at every 10-min interval to facilitate dissolution of microcarriers (Fig. 6). At 30 min (see Note 12), centrifuge at 179
g, 5 min, discard supernatant and resuspend in 10 mL PBS.

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Fig. 6 Bright field images of microcarrier dissolution process, with microcarriers been fully dissolved by 30 min and all cells were released

Fig. 7 Cell growth curve of hUCMSCs on 3D TableTrix® microcarriers in 3D FloTrix® vivaSPIN bioreactor. Cell number on day 0 is the initial seeding number. Numbers of cells for days 1–3 are calculated based on sampling, and for day 4 is based on the total number of cells harvested. All cell numbers reported are total live cell numbers

10. Take a sample of cell suspension and mix with 0.1% Trypan blue solution at 1:1 ratio. Count cells to calculate the total cell number in the culture vessel and cell viability (see Note 13, Figs. 6 and 7).

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3.4 Cell Harvesting on Day 4

1. At 96 h, prepare 1.5 L concentrated 3D FloTrix® Digest solution and transfer to a sterile feed bottle. Connect this bottle via luer connector to tube 1. 2. Connect a sterile waste bottle via luer connector to tube 3. Fit silicon tubing of tube 3 to pump 2 in the desired direction. Unclamp the Robert clamp and pump out 4 L of spent medium in the vessel to the waste bottle using pump 2 at 300 mL/min. Clamp and remove the waste bottle. 3. Feed in the concentrated 3D FloTrix® Digest solution using pump 1 at 300 mL/min. 4. Increase agitation speed to 50 rpm during microcarrier dissolution process. 5. Connect a sterile harvest bottle via luer connector to tube 2. Fit the silicon tubing of tube 2 in the desired direction on pump 2. When all microcarriers are fully dissolved (usually within 30–40 min, see Note 12), pump the cell suspension in the culture vessel to the harvest bottle at 300 mL/min. A total of 2.5 L cell suspension could be harvested. 6. Centrifuge cell suspension at 179
g, 5 min, discard supernatant and resuspend in 400 mL PBS. 7. Take a sample and mix with 0.1% Trypan blue solution at 1:1 ratio. Count cells to calculate the total cell number in the culture vessel and cell viability (Figs. 7 and 8). 8. Repeat step 6 for another two times but resuspend cell pellets with PBS at 2
106/mL instead. Discard supernatant and resuspend in complete medium or cells cryopreservation solution at desired density.

Fig. 8 Cell viability of hUCMSCs on 3D TableTrix® microcarriers in 3D FloTrix® vivaSPIN bioreactor

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Fig. 9 Immunophenotypic characterization of hUCMSCs. These cells expressed antigens CD73, CD90, and CD105, but not antigens CD14, CD-19, HLA-DR, CD34, and CD45

9. Stain cells harvested (about 106 cells) for expression of cell surface markers (PE-conjugated anti-human CD73, FITCconjugated anti-human CD90, PE-conjugated anti-human CD105, FITC-conjugated anti-human CD14, PE-conjugated anti-human CD19, FITC-conjugated anti-human CD34, APC-conjugated anti-human CD45, and PE-conjugated antihuman HLA-DR CD19) for immunophenotype characterization by fluorescence-activated cell sorting (FACS) according to [11] (Fig. 9).

4

Notes 1. Cell viability should be at least 95%. Mix a sample of cell suspension with Trypan blue solution at 1:1 ratio and count cells. Cells should be counted within 3 min. Non-viable cells are stained blue. 2. pH sensor should be fully immersed in calibration solutions (pH 4.00 and pH 7.01, respectively) for several times. 3. NaOH solution has strong causticity. Please exercise safety measures at all times. 4. Tube 1 for material feed, tube 2 for complete liquid discharge and harvest, tube 3 for medium exchange (this tube is fitted with a filter device to prevent microcarriers from been discharged), tube 4 for sampling, tube 5 for air ring sparger, tube 6 for direct air vent, tube 7a and 7b for pressure testing (to be connected with one short silicon tubing after pressure testing), and tube 8 for air vent condenser.

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5. Seal all silicon tubes with hemostatic forceps. Remove the short silicon tubing from tube 5b and 5c. Connect an air compressor to tube 5b, and a pressure gauge to tube 5c. Pump in filtered compressed air until the pressure gage reaches between 0.2 and 0.5 bar. Clamp tube 5b with hemostatic forceps and maintain air pressure for 15 min. If the vessel is not airtight, the pressure will drop. Check for air leakage, and tighten any loose bolts. If vessel is airtight, decompress to room pressure by removing the air pressure gauge from tube 5c. Disconnect air compressor from 5b. Connect 5b and 5c together with one short silicon tubing to form a closed loop. 6. Always stay by the equipment and watch the pressure gauge closely when performing pressure check. Be careful when introducing compressed air and do not let the air pressure in the vessel raise above 1 bar, in case the vessel crack and explode. 7. Robert clamps may open during autoclave due to high temperature. Clamp silicon tubes using hemostatic forceps in conjunction to Robert clamps as an additional measure. 8. After autoclave, check all bolts on the top plate to make sure they are tightly secured and check the integrity and dryness of all air venting filters. If any filter is wet or not intact, change and autoclave again. 9. All pumps rotate clockwise. Make sure the tubing directing is fitted for liquid flow in the intended direction. 10. Spray luer connectors with 75% ethanol before connection and after breaking connection. 11. The objective of this sterility check is to check if the bioreactor is fully aseptic. Take a sample of the prepared complete DMEM before feeding into the bioreactor as a control. 12. All microcarriers should be fully dissolved; extend the dissolution process to not more than 60 min if there is still microcarrier particles visible at 30 min. 13. Cells should be counted within 3 min. Non-viable cells are stained blue and cell viability is counted as percentage of viable cells in the total cell count, cell viability% ¼ living cells/total cells
100%.

Acknowledgments This work is financially supported by Beijing Municipal Science & Technology Commission (Z181100001818005) and R&D program (RC21-01) of Beijing CytoNiche Biotechnology Co. Ltd.

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References 1. Yin S, Ji C, Wu P, Jin C, Qian H (2019) Human umbilical cord mesenchymal stem cells and exosomes: bioactive ways of tissue injury repair. Am J Transl Res 11(3):1230 2. Li T, Xia M, Gao Y, Chen Y, Xu Y (2015) Human umbilical cord mesenchymal stem cells: an overview of their potential in cellbased therapy. Expert Opin Biol Ther 15 (9):1293–1306. https://doi.org/10.1517/ 14712598.2015.1051528 3. Yaghoubi Y, Movassaghpour A, Zamani M, Talebi M, Mehdizadeh A, Yousefi M (2019) Human umbilical cord mesenchymal stem cells derived-exosomes in diseases treatment. Life Sci 233:116733. https://doi.org/10. 1016/j.lfs.2019.116733 4. Lee WY, Wang B (2017) Cartilage repair by mesenchymal stem cells: clinical trial update and perspectives. J Orthop Translat 9:76–88. https://doi.org/10.1016/j.jot.2017.03.005 5. Lam AT, Lee AP, Jayaraman P, Tan KY, Raghothaman D, Lim HL, Cheng H, Zhou L, Tan AH, Reuveny S, Oh S (2021) Multiomics analyses of cytokines, genes, miRNA, and regulatory networks in human mesenchymal stem cells expanded in stirred microcarrier-spinner cultures. Stem Cell Res 53:102272. https://doi.org/10.1016/j.scr. 2021.102272 6. Mabvuure N, Hindocha S, Khan WS (2012) The role of bioreactors in cartilage tissue

engineering. Curr Stem Cell Res Ther 7 (4):287–292 7. Yan X, Zhang K, Yang Y, Deng D, Lyu C, Xu H, Liu W, Du Y (2020) Dispersible and dissolvable porous microcarrier tablets enable efficient large-scale human mesenchymal stem cell expansion. Tissue Eng Part C Methods 26 (5):263–275. https://doi.org/10.1089/ten. TEC.2020.0039 8. Zamani F, Zare Shahneh F, Aghebati-Maleki L, Baradaran B (2013) Induction of CD14 expression and differentiation to monocytes or mature macrophages in Promyelocytic cell lines: new approach. Adv Pharm Bull 3 (2):329–332. https://doi.org/10.5681/apb. 2013.053 9. Wang K, Wei G, Liu D (2012) CD19: a biomarker for B cell development, lymphoma diagnosis and therapy. Exp Hematol Oncol 1 (1):1–7 10. Ding DC, Chang YH, Shyu WC, Lin SZ (2015) Human umbilical cord mesenchymal stem cells: a new era for stem cell therapy. Cell Transplant 24(3):339–347. https://doi.org/ 10.3727/096368915X686841 11. Kassis I, Zangi L, Rivkin R, Levdansky L, Samuel S, Marx G, Gorodetsky R (2006) Isolation of mesenchymal stem cells from G-CSFmobilized human peripheral blood using fibrin microbeads. Bone Marrow Transplant 37 (10):967–976. https://doi.org/10.1038/sj. bmt.1705358

Methods in Molecular Biology (2022) 2436: 127–134 DOI 10.1007/7651_2021_410 © Springer Science+Business Media, LLC 2021 Published online: 04 June 2021

Optimized Method to Improve Cell Activity in 3D Scaffolds Under a Dual Real-Time Dynamic Bioreactor System Flavia Pedrini, Moema A. Hausen, and Eliana A. R. Duek Abstract Bioreactor systems that allow the simulation of in vivo variables in a controlled in vitro environment, were a great advance in the field of tissue engineering. Due to the dynamic-mechanical features that some tissues present, 3D-engineered constructs often do not exhibit the biomechanical properties of these native tissues. Thus, a successful approach must not only achieve tissue repair but also restore its function after injury. Here, we describe a method to improve cell activity in 3D scaffolds in a dynamic bioreactor system through the application of mechanical compression and fluid flow for tissue engineering approaches. Key words 3D Scaffolds, Bioreactor, Cell culture, Fluid flow, Mechanical compression, Tissue Engineering

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Introduction Tissue engineering emerged as an alternative approach that encompasses the architecture of bioartificial tissues in vitro through the implantation of cells on 3D scaffolds [1]. A key factor in the generation of these 3D constructs is the application of mechanical stimuli during maturation to regulate the nascent tissue to a functional activity similar to the quiescent one. In this context, bioreactor-based systems have gained particular interest as they produce clinically effective tissue-based constructs [2, 3]. An important finding was that, when mechanical compression is applied under static conditions, its effects are harmful to cell growth, while dynamic compression promotes cell activity [4]. Thus, a bioreactor should be able to meet the following requirements: (a) intensify mass transfer through perfusion strategies that generate a dynamic environment that promotes cell proliferation and differentiation, and (b) subject the tissue to physiologically relevant loads that can accelerate the production of extracellular matrix in vitro [5]. Studies indicate that when fluid dynamics are applied, cells experience stimuli to which they respond more actively (Fig. 1) [6–9]. In addition to nutrient diffusion, fluid flow is also a way to induce shear stress and give rise to 3D

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Fig. 1 Laser scanning confocal microscopy images of aggrecan labeling under static conditions or in bioreactor under fluid flow. Indirect labeling for aggrecan is identified in red by Alexa Fluor 647. Bars 50 μm. (Adapted with permission from Springer: Biotechnology Letters, Ref. 6, Enhancement of cartilage extracellular matrix synthesis in Poly(PCL-TMC)urethane scaffolds: a study of oriented dynamic flow in bioreactor, Pedrini et al., 2020)

constructions with greater resistance than those developed without any mechanical stimulation [10–13]. So, these stimuli can act as antagonists for more expensive approaches, which often use growth factors, in a more specific and targeted way. A more direct method of mechanical stimulation makes use of models of compression bioreactors, which can vary from uniaxial to multiaxial and allow for the insertion of different variables which act directly on the scaffolds. Since there are no standardized loading protocols, the results obtained by the several studies previously published, impose a difficult task to compare these protocols. However, the dynamic loading parameters including amplitude, frequency and load duration clearly influence biomechanical and biochemical results in these studies [14–19]. The intermittent compressive load, for example, is rather efficient than a continuous one, since the resting period allows the cells to respond to the mechanical stimuli during this gap [15, 16, 20]. Thus, considering all these aspects, the method here described focuses on the application of two real-time modes of mechanical stimulation: compressive loading and fluid dynamic. Both modes, when applied simultaneously and in the chosen parameters, are able to provide an environment that improves cell activity in 3D scaffolds when compared to standard static culture approaches (Fig. 2).

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Fig. 2 Laser scanning confocal microscopy images of aggrecan labeling under static conditions versus dynamic-mechanical bioreactor, under fluid flow and mechanical compression. Indirect labeling for aggrecan is identified in red by Alexa Fluor 647. Bars 50 μm

2

Materials Cell type and culture techniques should be chosen according to the application. The scaffold material can include natural or synthetic biomaterials. The method described refers to parameters tested in a modified polyurethane for approaches in cartilage tissue engineering.

2.1 Bioreactor System

All components used in the bioreactor system are included in the equipment. The dual real-time compressive and fluid flow modes were mounted as described in this section. 1. ElectroForce® BioDynamic 5200® (Bose/TA Instruments) bioreactor, connected to a peristaltic pump and assembled to 22N load cells. The bioreactor possesses four force modules, each one connected to a chamber (Fig. 3) (see Note 1). 2. 180 mL of Dulbecco’s Modified Eagle’s (DMEM) supplemented with fetal bovine serum 10% (FBS) and antibiotics (penicillin 10.000 units, streptomycin 10 mg, and amphotericin B 25 μg per mL), for each bioreactor chamber (see Note 2). 3. 3-Stop PVC Pump tubings hermetically mounted on the chambers (see Note 3). 4. Three way stopcocks connected to the pump tubings (see Note 4). 5. 0.22 μM syringe filters. 6. A cell culture incubator large enough to have the entire bioreactor system housed in it, connected to a CO2 gas supply.

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Fig. 3 Bioreactor system (ElectroForce® BioDynamic 5210®, Bose/TA Instruments). Samples are placed between the upper and lower columns (a) and the system is connected by a peristaltic pump. Regardless of flow direction, the medium is oriented to be directly pumped throughout the scaffolds. Cell loads force sensors are directly connected to each chamber (b). (Adapted with permission from Springer: Biotechnology Letters, Ref. 6, Enhancement of cartilage extracellular matrix synthesis in poly(PCL-TMC)urethane scaffolds: a study of oriented dynamic flow in bioreactor, Pedrini et al., 2020)

3

Methods All the procedures of sample preparation and bioreactor culture must follow Good Laboratory Practice (GLP) within the biosafety rules for the type of cell to be used, according to the application of interest (usually SL-1 or SL-2 cell types). The method described can be applied to several types of scaffold materials; therefore, according to their mechanical properties, the parameters of force and flow must be previously tested.

3.1 Sample Preparation

1. Sterilize the scaffolds prior to seed them with cells. The scaffold sterilization method must respect its physicochemical properties, so the most common ones are autoclaving or UV-C bath (see Note 5). 2. Seed the scaffolds with the desired cell type according to the application of interest. 3. Keep the scaffolds under preculture in static conditions during 4 days, following standard procedures [21] (see Note 6).

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3.2 Sample Transfer and Chamber Assembly

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After static preculture, transfer the samples to the bioreactor chambers under sterile laminar flow environments. Extreme operational care should be taken when placing 3D scaffolds to avoid cell scratching. 1. Remove one side of the acrylic seal plate of the bioreactor chamber. Separate each column enough to insert the sample. Mount the flow loop tubing system at input and output valves (see Note 7). 2. Add 180 mL of DMEM medium. Remove all air bubbles inside the tubing system (see Note 8). 3. Place the scaffold between the columns with the aid of tweezers, hold the scaffold and approach the upper column close enough to couple it. Do not tighten. 4. Add the seal acrylic plate to the chamber and thread it tightly. Observe the chamber seal o-ring correct positioning. 5. Vertically place the chamber on its specific holder. Align the upper column to the bottom one carefully to avoid sample displacement. 6. Add an 0.22 μM syringe filter to the upper aperture that contacts the remaining air volume in the chamber. 7. Place the assembled chamber into the bioreactor system. The specific mechanic load module and flow rate must be set up according to the application of interest.

3.3 Mechanical Compression Load and Fluid Flow Parameters

One must have in mind that compression and fluid flow parameters must comply with the material’s mechanical properties. Thus, cellfree previous tests are essential to determine the assays specifications. 1. Establish a route of the fluid flow through the bioreactor chamber. To evaluate the flow directly throughout the scaffold, the tubing system must be assembled in the upper and lower columns (see Note 9). 2. Setup the peristaltic pump to a continuous flow rate of 0.4 mL/min (see Note 10). 3. Mount the load cell in the bioreactor according to the applied force range desired as indicated in the equipment reference manual. 4. Setup the loading parameters (see Note 11): Force value: 1 N; Frequency: 0.015 Hz; Duration: 1 h continuous; Maximum duration: 18 h (see Note 12).

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Notes 1. It must be considered that this bioreactor can be configured to operate in different modes of action, including compression, tension and fluid flow. For each mode, the components used in the assembly of the equipment’s chambers are different. In the described method, the association of the compression modes and fluid flow was chosen. The total number of mounted chambers are suggested when sample comparison is a desired parameter to be evaluated. 2. Medium characteristics may vary due to cell type. 3. The hermetic feature should be previously tested before autoclaving through a manual sphygmomanometer. 4. The open/close apertures setup must allow the fluid flow oriented as desired, that is, from top to bottom. 5. 3D porous scaffolds of materials sensitive to the autoclave temperature can be disinfected with alcohol baths. To facilitate the alcohol flow within the scaffold, the material can be immersed in alcohol and subjected to vacuum. Successive washes in saline solution, also subjected to the same vacuum, will be necessary for the removal of alcohol. 6. Cell attachment and matrix synthesis during the preculture period are important for a more effective mechanotransduction and resultant tissue stiffening when under the dynamicmechanical environment [20]. One must also observe that differentiated cell types must have distinct conduct as compared to undifferentiated ones. Mature cells that exhibit high matrix expression should be treated with additional care to properly adapt the medium content. 7. According to the desired flow direction, one must choose different inputs and outputs apertures, and previous in/out flow tests should be performed. 8. If an expressive amount of medium is lost during air removal, the medium must be completed to a final volume of 180 mL. The o-ring must be completed dry with no medium residue to avoid contamination. 9. For convenience, create a schematic map of the designed flow orientation. 10. Cell adhesion can be affected according to the cell type and material properties, such as roughness, porosity, and chemical composition. Thus, in previous test trials, the ideal flow rate for each specific purpose should be assessed. 11. The main difference in this protocol is that the system is set up for a sinusoidal displacement approach until it detects a specific

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determined force, instead of amplitude parameters usually attributed to this type of assay [20], since reproducibility can be compromised due slight changes in sample dimensions. By this way, submitting each specimen to a specific force of 1 N will guarantee an accurate homogeneous comparison between them. 12. The 1 N was applied 3 times per day, each time during 1 h, with each new application preceded by a 1 h constraint-free rest.

Acknowledgments The authors thanks to UFSCar-Sorocaba/PPGBMA for support and to CAPES grant 1724839. References 1. Kingsley G (2001) Tissue engineering. In: Encyclopedia of Physical Science and Technology. Academic Press, Cambridge, Massachusetts, pp 133–143 2. Paez-Mayorga J, Herna´ndez-Vargas G, RuizEsparza GU, Iqbal HMN, Wang X, Zhang YS, Parra-Saldivar R, Khademhosseini A (2019) Bioreactors for cardiac tissue engineering. Adv Healthc Mater 8(7):e1701504. https://doi. org/10.1002/adhm.201701504. Epub 2018 May 8 3. Schu¨rlein S, Al Hijailan R, Weigel T, Kadari A, Ru¨cker C, Edenhofer F, Walles H, Hansmann J (2017) Generation of a human cardiac patch based on a Reendothelialized biological scaffold (BioVaSc). Adv Biosys 1(3):1600005 4. Grodzinsky AJ, Levenston ME, Jin M, Frank EH (2000) Cartilage tissue remodeling in response to mechanical forces. Annu Rev Biomed Eng 2:691–713. https://doi.org/10. 1146/annurev.bioeng.2.1.691 5. Cartmell S, El Haj A (2005) Mechanical bioreactors for bone tissue engineering. In: Chaudhuri J, Al-Rubeai M (eds) Bioreactors for tissue engineering. Dordrecht, Springer. https://doi.org/10.1007/1-4020-3741-4_8 6. Pedrini F, Hausen M, Gomes R, Duek E (2020) Enhancement of cartilage extracellular matrix synthesis in poly(PCL-TMC)urethane scaffolds: a study of oriented dynamic flow in bioreactor. Biotechnol Lett 42 (12):2721–2734. https://doi.org/10.1007/ s10529-020-02983-1. Epub 2020 Aug 12 7. Han TTY, Flynn LE (2020) Perfusion bioreactor culture of human adipose-derived stromal cells on decellularized adipose tissue scaffolds

enhances in vivo adipose tissue regeneration. J Tissue Eng Regen Med 14(12):1827–1840. https://doi.org/10.1002/term.3133 8. Bhaskar B, Owen R, Bahmaee H et al (2018) Design and assessment of a dynamic perfusion bioreactor for large bone tissue engineering scaffolds. Appl Biochem Biotechnol 185:555–563. https://doi.org/10.1007/ s12010-017-2671-5 9. Schmid J, Schwarz S, Meier-Staude R et al (2018) A perfusion bioreactor system for cell seeding and oxygen-controlled cultivation of three-dimensional cell cultures. Tissue Eng Part C Methods 24(10):585–595. https:// doi.org/10.1089/ten.TEC.2018.0204 10. Li D, Tang T, Lu J, Dai K (2009) Effects of flow shear stress and mass transport on the construction of a large-scale tissue-engineered bone in a perfusion bioreactor. Tissue Eng Part A 15(10):2773–2783. https://doi.org/10. 1089/ten.TEA.2008.0540 11. Yeatts AB, Fisher JP (2011) Bone tissue engineering bioreactors: dynamic culture and the influence of shear stress. Bone 48(2):171–181. https://doi.org/10.1016/j.bone.2010.09. 138. Epub 2010 Oct 13 12. Rotenberg MY, Ruvinov E, Armoza A, Cohen S (2012) A multi-shear perfusion bioreactor for investigating shear stress effects in endothelial cell constructs. Lab Chip 12 (15):2696–2703. https://doi.org/10.1039/ c2lc40144d. Epub 2012 May 23 13. McCoy RJ, O’Brien FJ (2010) Influence of shear stress in perfusion bioreactor cultures for the development of three-dimensional bone tissue constructs: a review. Tissue Eng

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Part B Rev 16(6):587–601. https://doi.org/ 10.1089/ten.TEB.2010.0370. Epub 2010 Oct 12 14. Shelton JC, Bader DL, Lee DA (2003) Mechanical conditioning influences the metabolic response of cell-seeded constructs. Cells Tissues Organs 175(3):140–150. https://doi. org/10.1159/000074630 15. Chowdhury TT, Bader DL, Shelton JC, Lee DA (2003) Temporal regulation of chondrocyte metabolism in agarose constructs subjected to dynamic compression. Arch Biochem Biophys 417(1):105–111. https://doi.org/ 10.1016/s0003-9861(03)00340-0 16. Kisiday JD, Jin M, DiMicco MA, Kurz B, Grodzinsky AJ (2004) Effects of dynamic compressive loading on chondrocyte biosynthesis in self-assembling peptide scaffolds. J Biomech 37(5):595–604. https://doi.org/10.1016/j. jbiomech.2003.10.005 17. Kaupp JA, Weber JF, Waldman SD (2012) Mechanical stimulation of chondrocyteagarose hydrogels. J Vis Exp 27(68):e4229. https://doi.org/10.3791/4229 18. Wu P, DeLassus E, Patra D, Liao W, Sandell LJ (2013) Effects of serum and compressive loading on the cartilage matrix synthesis and spatiotemporal deposition around chondrocytes in

3D culture. Tissue Eng Part A 19 (9-10):1199–1208. https://doi.org/10. 1089/ten.tea.2012.0559. Epub 2013 Feb 14 19. Nebelung S, Gavenis K, Rath B, Tingart M, Ladenburger A, Stoffel M, Zhou B, MuellerRath R (2011) Continuous cyclic compressive loading modulates biological and mechanical properties of collagen hydrogels seeded with human chondrocytes. Biorheology 48 (5):247–261. https://doi.org/10.3233/BIR2012-0597 20. Anderson DE, Johnstone B (2017) Dynamic mechanical compression of chondrocytes for tissue engineering: a critical review. Front Bioeng Biotechnol 5:76. https://doi.org/10. 3389/fbioe.2017.00076 21. Pamies D, Bal-Price A, Simeonov A, Tagle D, Allen D, Gerhold D, Yin D, Pistollato F, Inutsuka T, Sullivan K, Stacey G, Salem H, Leist M, Daneshian M, Vemuri MC, McFarland R, Coecke S, Fitzpatrick SC, Lakshmipathy U, Mack A, Wang WB, Yamazaki D, Sekino Y, Kanda Y, Smirnova L, Hartung T (2017) Good cell culture practice for stem cells and stem-cell-derived models. ALTEX - Alternatives to animal experimentation 34(1):95–132. https://doi.org/10. 14573/altex.1607121

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In Vitro 3D Mechanical Stimulation to Tendon-Derived Stem Cells by Bioreactor Ziming Chen, Peilin Chen, Rui Ruan, and Minghao Zheng Abstract Bioreactors can offer an advanced platform to provide conditions that mimic the native microenvironment, which can also provide stretching environment for mechanobiology research. Tendon-derived stem cells (TDSCs) are a type of mechanosensitive and multipotent cells, which behave differently in diverse mechanical stretching environments. We have proved the in vitro three-dimensional (3D) mechanical stimulation could closely mimic the stretching environment in vivo. Thus, here we describe applying a customized bioreactor to provide 3D force for mechanical stimulation on TDSC in vitro. Key words Bioreactor, Mechanical loading, Mechanobiology, RT-qPCR, Tendon formation

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Introduction Mechanobiology is a multidisciplinary subject including biology, engineering, and physics, which investigates the influence of mechanical stress in life science. Cells have the ability to sense force, and in turn, respond to the extracellular environment [1]. Thus, mechanobiology is an essential discipline with wide application prospect. Various mechanical stimulation types are discussed nowadays, including compression stimulation, tensile strain, ventilation pressure, flow shear stress, and so forth [2–4]. For tendon, a connective tissue that attaches muscle to bone, it experiences numerous repetitive stretching over a long period of time. Thus, here we use stretching tensile strain to mimic its nature environment. Tendon-derived stem cell (TDSC) is a type of mechanosensitive and multipotent cells, which behaves differently in diverse mechanical stretching environments [5]. Thus, it is an idea source for mechanobiology study. A proper loading protocol is essential for tendon differentiation of TDSCs [6]. Two-dimensional (2D) loading models, which are applied to monolayer cells, are the most commonly used for tendon study. However, in our

Ziming Chen and Peilin Chen contributed equally to this work.

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previous study, we demonstrated that the three-dimensional (3D) uniaxial loading protocol could provide a better environment for the cell–matrix interactions, which closely mimic the mechanical microenvironment of TDSCs, and preferentially generate a tenogenic response [7]. In this chapter, we provided 2 options for 3D uniaxial stretching to TDSCs, including the scaffold-based method and scaffold-free method. The scaffold-based method was using CelGro® scaffold, a unique collagen scaffold providing a 3D space for cell attachment [8]. The scaffold-free method was modified by our previous protocol and designed to stimulate TDSCs to generate extracellular matrix and then was formed to a tendon organoid [9, 10]. Our data showed both of these two options can induce tenogenesis of TDSCs. These two methods have their own advantages. The scaffold-free method may be time consuming but mimic natural process. On the other hand, the scaffold-based method is time effective and enables potential development of surgical implantable biological devices. Other types of mechanosensitive cells, such as cardiac myocytes, endothelial cells, and osteocytes, can be investigated by this method as well [11–13]. However, some cells without producing extracellular matrix ability can only be seeded on scaffolds. Moreover, investigators should use specific loading regime and detect specific outcome events on other types of cells. In the present chapter, we use tenogenesis markers to prove our loading regime can enhance tenogenesis for TDSCs than static culture (see Fig. 1).

Fig. 1 Tenogenesis marker in (a) scaffold-free model and (b) scaffold-based model. RNA was extracted from 1 cm from the middle of samples. Individual gene-expression levels were normalized against the internal control, 36B4, and then normalized to gene-expression levels from static cultures

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Materials A bioreactor which can provide uniaxial stretching: Autoclave (1.5 h at 134
C) the bioreactor culture chamber, sample hooks, glass cover, and screws before use. Sterilize the motor module and connection cable by ultraviolet rays (UV) exposure overnight.

2.1 Isolation of Mice TDSCs

1. 6–8-week-old C57BL/6 mice. 2. T-25 cell culture flask. 3. Centrifuge. 4. 70% ethanol. 5. Phosphate buffer saline (PBS). 6. Sterile dissection kit for small animals. 7. Wash buffer: 100 U/mL penicillin and 100 μg/mL streptomycin in PBS. 8. 3 mg/mL Type 1 collagenase. 9. 70 μm cell strainer. 10. Complete medium: Alpha Modified Eagle Medium supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, 100 μg/mL streptomycin. 11. Fluorescence-activated Cell Sorting (FACS) buffer: PBS with 5% FBS. 12. Conjugated antibodies for flow cytometry: CD44, CD45, CD90, and CD34.

2.2 Mechanical Stimulation

1. TDSCs: Passage 2–4.

2.2.1 Scaffold-Free Mechanical Stimulation

3. Stimulation medium: 25 ng/mL connective tissue growth factor and 4.4 mg/mL ascorbic acid in complete medium.

2. T-75 cell culture flask.

4. Complete medium. 5. PBS: prewarmed to 37
C. 6. Sterile tweezer. 7. Trypsin-EDTA solution: 0.25% trypsin and 0.02% EDTA. 8. 100  20 mm Petri dish. 2.2.2 Scaffold-Based Mechanical Stimulation

1. TDSCs: Passage 2–4. 2. Complete medium. 3. 100  20 mm Petri dish. 4. CelGro® collagen scaffold. 5. Customed culture chamber: 1.5 cm  2 cm, autoclave before use.

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2.3 Extraction of RNA for Validation of the System

1. TRIzol reagent. 2. PBS. 3. Clean scissors: autoclave before use. 4. Clean tweezers: autoclave before use. 5. A 10 mL tissue grinder: include a mortar and a pestle; autoclave before use. 6. PureLink™ RNA Mini Kit. 7. Liquid nitrogen. 8. Ethanol: Molecular Biology grade. 9. 70% ethanol in RNase-free water: Molecular Biology grade. 10. Chloroform: Molecular Biology grade. 11. 2 mL RNase-free tubes.

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Methods Carry out all procedures at room temperature unless otherwise specified. All medium should be prewarmed to 37
C before adding to cells.

3.1 Isolation of Mice TDSCs

1. Euthanize the 6–8-week-old C57BL/6 mice by cervical dislocation. Shave the hair and sterilize hind limbs with 70% ethanol spray. 2. Isolate the hind limb at femur head level. Remove the surrounding adipose tissue and blunted isolation of connective tissue to expose patellar tendon and Achilles tendon (see Note 1). 3. Isolate the mid portion of Patellar and Achilles tendon and immediately rinse them by wash buffer. Isolated tendon tissue was homogenized and digested by 3 mg/mL Type 1 collagenase for 3 h (see Note 2). 4. Filter the digested tendon remains by 70 μm cell strainer, and remove residual tendon fibers. 5. After centrifuging, culture the isolated cells with complete medium in T-25 flask. Incubation conditions are 37
C in a humidified atmosphere and containing 5% CO2. 6. Make a single cell suspension at 107 cells/mL in FACS buffer (see Note 3). 7. For each staining sample, put 100 μL of 106 cells into a sorter tube. For each antibody combination, add conjugated antibodies (CD44, CD45, CD90, and CD34) to the samples at final concentration of 10 μg/mL, and incubate for 30 min in dark at

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4
C. Wash cells after incubation with ice-cold PBS for three times. 8. Add secondary antibodies to the samples in 2 μg/mL and incubate for 30 min at 4
C in dark. Wash samples again for three times with PBS and resuspend by ice-cold FACS buffer. The samples were then analyzed by flow cytometry within 1 h. 3.2 Mechanical Stimulation 3.2.1 Scaffold-Free Mechanical Stimulation

1. Add 8 mL complete medium to the warmed-up cells (mice TDSCs, one million cells, passage 2–4, to 37
C). And use a pipette to transfer cells to a 15 mL centrifuge tube. Centrifuge at 350  g for 5 min. 2. Decant the medium, and resuspend cells gently in 1–2 mL of complete medium. Gently pipette up and down for several times to completely resuspend it (see Note 4). 3. Transfer resuspended cells to a T-75 flask. Add complete medium to the flask to reach a total volume of 10 mL. Place the flask into the incubator and culture at 37
C with 5% CO2. Cell culture should be done until the cells are cultured to 100% confluence (see Note 5). 4. Discard the complete medium. Add 10 mL of stimulation medium slowly and avoid touching the cells attached to the bottom of the flask. 5. Culture the cells in stimulation medium for 6 days (see Note 6) to sufficiently generate the cell sheet at 37
C with 5% CO2 (see Note 7). 6. Discard the stimulation medium completely, and wash the monolayer cell sheet with prewarmed PBS by swirling the flask. Then discard the PBS. Use a pipette to add 1 mL of trypsin-EDTA solution to the corner of the flask. 7. Gently tap the corner of the flask to detach the cell sheet until the corner of the cell sheet starts to peel off from the bottom of the flask (see Fig. 2a). 8. Add 9 mL of complete medium to stop the trypsinization. Keep swirling the flask to completely peel off the cell sheet. 9. Pour the whole cell sheet into a Petri dish with medium. Use a sterile tweezer to pick up one corner of the cell sheet and rotate in a clockwise direction for 15 times. Then pick up another end of the cell sheet and rotate in an anti-clockwise direction for ten times to firmly generate a tendon-like 3D construct (see Fig. 2a). 10. Connect the hooks by the connecter and adjust the distance between two hooks to 2 cm. Gently wind the 3D TDSCs construct on the assembled hook for three times on each hook.

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Fig. 2 Schematic diagram of (a) scaffold-free model and (b) scaffold-based model

11. Anchor securely the hooks with cell construct onto the chamber of the bioreactor by tightening the screws on two ends. Fill up the chamber with complete medium. Cut the hooks connecter by sterile scissors. Put the glass lid on the chamber. Culture overnight at 37
C with 5% CO2 (see Note 8). 12. Connect the wires for the actuator. Switch on the power and corresponding channel controller to start mechanical stimulation. Check the indicator lights and assure the bioreactor can function properly. Move the bioreactor into the incubator and subject the 3D cell construct to mechanical stretching for 6 days (stretch program: 6% stretching, 0.25 Hz, 8 h, followed by 16 h rest). 3.2.2 Scaffold-Based Mechanical Stimulation

1. Take a 30 mm  40 mm CelGro® scaffold and make the rough side of CelGro® up. Put it on the bottom part of the culture chamber. Firmly assemble the culture chamber (see Fig. 2b) (see Note 9). 2. Thaw up the cells (mice TDSCs, one million cells, passage 2–4, to 37
C) as descripted in Subheading 3.2.1, steps 1 and 2. 3. Seed cells (Quantity: 4  105) into CelGro® culture chamber. Put the culture chamber in a Petri dish and culture 24 h at 37
C with 5% CO2 for cell attachment in CelGro®. 4. Roll the 3D CelGro® collagen-tenocyte construct up to 25  5 mm (see Fig. 2b). 5. Use the sterile clamp to anchor the collagen-tenocyte construct onto the chamber of the bioreactor by tightening the screws on two ends. Fill up the chamber with complete medium. Put the glass lid on the chamber. Culture overnight at 37
C with 5% CO2.

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6. Assemble and switch the bioreactor on as described in Subheading 3.2.1, step 12. 3.3 Extraction of RNA for Validation of the System

1. Detach the sample from bioreactor by two clean tweezers, and then wash the sample for three times by PBS (see Note 10). 2. Dissect a sample of 1 cm in the middle by a clean scissor (see Note 11). 3. Snap-freeze the sample in liquid nitrogen. 4. Quickly move the sample into a clean homogenization.

mortar

for

5. Add 1 mL TRIzol reagent into the mortar. 6. Vertically press the sample with a rocking motion to homogenize the sample by a pestle, and then keep grinding the tissue for about 5 min (see Note 12). 7. Transfer the homogenized samples into a 2 mL tube and incubate for 10 min at room temperature. 8. Add 200 μL chloroform into the tube. Shake the tube vigorously for 15 s by hand. And then incubate the sample for 3 min at room temperature. 9. Centrifuge for 15 min at 4
C, 12,000  g. Transfer the colorless upper aqueous phase into a clean 2 mL tube (see Note 13). 10. Add same volume 70% ethanol to the aqueous phase. Vortex to mix well and then spin down. 11. Transfer up to 700 μL of sample to the spin cartridge with the collection tube. And then centrifuge at 12,000  g for 15 s at room temperature. Discard the liquid in collection tube and reinsert the spin cartridge. Repeat this step until all the samples are processed (see Note 14). 12. Add 700 μL Wash Buffer I, and then centrifuge at 12,000  g for 15 s. Insert the spin cartridge into a new collection tube. 13. Add 500 μL Wash Buffer II, and then centrifuge at 12,000  g for 15 s. Discard the liquid in collection tube. Repeat this step once. 14. Centrifuge at 12,000  g for 2 min to dry the membrane. Insert the spin cartridge into a recovery tube. Add 30 μL RNase-free water to the center of the spin cartridge. Incubate for 1 min (see Note 15). 15. Centrifuge the spin cartridge for 2 min at 12,000  g. Now the RNA is in the recovery tube and ready for RT-qPCR (Table 1).

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Table 1 Primer sequences used for RT-qPCR analysis Primer sequence Gene

Forward 50 ! 30

Reverse 50 ! 30

COL1A1

TGACTGGAAGAGCGGAGAGT

GTTCGGGCTGATGTACCAGT

Scleraxis

CCCAAACAGATCTGCACCTT

GGCTCTCCGTGACTCTTCAG

Mohawk

GTCCGGCAGCCAGATTTAAG

TCGCTGAGCTTTCCCCTTTA

Tenomodulin

CCGCAGAAAAGCCTATTGAA

GACCACCCATTGCTCATTCT

36B4

CTTCCCACTTGCTGAAAAGG

CGAAGAGACCGAATCCCATA

4

Notes 1. Ensure complete separation of tendons, and avoid any residual muscles or other tissues. Otherwise, it may affect the purity of the TDSCs finally obtained. 2. Vortex 10 s per 30 min when digest. 3. Keep cells on ice at all times for the following staining procedure. 4. It is important to completely resuspend it. Evenly distributed cell is one of the keys to form a quality cell sheet. 5. Change the medium every 3 days. 6. Change the medium every 3 days. Operation when changing medium must be gentle. The medium flow should not directly touch the cells as it could tear the cell sheet. 7. Extracellular matrix will become thick and present to be cloudy when observed from the bottom of the flask after stimulated by stimulation medium, which means the cell sheet is sufficiently generated. Ideally, 6 days should be enough to observe the margin of the cell sheet is rolled up. Rolled edge is the signal to move to the next step. 8. Culture overnight at 37
C with 5% CO2 before stretching can shrink the edge of the sample and more stably form the 3D construct. 9. Add medium into the chamber firstly to ensure firmly assembled and no leaking for the culture chamber. 10. Washing collagen-tenocyte construct should be gentler than washing scaffold-free tendon-like 3D construct and do not unfold the construct when washing the collagen-tenocyte construct.

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11. Uniaxial stretching provides non-uniform deformation field on the sample [14]. Thus, choosing different parts of the sample for RNA extraction may lead to different results. 12. The aim of grinding is to obtain a uniform powder without large sample pieces which can pose an adverse impact on efficiency of RNA extraction in subsequent steps. 13. Count the volume of aqueous phase when transfer. 14. Use PureLink™ RNA Mini Kit from this step. Spin cartridge and collection tube can be found in the kit. Before use, add ethanol (Molecular Biology grade) to Wash Buffer II as instructed in product manual. 15. Recovery tube can be found in the PureLink™ RNA Mini Kit.

Acknowledgements The authors acknowledge financial support from Australian Research Council Industrial Transformation Training Centre for Personalised Therapeutics Technologies (IC170100016). References 1. Lim CT, Bershadsky A, Sheetz MP (2010) Mechanobiology. J R Soc Interface 3(Suppl 3):S291–S293. https://doi.org/10.1098/rsif. 2010.0150.focus 2. Li Y, Chen M, Hu J, Sheng R, Lin Q, He X, Guo M (2021) Volumetric compression induces intracellular crowding to control intestinal organoid growth via Wnt/β-catenin signaling. Cell Stem Cell 28(1):63–78.e67. https://doi.org/10.1016/j.stem.2020.09. 012 3. Spassov SG, Kessler C, Jost R, Schumann S (2019) Ventilation-like mechanical strain modulates the inflammatory response of BEAS2B epithelial cells. Oxidative Med Cell Longev 2019:2769761. https://doi.org/10.1155/ 2019/2769761 4. Lee J, Estlack Z, Somaweera H, Wang X, Lacerda CMR, Kim J (2018) A microfluidic cardiac flow profile generator for studying the effect of shear stress on valvular endothelial cells. Lab Chip 18(19):2946–2954. https:// doi.org/10.1039/c8lc00545a 5. Bi Y, Ehirchiou D, Kilts TM, Inkson CA, Embree MC, Sonoyama W, Li L, Leet AI, Seo BM, Zhang L, Shi S, Young MF (2007) Identification of tendon stem/progenitor cells and the role of the extracellular matrix in their niche. Nat Med 13(10):1219–1227. https:// doi.org/10.1038/nm1630

6. Wang T, Lin Z, Day RE, Gardiner B, LandaoBassonga E, Rubenson J, Kirk TB, Smith DW, Lloyd DG, Hardisty G, Wang A, Zheng Q, Zheng MH (2013) Programmable mechanical stimulation influences tendon homeostasis in a bioreactor system. Biotechnol Bioeng 110 (5):1495–1507. https://doi.org/10.1002/ bit.24809 7. Wang T, Thien C, Wang C, Ni M, Gao J, Wang A, Jiang Q, Tuan RS, Zheng Q, Zheng MH (2018) 3D uniaxial mechanical stimulation induces tenogenic differentiation of tendon-derived stem cells through a PI3K/ AKT signaling pathway. FASEB J 32 (9):4804–4814. https://doi.org/10.1096/fj. 201701384R 8. Allan B, Ruan R, Landao-Bassonga E, Gillman N, Wang T, Gao J, Ruan Y, Xu Y, Lee C, Goonewardene M, Zheng M (2021) Collagen membrane for guided bone regeneration in dental and orthopedic applications. Tissue Eng A 27(5–6):372–381. https://doi. org/10.1089/ten.TEA.2020.0140 9. Chen Z, Chen P, Ruan R, Chen L, Yuan J, Wood D, Wang T, Zheng MH (2020) Applying a Three-dimensional uniaxial mechanical stimulation bioreactor system to induce tenogenic differentiation of tendon-derived stem cells. J Vis Exp 162. https://doi.org/10. 3791/61278

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10. Chen P, Chen Z, Mitchell C, Gao J, Chen L, Wang A, Leys T, Landao-Bassonga E, Zheng Q, Wang T, Zheng M (2021) Intramuscular injection of Botox causes tendon atrophy by induction of senescence of tendon-derived stem cells. Stem Cell Res Ther 12(1):38. https://doi.org/10.1186/s13287-02002084-w 11. Saucerman JJ, Tan PM, Buchholz KS, McCulloch AD, Omens JH (2019) Mechanical regulation of gene expression in cardiac myocytes and fibroblasts. Nat Rev Cardiol 16 (6):361–378. https://doi.org/10.1038/ s41569-019-0155-8 12. Fang Y, Wu D, Birukov KG (2019) Mechanosensing and mechanoregulation of

endothelial cell functions. Compr Physiol 9 (2):873–904. https://doi.org/10.1002/cphy. c180020 13. Uda Y, Azab E, Sun N, Shi C, Pajevic PD (2017) Osteocyte mechanobiology. Curr Osteoporos Rep 15(4):318–325. https://doi. org/10.1007/s11914-017-0373-0 14. Morita Y, Sato T, Higashiura K, Hirano Y, Matsubara F, Oshima K, Niwa K, Toku Y, Song G, Luo Q, Ju Y (2019) The optimal mechanical condition in stem cell-to-tenocyte differentiation determined with the homogeneous strain distributions and the cellular orientation control. Biol open 8(5):bio039164. https://doi.org/10.1242/bio.039164

Methods in Molecular Biology (2022) 2436: 145–156 DOI 10.1007/7651_2021_411 © Springer Science+Business Media, LLC 2021 Published online: 22 June 2021

Microcarrier-Supported Culture of Chondrocytes in Continuously Rocked Disposable Bioreactor Kamil Wierzchowski and Maciej Pilarek Abstract Disposable wave-assisted bioreactors are devices originally designed for scaling-up cultures of extremely fragile animal cells. In such bioreactors, agitation is achieved by continuous horizontal oscillations of disposable culture bag-like container fixed in a rocker unit. The continuous rocking movement of the container induces waves in the two-phase (i.e., gas–liquid) culture system composed of CO2-enriched air and aqueous culture medium. Such continuously oscillating devices can be utilized for supporting homogeneity in systems for in vitro propagation of animal anchorage-dependent, that is, adherent, cells, like CP5 chondrocytes cells. As most of in vitro cultured cells exhibit anchorage-dependency toward solid surface, the suitable interface can be provided by beads of microcarriers made of polymers and characterized by large surface-to-volume ratio. This chapter describes a methodology for efficient propagation of CP5 chondrocytes on Cytodex 3 microcarriers performed in ReadyToProcess WAVE 25 disposable bioreactor, as well as all useful procedures for daily monitoring the growth of CP5 chondrocytes. Key words Anchorage-dependent cells, Bioprocess intensification, Chondrocytes, Disposable bioreactor, Microcarrier beads, Wave-assisted agitation

1

Introduction In the case of scaling-up bioprocesses focused on anchoragedependent (i.e., adherent) animal cells, the static culture systems must be substituted by gentle-agitated systems to ensure sufficient level of aeration in high cell density cultures. Instead of commonly applied rotating stirrers applied in classical bioreactor systems, continuously oscillating devices can be utilized for supporting homogeneity in disposable (i.e., single-use) bioreactor systems originally designed for in vitro maintaining of extremely fragile animal cells [1]. Without a doubt, the disposable bioreactors support notable reduction of microbial contamination, as well as reduce the developing cost of the plant constructions, in contrast to typical bioreactors equipped with large-volume vessels made of glass or stainless steel [2, 3]. In the case of single-use bioreactors the agitation is achieved mainly by continuous horizontal oscillations of disposable culture bag fixed in a rocker unit [4, 5]. The continuous rocking of the rocker generates, as well as escalates, waves in the

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two-phase (i.e., gas–liquid) culture system composed of CO2enriched air and aqueous culture medium, closed inside the disposable bag-like container. Continuously generated waves cause agitation of large volumes of medium and facilitate to disperse gases, nutrients, extracellularly secreted bioproducts in the liquid phase [6–8], what finally enhances the homogeneity of whole waving culture system. To enhance the proliferation yield of adherent animal cells exhibiting anchorage-dependency toward solid surface, that is, majority of mammalian, avian, fish, and insect cells, an easy access of biomass to solid-state surface must be provided [9]. Microcarriers, in the form of μm-/mm-scale beads made of biocompatible polymer, characterized by large surface-to-volume ratio, are recognized as the most efficient proliferation supporters in cultures of adherent cells [10, 11]. Moreover, depending on the type of applied microcarriers, adherent cells may grow in monolayered form on the outer surface of beads or migrate inside micropores of porous beads suspended in the culture medium [12, 13]. The chondrocytes and cartilage tissue are responsible for the skeleton stabilization and connecting elements of the skeletal system. Due to slow in vivo regeneration of joint cartilage, the most effective way to rebuild degraded cartilage is chondrocyte reimplantation into damaged joints [14, 15]. Currently, new methods for efficient and low-cost in vitro cultures of chondrocytes are seeking with a wide scope of possible applications of bioprocessed chondrocytes in medicine [15, 16]. In this chapter, a detailed methodology for efficient proliferation of CP5 chondrocytes on Cytodex 3 microcarrier beads, performed in ReadyToProcess WAVE 25 disposable bioreactor has been presented. Some procedures useful for daily monitoring the proliferation of CP5 chondrocytes, by measurement of density, viability of cells and their metabolic activity, the specific glucose consumption rate, the level of lactate dehydrogenase activity in culture medium, as well as levels of DO and pH, have been comprehensively described in detail.

2

Materials All culture media components must be certified as cell culture approved. All applied liquid media must be prepared under sterile conditions. Inoculation of the disposable bag-like container with cells must be performed in a biological safety cabinet under sterile conditions. Cell-containing samples harvested from the disposable culture bag must be sterilely handled. Analysis of harvested cell-free samples of culture medium may be proceeded outside biological safety cabinet, but in a reasonably clean-class laboratory, as it is usually provided for cell or tissue culture.

Microcarrier-Supported Culture of Chondrocytes in Disposable Bioreactor

2.1 Microcarrier Conditioning

147

1. Biological safety cabinet Herasafe™ KS 12 (Thermo Fisher Scientific, USA). 2. Laboratory autoclave (Biolab Scientific Ltd., Canada). 3. Mixer Vortex MX-S (Biologix Group, People’s Republic of China). 4. Manual pipette controller and presterilized disposable serological pipettes (25 mL, 10 mL, and 5 mL). 5. Automatic laboratory pipettes and presterilized disposable pipette tips (1 mL and 200 μL) placed in the racks. 6. 50 mL Falcon centrifuge tubes. 7. Cytodex 3 (Cytiva, USA), as microcarriers made of pig gelatine–coated cross-linked dextran beads, characterized by diameter equaled to 175 μm, surface area dry weight equaled to 0.27 m2 g
1 and beads quantity in dry weight at level of 4.3  106 g
1. 8. Dulbecco’s phosphate-buffered saline free of Ca2+ and Mg2+ cations (DPBS, Thermo Fisher Scientific, USA). 9. Dulbecco’s Modified Eagle’s Medium (DMEM, Thermo Fisher Scientific, USA) containing 4.5 g L
1 of glucose, 10% of inactivated fetal bovine serum (FBS, Thermo Fisher Scientific, USA), and 1% of commercial antibiotic–antimycotic mixture (PenStrep, Thermo Fisher Scientific, USA).

2.2 Preparation of CP5 Cell Inoculum

1. Biological safety cabinet Herasafe™ KS 12 (Thermo Fisher Scientific, USA). 2. CO2 incubator HF90 (Heal Force, People’s Republic of China). 3. Laboratory centrifuge for Falcon tubes MPW-352RH (MPW, Poland). 4. Reverse microscope Eclipse TS100 (Nikon, Japan). 5. Disposable Cellbag™ 2L vessel compatible with ReadyToProcess WAVE™ 25 system (Cellbag, Cytiva, USA). 6. Manual pipette controller and pre-sterilized disposable serological pipettes (10 mL and 5 mL). 7. Automatic laboratory pipettes and presterilized pipette tips (1 mL and 200 μL) placed in the racks. 8. 50 mL Falcon centrifuge tubes. 9. Articular cartilage progenitor cell line isolated from Bos taurus (CP5, ECACC, UK). 10. Dulbecco’s phosphate-buffered saline free of Ca2+ and Mg2+ cations (DPBS, Thermo Fisher Scientific, USA). 11. Dulbecco’s Modified Eagle’s Medium (DMEM, Thermo Fisher Scientific, USA) containing 4.5 g L
1 of glucose, 10%

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of inactivated fetal bovine serum (FBS, Thermo Fisher Scientific, USA), and 1% of commercial antibiotic–antimycotic mixture (PenStrep, Thermo Fisher Scientific, USA). 12. 0.05% trypsin–EDTA containing phenol-red (Thermo Fisher Scientific, USA). 2.3

Cell Culture

1. ReadyToProcess WAVE™ 25 (WAVE 25, Cytiva, USA) bioreactor system. 2. Disposable Cellbag™ 2L bag-like container adapted for WAVE 25 system. 3. Disposable 5 mL Luer Lock tip syringe (BD, Germany).

2.4 Analytical Methods

1. Biological safety cabinet Herasafe™ KS 12 (Thermo Fisher Scientific, USA).

2.4.1 Preparation of Samples for Analysis

2. Mixer Vortex MX-S (Biologix Group, People’s Republic of China). 3. Pipette controller, disposable serological pipettes (25 mL, 10 mL, and 5 mL), laboratory pipettes, pipette tips (1 mL and 200 μL), and pipette aids. 4. 2 mL Eppendorf tubes. 5. Syringe filter φ ¼ 0.2 μm WhatMan® Puradisc (Thermo Fisher Scientific, USA). 6. Dulbecco’s phosphate-buffered saline free of Ca2+ and Mg2+ cations (DPBS, Thermo Fisher Scientific, USA). 7. Dulbecco’s Modified Eagle’s Medium (DMEM, Thermo Fisher Scientific, USA) containing 4.5 g L
1 of glucose, 10% of inactivated fetal bovine serum (FBS, Thermo Fisher Scientific, USA), and 1% of commercial antibiotic–antimycotic mixture (PenStrep, Thermo Fisher Scientific, USA). 8. 0.05% trypsin–EDTA with phenol red (Thermo Fisher Scientific, USA).

2.4.2 Cell Staining

1. Reverse microscope Eclipse TS100 (Nikon, Japan). 2. Bu¨rker-Tu¨rk haemocytometer (Brand, Germany). 3. 0.4% trypan blue aqueous solution (Thermo Fisher Scientific, USA).

2.4.3 Activity of Intracellular Oxidoreductases

1. CO2 incubator HF90 (Heal Force, People’s Republic of China). 2. UV-VIS spectrophotometer GENESYS 20 (Thermo Fisher Scientific, USA).

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149

3. Freshly sterilized conditioned Cytodex 3 microcarriers (Cytiva, USA). 4. Resazurin-based PrestoBlue™ assay (PrestoBlue, Thermo Fisher Scientific, USA). 2.4.4 Glucose Consumption Rate

1. UV-VIS spectrophotometer GENESYS 20 (Thermo Fisher Scientific, USA). 2. BioMaxima-glucose enzymatic assay (BioMaxima, Poland). 3. Double-distilled water.

2.4.5 Activity of Lactate Dehydrogenase (LDH)

1. UV-VIS spectrophotometer GENESYS 20 (Thermo Fisher Scientific, USA). 2. BioMaxima-LDH enzymatic assay (BioMaxima, Poland). 3. Double-distilled water.

2.5 General Description of the Setup 2.5.1 ReadyToProcess WAVE™ 25 Bioreactor System

The schematic diagram of ReadyToProcess WAVETM25 system (WAVE 25, Cytiva, USA) setup has been presented in Fig. 1. WAVE 25 was equipped with a disposable pre-sterilized polymerbased 2 L Cellbag™ bag-like container (Cellbag, Cytiva, USA), which is characterized by working volume ranging from 0.1 L to 1.0 L of culture medium, as recommended by supplier of the system. The schematic diagram of Cellbag container and its ports configuration has been presented in Fig. 2. The thermostatic rocker performed continuously oscillating movements of whole Cellbag container, according to variously set conditions of wave-type agitation defined by: angle of oscillation, frequency of oscillation, and rocking motion acceleration. The control unit (CBCU) allowed to automatic measuring and controlling of crucial parameters of culture conditions, such as pH level and dissolved oxygen (DO) concentration (via miniaturized spectrophotometric sensors

Fig. 1 Schematic diagram of WAVE 25 bioreactor system

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Fig. 2 Schematic diagram of Cellbag container with arrangement of inlet and outlet ports: (1) Cellbag rods, (2, 6, 7) various pipe-ports, (3) outlet vent filter with pressure control valve, (4) inlet vent filter, (5) Screwcap port, (8) CLAVE™ sampling port

Table 1 The range of operating parameters that can be applied in WAVE 25 system Operating parameters

Range of parameter

Unit

Angle of oscillations

2–12



Frequency of oscillations

2–40

min
1

Gas flow rate

0.1–1.0

L min
1

Volume of liquid phase

0.1–1.0

L

Temperature

20–40



O2 concentration in gas phase

0–50

%

CO2 concentration in gas phase

0–15

%

Rocking motion acceleration

15–100

%

C

built-in inside the bottom of Cellbag), the concentration of O2 and CO2 in the inlet gas mixture, as well as the total gas flow rate. The range of operating parameters that can be applied in system of WAVE 25 has been presented in Table 1.

Microcarrier-Supported Culture of Chondrocytes in Disposable Bioreactor

3

151

Methods

3.1 Rehydration, Conditioning, and Sterilization of Microcarriers

1. Weigh 300 mg of Cytodex 3 microcarrier beads and transfer to 50 mL centrifuge Falcon tube. 2. Add 30 mL of Ca2+ and Mg2+ free DPBS to the 50 mL Falcon tube containing microcarriers, suspend them carefully and incubated at room temperature. After 4 h of microcarriers incubation in DPBS, slowly pipet out the supernatant. Next, add 20 mL of fresh Ca2+ and Mg2+ free DPBS into microcarrier-containing Falcon tube, and vortex shortly. After ca. 5 min, pipet out old DPBS and add 30 mL of fresh Ca2+ and Mg2+ free DPBS. 3. Autoclave the microcarriers at 121  C with 1 bar overpressure for 25 min. 4. Wait until the Falcon tube with microcarriers cool down and carefully pipet out the DPBS under the biological safety cabinet. Next, add 30 mL of sterile DMEM culture medium, vortex shortly, and leave for incubation at room temperature to accomplish microcarriers conditioning. After 30 min, slowly pipet out supernatant and add 30 mL of sterile fresh DMEM culture medium. Suspend the microcarriers by vortexing and then transfer them to the Cellbag container according to sterility regime (see Note 1).

3.2 Preparation of CP5 Chondrocyte Inoculum

1. Remove CP5 cell-containing culture flask from the incubator and carry out microscope observation. If confluence of CP5 cells in the culture flask is between 70% and 90%, the chondrocytes are ready for the next steps. 2. Transfer the culture flask to the biological safety cabinet. Carefully pipet out the culture medium from culture flask and flush still adhered CP5 cells twice with 5 mL of fresh Ca2+ and Mg2+ free DPBS. Next, detach CP5 cells by addition of 3 mL of trypsin–EDTA and leave shortly at room temperature to accomplish trypsinization. After ca. 5 min, add 5 mL of DMEM culture medium and suspend cells by gently shaking culture flask. 3. Immediately transfer suspended cells into new centrifuge Falcon tube and separate cells from medium by centrifugation (3800  g, 5 min). Then carefully decant supernatant and resuspend cells in 5 mL of fresh DMEM culture medium. 4. Count the number of cells using the methodology described in Subheading 3.4.2. Dilute cells to reach cell density of 6  105 cell mL
1. 5. Sterilely transfer 30 mL of diluted cells into microcarriercontaining Cellbag container prepared according to

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Subheading 3.1 (see Note 2). Incubate inoculated system at 37  C for ca. 4 h, that is, CP5 cells must completely attach to the outer surface of beads of the conditioned microcarriers. After this time, add 300 mL of DMEM culture medium to obtain final volume of the culture system inside Cellbag container (see Note 3). 3.3 Maintaining CP5 Cells in Bioreactor System

1. Install Cellbag container which contains microcarriers inoculated with CP5 in the rocker of WAVE 25 system. 2. Set the operating parameters (see Table 1) in control unit of WAVE 25 system (see Note 4). 3. Next, stabilize the temperature of culture system at level of 37  C, as well as DO concentration at the value at 100% saturation of the aqueous phase (for gas mixture composed of 21% O2, 5% CO2, and 74% N2). 4. Maintain oscillatory wave-agitated culture for 7 days, with daily monitoring and harvesting 5 mL samples of culture medium (see Note 5). To fully quantitatively characterize the culture parameters for each day of bioprocess, determination the values of cells density, viability of living cells, metabolic activity of cells, and specific glucose consumption rate (see Subheadings 3.4.2–3.4.4) as quantitative characteristics of biomass, as well as determination the values of DO, pH level and activity of lactate dehydrogenase (see Subheadings 3.4.5 and 3.4.6) as quantitative characteristics of culture medium, should be performed.

3.4 Analytical Methods 3.4.1 Preparation of Samples for Analysis

1. Use 3 mL of each of collected samples, in a form of suspension of cell-occupied microcarriers, to analyze metabolic activity of cells. 2. For other 2 mL of each of collected samples, separate celloccupied microcarriers from the culture medium by carefully pipetting out the culture medium. Filter culture medium through 0.22 μm syringe filter into new and sterile 2 mL Eppendorf tube. Sample of filtered culture medium will be used for analysis of specific glucose consumption rate and level of activity of lactate dehydrogenase. 3. Rinse the separated microcarriers occupied by CP5 cells with 2 mL of fresh Ca2+ and Mg2+ free DPBS for 5 min twice. Carefully pipet out DPBS, then add 1 mL of trypsin–EDTA, and gently vortex once a minute. After 5 min incubation at room temperature, add 1 mL of fresh culture medium (see Note 6). Vortex Falcon tube and immediately carefully pipet out cell-containing liquid above microcarriers (see Note 7). Cell-containing liquid phase will be used for counting cells, as well as for determination of living cells viability.

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1. Mix 50 μL of detached-cell suspension and 50 μL of 0.4% trypan blue aqueous solution, and immediately vortex it. 2. Next, incubate cell–dye mixture for 3 min at 37  C. 3. Count live (i.e., unstained) and dead (i.e., blue-stained) cells under reverse microscope equipped with the hemocytometer grid. 4. Calculate the value of cell density according to the following equation:
x X ¼  d  5  105 cells mL
1 k where X is total concentration of cells (i.e., value summarizing both stained and unstained cells), x is the total number of cells counted in the grid of the hemocytometer, k is the number of grid squares with cells, and d is the dilution of the sample containing cells. 5. Calculate the value of cells viability as follows. z Z ¼ ð%Þ x where Z is the viability of living cells, and z is the number of living (i.e., unstained) cells.

3.4.3 Metabolic Activity of Cells Adhered to Microcarriers

1. Mix 0.1 mL of PrestoBlue™ reagent and 0.9 mL of suspension of cell-occupied microcarriers (in the case of test samples) or 0.9 mL of suspension of freshly sterilized conditioned microcarriers (in the case of reference (i.e., blank) samples), and immediately vortex it shortly. 2. Next, incubate all samples for 2 h at 37  C. 3. After incubation measure the absorbance of all samples, that is, both test and reference ones, using UV-VIS spectrophotometer at 570 nm versus referenced wavelength equaled to 600 nm. 4. Calculate the values of metabolic activity of cells according to the following equation: A W ¼ ðA 570
A 570REF Þ
ðA 600
A 600REF Þ ½

a m ¼ 37, 04  A W μkat L
1 where AW is specific absorbance of the test sample, A570 is absorbance of the test sample at 570 nm, A570REF is absorbance of the reference at 570 nm, A600 is absorbance of the test sample at 600 nm, A600REF is absorbance of the reference sample at 600 nm, and am is metabolic activity of cells.

3.4.4 Specific Glucose Consumption Rate

1. Mix 20 μL of filtered culture medium collected daily from the cultures (in the case of test samples) or 20 μL of double-

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distilled water (in the case of blank sample) and 1 mL BioMaxima-glucose reagent, and immediately vortex it. 2. Next, incubate all samples for 20 min at 25  C. 3. Measure absorbance of all samples using UV-VIS spectrophotometer at 500 nm. 4. Calculate the values of specific glucose consumption rate according to following equation: r ∗ glc=cell ¼


C glc1
C glc2 g h
1 cell
1 X ∙Δt

where r*glc/cell is the specific glucose consumption rate, Cglc1, Cglc2 is the glucose concentration in the culture medium, and Δt is the time interval between two consecutive measurements of Cglc. 3.4.5 Activity of Lactate Dehydrogenase

1. Mix 1 mL of BioMaxima-LDH reagent and 20 μL of filtered culture medium harvested daily from cultures (in the case of test samples) or 20 μL of double-distilled water (in the case of blank sample) in directly in the disposable UV-Vis measuring cuvette, and vortex it shortly. 2. Next, as soon as it only possible, read the absorbance in 1-min intervals using UV-Vis spectrophotometer at 340 nm. 3. Calculate the values of activity of lactate dehydrogenase according to the following equation:
aLDH ¼ 267:2  ΔA μkat L
1 where aLDH is the activity of lactate dehydrogenase, and ΔA is the absorbance change per minute.

3.4.6 DO and pH Level Measurement

4

1. DO and pH level can be automatically measured by miniaturized spot-like sensors built-in inside the bottom of Cellbag container. The correctness of both values is verified and certified by the manufacturer of the culture bag according to setting the blank data for both sensors, which is printed individually on each applied Cellbag container, fixed to WAVE 25 system.

Notes 1. The most suitable way to transfer suspension of conditioned microcarriers from Falcon centrifuge tube into Cellbag container is manually pipetting them via Screwcap port. 2. Small volume (i.e., ca. 60 mL) of cell–microcarrier suspension introduced into Cellbag container provides proper conditions for contact between CP5 cells and beads of microcarrier which

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allows for sufficient adhesion of CP chondrocytes, as anchorage-dependent cells, onto microcarrier surface. 3. The addition of DMEM culture medium up to reach the total value of 300 mL liquid phase inside Cellbag container, allows obtaining starting cell concentration level equaled to 6  104 cell mL
1. 4. Based on our experience, the most suitable operating parameters supporting the most efficient propagation of CP5 cells biomass are as following: angle of oscillations equaled to 60, frequency of oscillations equaled to 20 min
1, gas flow rate equaled to 0.55 L min
1, volume of liquid phase equaled to 300 mL, temperature equaled to 37  C, O2 concentration in gas phase equaled to 21%, CO2 concentration in gas phase equaled to 5% and rocking motion acceleration equaled to 30%. In the case of other mammalian cells, the most suitable operating parameters will be depend strongly on resistance of given cells to hydrodynamic shear stress. Because of that, operating parameters must be individually designated for each cell line. 5. The most suitable way to harvest sample of culture medium from Cellbag container is to use syringe with Luer Lock tip and connect it to CLAVE™ sampling port, what allows collecting sample in sterile conditions outside the biological safety cabinet. 6. Addition of fresh DMEM medium allows to neutralize the proteolytic effects caused by trypsin. 7. The sedimentation time for microcarriers equaled to ca. 2 min. After microcarrier beads sediment to the bottom of Falcon centrifuge tube, cells can separate from microcarriers to slowly manual pipette out the cell-containing liquid phase over the layer of beads and transfer it into 2 mL Eppendorf tube. References 1. Shukla A, Tho¨mmes J (2010) Recent advances in large-scale production of monoclonal antibodies and related proteins. Trends Biotechnol 28:253–261. https://doi.org/10.1016/j. tibtech.2010.02.001 2. Kaiser S, Kraume M, Eibl D, Eibl R (2015) Single-use bioreactors for animal and human cells. In: Ai-Rubeai M (ed) Animal cell culture. Cell engineering. Springer, Basel, Switzerland 3. Wierzchowski K, Grabowska I, Pilarek M (2020) Efficient propagation of suspended HL-60 cells in a disposable bioreactor supporting wave-induced agitation at various Reynolds number. Bioproc Biosystems Eng

43:1973–1985. https://doi.org/10.1007/ s00449-020-02386-6 4. Singh V (2001) Method for culturing cells using wave-induced agitation, US Patent 6190913 5. Wierzchowski K, Grabowska I, Pilarek M (2019) Propagation of non-adherent HL-60 cells in batch cultures maintained in static and wave-type agitated systems. Chem Process Eng 40:167–177. https://doi.org/10.24425/cpe. 2019.126109 6. Eibl R, Eibl D (2011) Single-use technology in biopharmaceutical manufacture. Wiley, Hoboken

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7. Larroche C, Sanroman MA, Du G, Pandey A (2016) Current developments in biotechnology and bioengineering: bioprocess, bioreactors and controls. Elsevier, Amsterdam 8. Pilarek M, Sobieszuk P, Wierzchowski K, Da˛bkowska K (2018) Impact of operating parameters on values of a volumetric mass transfer coefficient in a single-use bioreactor with waveinduced agitation. Chem Eng Res Des 136:1–10. https://doi.org/10.1016/j.cherd. 2018.04.012 9. Kubis HP, Scheibe EJ, Decker B, Hufendiek K, Hanke N, Gros G, Meissner JD (2015) Primary skeletal muscle cells cultured on gelatin bead microcarriers develop structural and biochemical features characteristic of adult skeletal muscle. Cell Biol Int 40:364–374. https://doi. org/10.1002/cbin.10565 10. Li B, Wang X, Wang Y, Gou W, Yuan X, Peng J, Gou Q, Lu S (2015) Past, present, and future of microcarrier-based tissue engineering. J Orthop Translat 3:51–57. https://doi.org/ 10.1016/j.jot.2015.02.003 11. Tavassoli H, Alhosseini AN, Tay A, Chan PY, Oh AKW, Warkiani ME (2018) Large-scale production of stem cells utilizing microcarriers: a biomaterials engineering perspective from academic research to commercialized products.

Biomaterials 181:333–346. https://doi.org/ 10.1016/j.biomaterials.2018.07.016 12. Chen XY, Chen JY, Tong XM, Mei JG, Chen YF, Mou XZ (2020) Recent advances in the use of microcarriers for cell cultures and their ex vivo and in vivo applications. Biotechnol Lett 42:1–10. https://doi.org/10.1007/ s10529-019-02738-7 13. Wierzchowski K, Kuz´min´ska A, Pilarek M (2021) Intensification of chondrocytes proliferation by microcarriers and wave-induced mixing: Reynolds number influence on CP5 cells growth. Chem Eng Process 166: 108472. https://doi.org/10.1016/j.cep. 2021.108472 14. Stockwell RA (1978) Chondrocytes. J Clin Pathol Suppl (R Coll Pathol) 31:7–13 15. Phull AR, Eo SH, Abbas Q, Ahmed M, Kim SJ (2016) Applications of chondrocyte based cartilage engineering: an overview. Biomed Res Int 2016:1879837. https://doi.org/10. 1155/2016/1879837 16. Shah SS, Mithoefer K (2020) Scientific developments and clinical applications utilizing chondrons and chondrocytes with matrix for cartilage repair. Cartilage. (in press). https:// doi.org/10.1177/1947603520968884

Methods in Molecular Biology (2022) 2436: 157–165 DOI 10.1007/7651_2021_398 © Springer Science+Business Media, LLC 2021 Published online: 06 May 2021

Tracheal In Vitro Reconstruction Using a Decellularized Bio-Scaffold in Combination with a Rotating Bioreactor Georgia Pennarossa, Matteo Ghiringhelli, Fulvio Gandolfi, and Tiziana A. L. Brevini Abstract Long-segment airway stenosis as well as their neoplastic transformation is life-threatening and still currently represent unsolved clinical problems. Indeed, despite several attempts, definitive surgical procedures are not presently available, and a suitable tracheal reconstruction or replacement remains an urgent clinical need. A possible innovative strategic solution to restore upper airway function may be represented by the creation of a bioprosthetic trachea, obtained through the combination of tissue engineering and regenerative medicine. Here we describe a two-step protocol for the ex vivo generation of tracheal segments. The first step involves the application of a decellularization technique that allows for the production of a naturally derived extracellular matrix (ECM)-based bio-scaffold, that maintains the macro- and micro-architecture as well as 9 the matrix-related signals distinctive of the original tissue. In the second step chondrocytes are seeded onto decellularized trachea, using a rotating bioreactor to ensure a correct scaffold repopulation. This multi-step approach represents a powerful tool for in vitro reconstruction of a bioengineered trachea that may constitute a promising solution to restore upper airway function. In addition, the procedures here described allow for the creation of a suitable 3D platform that may find useful applications, both for toxicological studies as well as organ transplantation strategies. Key words Bioprosthetic trachea, Chondrocytes, Decellularization, ECM-based bio-scaffold, Rotating bioreactor, Tissue engineering

1

Introduction Upper airway dysfunctions and diseases, such as stenosis and neoplastic transformation, represent life-threatening conditions and seriously affect the duration and quality of life due to altered breathing, speech, and swallowing [1]. Several studies have proposed different techniques to help cartilage repairing, however definitive surgical procedures are not presently available and a suitable tracheal reconstruction or replacement remains an unmet clinical need [2]. In this context, tissue engineering represents one of the most promising approach to generate bioprosthetic tracheas, suitable for organ transplantation. During the past 50 years, several attempts have been made to cope with the challenging problem of reconstructing long segmental tracheal defects and diverse

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Fig. 1 Decellularization protocol and macroscopic images illustrating changes in trachea color, turning from red to white, while maintaining original shape

synthetic materials have been evaluated for the production of tracheal scaffolds [3]. Although promising, the results obtained met limited success because of immunological complications and bacterial infections. In addition, the materials tested lacked many of the organ-specific biomechanical properties, namely flexibility, strength to avoid collapse, and the formation of airtight seals [4, 5]. We here describe a two-step protocol for the ex vivo creation of a bioprosthetic trachea. The first step involves a decellularization technique that allows for the production of a naturally derived extracellular matrix (ECM)-based porcine bio-scaffold (Fig. 1). In the second step, human chondrocytes are seeded onto the decellularized trachea, using a rotating bioreactor to ensure a correct scaffold repopulation (Fig. 2). Nonimmunogenic tracheal bio-scaffolds were derived by using a physical-chemical method to successfully eliminate the cell compartment, while preserving the macro- and micro-architecture and maintaining an intact ECM protein composition [6, 7]. The bio-scaffold was obtained from the pig which is an ideal source of organs for xenotransplantation because of its anatomical and physiological similarities to humans [8, 9]. In addition, the adult porcine trachea has been recently demonstrated to match the biomechanical properties of the human organ, including bending stiffness, radial supporting force, longitudinal elongation, residual stress, and bursting strength [10]. In the second step, human chondrocytes were used to repopulate the porcine bio-scaffold to generate “semi-xenografts,” where the ECM-based scaffold is animal-derived, and the repopulating cells have human origin, thus combining the advantages of both xenotransplantation and tissue bioengineering. A key aspect of the protocol described is represented by the use of a rotating bioreactor that ensures a dynamic repopulation system with several important advantages compared to the static culture approach. Indeed, the adoption of a bioreactor favors and positively impacts on ex vivo cell and tissue re-organization, ensuring the

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Fig. 2 Preparation of the repopulating cells, injection into the lumen of the ECM-based trachea bio-scaffold and its placement into the rotating bioreactor for long-term culture

physiologically relevant physical signals, such as shear stress, compression, pressure, and stretch. In our opinion, this is an important aspect because the strategy adopted tries to mimic as closely as possible the mechanically dynamic environment experimented by chondrocytes in vivo, thus providing the ideal milieu for tracheal segment reconstruction [11]. Furthermore, the rotational bioreactor used in the present protocol is functionally superior to static or spinner flask culture, since it is able to create optimal laminar flow conditions and lower shear stress [11, 12]. They method here reported is simple and highly efficient. It paves the way for in vitro trachea reconstruction and organ transplantation. At the same time, it allows for the bio-fabrication of

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patient specific in vitro platforms that provide increased similarity to the in vivo physiology and pathology. The 3D semi-xenografts may be used for the development of new therapies against upper airway diseases and inflammation and can contribute to advances in the prevention and treatments of life-threatening infections such as COVID-19.

2

Materials Prepare all solutions immediately before use (unless indicated otherwise).

2.1 Porcine Trachea Collection

1. Tracheas collected from a slaughterhouse. 2. Sterile plastic containers. 3. Ice container. 4. Dulbecco’s phosphate-buffered saline (PBS): dissolve 8 g of NaCl (137 mM), 200 mg of KCl (2.7 mM), 1.44 g of Na2HPO4 (8 mM), and 240 mg of KH2PO4 (2 mM) in 800 mL of distilled water. Adjust pH to 7.4. Add distilled water until volume is 1 L. Sterilize solution with autoclave and store at +4
C. 5. Antibiotic/Antimycotic Solution.

2.2 Generation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold

1. 50 mL centrifuge polypropylene tubes. 2. Water bath. 3. Orbital shaker. 4. 500 mL plastic or glass bottles. 5. Deionized water (DI-H2O). 6. Sterile water. 7. Ethanol. 8. Antibiotic/Antimycotic Solution. 9. 1% sodium dodecyl sulfate (SDS): dissolve 5 g of SDS in 500 mL of DI-H2O. 10. 1% Triton X-100: add 5 mL in 495 mL of DI-H2O. 11. 2% deoxycholate: dissolve 10 g of deoxycholate in 500 mL of DI-H2O. 12. Dulbecco’s phosphate-buffered saline (PBS): dissolve 8 g of NaCl (137 mM), 200 mg of KCl (2.7 mM), 1.44 g of Na2HPO4 (8 mM) and 240 mg of KH2PO4 (2 mM) in 800 mL of distilled water. Adjust pH to 7.4. Add distilled water until volume is 1 L. Sterilize solution with autoclave and store at +4
C.

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13. Chondrocyte culture medium: Ham’s F-12 Nutrient Mixture, 10% Fetal Bovine Serum (FBS), 2 mM L-glutamine, 50 μg/mL ascorbic acid-2-phosphate (ASC), 5.0 ng/mL basic fibroblast growth factor (bFGF), 10,000 U/mL penicillin G; 2.5 μg/mL Amphotericin B. 2.3 Human Chondrocyte Propagation and Maintenance

1. Human chondrocyte cell line (see Note 1). 2. T75 cell culture flasks. 3. 15 mL centrifuge polystyrene tubes. 4. Centrifuge. 5. CO2 incubator. 6. Inverted microscope. 7. Antibiotic/Antimycotic Solution. 8. Dulbecco’s phosphate-buffered saline (PBS): dissolve 8 g of NaCl (137 mM), 200 mg of KCl (2.7 mM), 1.44 g of Na2HPO4 (8 mM) and 240 mg of KH2PO4 (2 mM) in 800 mL of distilled water. Adjust pH to 7.4. Add distilled water until volume is 1 L. Sterilize solution with autoclave and store at +4
C. 9. Trypsin-EDTA solution: dissolve 0.5 g of porcine trypsin and 0.2 g of EDTA 4Na in 1 L of HBSS with phenol red. 10. Chondrocyte culture medium: Ham’s F-12 Nutrient Mixture, 10% Fetal Bovine Serum (FBS), 2 mM L-glutamine, 50 μg/mL ascorbic acid-2-phosphate (ASC), 5.0 ng/mL basic fibroblast growth factor (bFGF), 10,000 U/mL penicillin G; 2.5 μg/mL Amphotericin B.

2.4 Repopulation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold with Human Chondrocytes and Bioreactor Setup

1. Autoclave. 2. 15 mL centrifuge polystyrene tubes. 3. Centrifuge. 4. CO2 incubator. 5. Cell counting chambers. 6. Inverted microscope. 7. Sterile decellularized tracheal ECM-based bio-scaffold. 8. Sterile rotating bioreactor. 9. Dulbecco’s phosphate-buffered saline (PBS): dissolve 8 g of NaCl (137 mM), 200 mg of KCl (2.7 mM), 1.44 g of Na2HPO4 (8 mM) and 240 mg of KH2PO4 (2 mM) in 800 mL of distilled water. Adjust pH to 7.4. Add distilled water until volume is 1 L. Sterilize solution with autoclave and store at +4
C. 10. Trypsin-EDTA solution: dissolve 0.5 g of porcine trypsin and 0.2 g of EDTA 4Na in 1 L of HBSS with phenol red.

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11. Chondrocyte culture medium: Ham’s F-12 Nutrient Mixture, 10% Fetal Bovine Serum (FBS), 2 mM L-glutamine, 50 μg/mL ascorbic acid-2-phosphate (ASC), 5.0 ng/mL basic fibroblast growth factor (bFGF), 10,000 U/mL penicillin G; 2.5 μg/mL Amphotericin B.

3

Methods All the procedures described below must be performed under sterile conditions. All instruments touching or in connection to tissues, cells, scaffolds, and bioreactors have to be sterilized. Cell manipulation must be carried out under laminar a flow hood and cell cultures have to be maintained at 37
C during their handling using thermostatically controlled stages. All studies were reviewed and approved by the Ethical Committee of the University of Milan. All animal experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals, published by the US National Institutes of Health (NIH).

3.1 Porcine Trachea Collection

1. Collect tracheal segments from gilts weighing approximately 120 kg. 2. Transfer tracheas in cold sterile PBS containing antibiotic/ antimycotic solution (5 mL/500 mL) and transport them to the laboratory using ice container.

3.2 Generation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold

1. Wash extensively tracheal graft in fresh PBS. 2. Completely remove the PBS, place the trachea in an empty 50 mL tube and store organ at 80
C for at least 24 h (see Note 2). 3. Thaw trachea at 37
C for 30 min using a water bath. 4. Transfer trachea in a bottle containing 500 mL of 1% SDS. Place the bottle onto an orbital shaker at 200 rpm and incubate for 3 h at room temperature. 5. Remove SDS solution from the bottle containing the trachea and wash it with 500 mL of DI-H2O for 40 min using an orbital shaker at 200 rpm. 6. Remove DI-H2O from the bottle and add 500 mL of 1% Triton X-100. Incubate trachea for 12 h at room temperature in 1% Triton X-100, using an orbital shaker at 200 rpm. 7. Remove Triton X-10 solution from the bottle containing the trachea and add 500 mL of DI-H2O twice.

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8. Remove the last washing DI-H2O from the bottle and incubate trachea in 500 mL of 2% deoxycholate for 12 h at room temperature, using an orbital shaker at 200 rpm. 9. Remove deoxycholate and wash trachea with DI-H2O for 6 h at room temperature, using an orbital shaker at 200 rpm. Changes DI-H2O every 2 h (see Note 3). 10. Sterilize the tracheal ECM-based bio-scaffold in a water solution containing 70% Ethanol and 2% antibiotic/antimycotic solution for 30 min at room temperature, using an orbital shaker at 200 rpm. 11. Extensively wash the ECM-based bio-scaffold with PBS supplemented with 4% antibiotic/antimycotic solution at room temperature using an orbital shaker at 200 rpm. 12. Before chondrocyte repopulation, immerse the tracheal ECM-based bio-scaffold in chondrocyte culture medium for at least 1 h at room temperature. 3.3 Human Chondrocyte Propagation and Maintenance

1. Maintain human chondrocyte cell line in 5% CO2 incubator at 37
C. 2. Monitor cells daily. 3. Once cell culture has reached 80% confluency, carefully aspirate the culture medium from T75 flask (see Note 4). 4. Wash cells three times with 7 mL of PBS supplemented with 1% antibiotic antimycotic solution. 5. Add 2 mL of trypsin-EDTA solution (see Note 5) and incubate at 37
C until cell monolayer begins to detach from the bottom of the tissue culture dish and cells dissociate (see Note 6). 6. Dilute cell suspension in 8 mL of chondrocyte culture medium to neutralize trypsin action. 7. Dislodge cells by repeatedly and gently pipetting. 8. Collect cell suspension in a 15 mL centrifuge polystyrene tube and centrifuge at 300  g for 5 min. 9. Remove supernatant and resuspended chondrocytes in 10 mL of fresh culture medium. 10. Plate cells in a new culture dish and culture at 37
C in 5% CO2 incubator. Keep the passage ratio between 1:2 and 1:4, depending on growth rate (see Note 7). 11. Change medium every 2–3 days. 12. Maintain cells in culture until they have reached 80% confluency and passage them.

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3.4 Repopulation of the Decellularized Tracheal ECM-Based Porcine Bio-Scaffold with Human Chondrocytes and Bioreactor Setup

1. Carefully remove culture medium from culture dish, wash cells three times with PBS and incubate in 2 mL of trypsin-EDTA solution at 37
C until cell monolayer begins to detach (for detailed procedure see Subheading 3.3, steps 3 and 4). 2. Add 8 mL of chondrocyte culture medium to cell suspension and collect chondrocytes in a 15 mL centrifuge polystyrene tube. 3. Count cells using a counting chamber under an inverted optical microscope at room temperature. 4. Calculate the volume of medium needed to resuspend chondrocytes to obtain a concentration of 1  106 cells/cm2 of tracheal lumen bio-scaffold (see Note 8). 5. Centrifuge cell suspension at 300  g for 5 min. Carefully aspirate supernatant and resuspend cell pellet in the previously calculated volume (see Subheading 3.2, step 4) of chondrocyte culture medium. 6. Repopulate the tracheal ECM-based bio-scaffold by injecting chondrocytes into the tracheal lumen and carefully transfer it into CO2 incubator. 7. Allow chondrocyte adhesion for 2 h under static conditions. 8. Connect the repopulating tracheal ECM-based bio-scaffold to an autoclaved bioreactor, set a rotation of 5 rpm (see Note 9) and culture at 37
C in 5% CO2 incubator. 9. Change culture medium every 48 h.

4

Notes 1. It is possible to apply the protocol here described to other cell types, such as tracheal epithelial cells (HTEpCs). 2. Tracheal segments can be stored at 80
C for longer periods without causing ECM alteration. 3. At the end of decellularization protocol, it is a good practice to verify the efficiency of the process, by confirming cellular compartment removal (e.g., DNA quantification, DAPI and/or hematoxylin and eosin staining) and the retention of intact ECM components (e.g., collagen, elastin, glycosaminoglycans, etc.). 4. Confluency normally takes between 7–10 days. If cells are not confluent after 10 days, they are not successfully growing. 5. The trypsin volume here reported is necessary for detaching cell cultured in a T75 flask. When working with smaller flask or dish, scale down the volumes accordingly. 6. It usually takes 3–5 min.

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7. The recommended seeding density is 5000–20,000 cells per cm2. 8. The formula to calculate the final volume depends on the specific type of chamber used. Cells/μL ¼ Average number of cells per small grid  chamber multiplication factor  dilution. 9. The bioreactor provides continuous rotation of the tracheal bio-scaffold, exposing cells to a covering film of alternating gas and liquid phases.

Acknowledgments This research was funded by Carraresi Foundation. The Authors are members of the COST Action CA16119 In vitro 3-D total cell guidance and fitness (CellFit) and of the Trans-COST Actions Task-Force on Covid-19. References 1. Go T, Jungebluth P, Baiguero S et al (2010) Both epithelial cells and mesenchymal stem cell-derived chondrocytes contribute to the survival of tissue-engineered airway transplants in pigs. J Thorac Cardiovasc Surg 139:437–443 2. Grillo HC (2002) Tracheal replacement: a critical review. Ann Thorac Surg 73:1995–2004 3. Aoki FG, Varma R, Marin-Araujo AE et al (2019) De-epithelialization of porcine tracheal allografts as an approach for tracheal tissue engineering. Sci Rep 9 4. Birchall M, Macchiarini P (2008) Airway transplantation: a debate worth having? https:// pubmed.ncbi.nlm.nih.gov/18431223/ 5. Macchiarini P (2004) Trachea-guided generation: De´ja` vu all over again? https://pubmed. ncbi.nlm.nih.gov/15224015/ 6. Pennarossa G, Ghiringhelli M, Gandolfi F et al (2020) Whole-ovary decellularization generates an effective 3D bioscaffold for ovarian bioengineering. J Assist Reprod Genet:1–11 7. Pennarossa G, Ghiringhelli M, Gandolfi F et al (2021) Creation of a bioengineered ovary:

isolation of female germline stem cells for the repopulation of a decellularized ovarian bioscaffold. Methods Mol Biol 2273:139–149 8. Ekser B, Cooper DKC, Tector AJ (2015) The need for xenotransplantation as a source of organs and cells for clinical transplantation. Int J Surg 23(Pt B):199–204 9. Michel SG, Madariaga MLL, Villani V, et al (2015) Current progress in xenotransplantation and organ bioengineering. https:// pubmed.ncbi.nlm.nih.gov/25496853/ 10. Shi HC, Deng WJ, Pei C et al (2009) Biomechanical properties of adult-excised porcine trachea for tracheal xenotransplantation. Xenotransplantation 16:181–186 11. Lin CH, Hsu S-h, Huang CE et al (2009) A scaffold-bioreactor system for a tissueengineered trachea. Biomaterials 30:4117–4126 12. Kojima K, Bonassar LJ, Roy AK et al (2003) A composite tissue-engineered trachea using sheep nasal chondrocyte and epithelial cells. FASEB J 17:823–828

Methods in Molecular Biology (2022) 2436: 167–182 DOI 10.1007/7651_2021_431 © Springer Science+Business Media, LLC 2021 Published online: 10 September 2021

Bioreactor-Based De-epithelialization of Long-Segment Tracheal Grafts Alba E. Marin-Araujo, Siba Haykal, and Golnaz Karoubi Abstract Tissue engineering techniques to generate a graft ex vivo is an exciting field of research. In particular, the use of biological scaffolds has shown to be promising in a clinical setting. In this approach, decellularized donor scaffolds are obtained following detergent-based enzymatic treatment to remove donor cells and subsequently repopulated with recipient specific cells. Herein, we describe our bioreactor-based partial decellularization approach to generate hybrid tracheal grafts. Using a short detergent-based treatment with sodium dodecyl sulfate (SDS), we remove the epithelium and maintain the structural integrity of the donor grafts by keeping the cartilage alive. The following will be a step-by-step description of the bioreactor system setup and partial decellularization protocol to obtain a de-epithelialized tracheal graft. Key words Biological scaffolds, Double-chamber perfusion bioreactor, Partial decellularization, Tissue engineering, Tracheal de-epithelialization

1

Introduction Tracheal transplantation using a tissue engineered graft is a promising alternative for treatment of extensive tracheal injury. The ultimate goal of tracheal tissue engineering is the generation of a functional construct to replace damaged upper airways. While synthetic grafts can provide structural integrity, recapitulation of the ECM components and architecture is challenging. An ideal approach allows for preservation of the extracellular matrix (ECM) which provides important biochemical cues required to support cellular function and maintains adequate biomechanical support [1, 2]. Thus, decellularization and recellularization of biological tracheal scaffolds is an exciting and promising area of research. While the specific details of each protocol are dependent on the tissue or organ of interest, the vast majority of decellularization protocols require a combination of physical, enzymatic, and/or chemical treatments [1]. In the trachea, fully decellularized scaffolds have come across challenges with stenosis due to

Siba Haykal and Golnaz Karoubi are co-senior authors.

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compromised cartilage [3, 4]. More recent studies suggest a plausible alternative approach of partial decellularization in which a hybrid graft is generated via removal of only the epithelium while maintaining the cartilage alive [5–7]. We have recently reported a novel bioreactor-based partial decellularization approach to generate hybrid tracheal grafts [8]. Using a short detergent-based treatment with sodium dodecyl sulfate (SDS), we remove the epithelium and maintain the structural integrity of the donor grafts by keeping the cartilage alive. The partial decellularization process is completed in an established double-chamber perfusion bioreactor system in which the inner tracheal lumen is treated with SDS while the outer chamber maintains the cartilage bathed in culture media [8–10]. Herein, we describe our tracheal bioreactor system and our de-epithelialization process in step-by-step detail.

2

Materials Place the bioreactor main components and the bioreactor extra components in autoclavable bags and autoclave. The autoclavable bags are to be opened and used inside the biosafety cabinet (BSC). Other materials and tools are to be used under sterile conditions in the biosafety cabinet as well but cannot be autoclaved (such as electronic components, pulleys, and other materials listed in Subheading 2.3). Those materials are to be disinfected with a 10% bleach solution followed by a wash in 70% ethanol solution before placement in the BSC.

2.1 Bioreactor Main Components (Fig. 1)

1. Stainless-steel body, lid, and silicone gasket. 2. Two chemical-resistant Viton® fluoroelastomer brown O-rings (3/32 fractional width #116). 3. Long and short rotating anchors. 4. Shaft holder, shaft, and corresponding rod. 5. Two stainless-steel rods with washer welded at one end.

2.2 Bioreactor Extra Components

1. Two tight-seal moisture-resistant plastic tube fitting reducers (1/800  1/1600 tube ID). 2. One plastic barbed swivel adapter tube fitting (1/800 tube ID  1/8 NPT male). 3. Three plastic barbed tube fitting adapters (1/800 tube ID  1/ 8 NPT male). 4. Seven plastic barbed tube fittings, polypropylene tee connector for 1/800 tube ID. 5. Four plastic barbed tube fittings, polypropylene 90 elbow for 1/800 tube ID.

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Fig. 1 CAD model of the double-chamber tracheal bioreactor. The bioreactor body and lid are shown as translucent. (a) Top view showing the position of the anchors and the trachea between them with a cable tie on each end for tracheal anchorage; (b) Lateral view showing the position of the motor, pulleys, belt, and shaft holder on the bioreactor right side

6. Two polycarbonate plastic tube caps for plugs. 7. High-purity silicone rubber soft tubing (1/800 ID 1/400 OD: Nine 5 cm long, thirteen 15 cm long, and three 25 cm long). 8. Tygon® 3-stop tubing S3™ E-LFL 1.85 mm ID. 9. Surgical instruments: three clamps, six forceps, four metal spatulas, two scissors, and one scalpel handle. 10. Stainless-steel instrument drying tray with rolled bead edge. 11. Two worm-drive clamps for firm hose and tube, steel screw, 1/200 wide band, 2–9/1600 to 3–1/200 clamp ID. 12. Silicone-based lubricant. 2.3

Other Materials

1. Timing belt (700 /35T, (3/800 ) Wide XL). 2. Small pulley: Timing belt pulley, XL series, 0.929“ OD, 10 teeth with “set-screw” for tightening. 3. Large pulley: Timing belt pulley, XL Series, 1.756“ OD with “set-screw” for tightening. 4. 37D L shaped gear motor bracket.

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5. 18–8 Stainless-steel socket head screw, M3  0.5 mm thread, 6 mm long for bracket and motor holding. 6. Super-corrosion-resistant flat-tip set screws, 316 stainless steel, 10–32 thread, 3/1600 long for shaft holder. 7. Four washers. 8. Teflon tape for pipe threads to ensure water-tight threads. 9. Sterile non-absorbable surgical silk suture #0–0 (2  20 cm). 10. Two polypropylene plastic narrow cable ties 400 Long. 11. Two polypropylene plastic standard cable ties 600 Long. 12. Three 125 mL sterile Erlenmeyer flasks with 2-hole rubber stopper. 13. One 250 mL sterile Erlenmeyer flask with 2-hole rubber stopper. 14. Three 500 mL Duran bottles for detergents and waste. 15. Five sterile hybridization bottles (for the tracheas). 16. USB temperature sensor (1/800 NPT stainless-steel housing with a 100 long probe). 17. Five 50 mL plastic pipettes. 18. Sterile cotton tip applicator. 2.4

Tools

1. Nine-piece metric hex L-key set. 2. Slip-joint pliers. 3. Flathead screwdriver.

2.5

Devices

1. 12 V 5 A switching power supply adapter. 2. 6 rpm precision spur gear motor. 3. Arduino 4-channel motor shield–1.2 A. 4. Electric coaxial cable. 5. Ismatec® Reglo peristaltic pump.

2.6 Antibiotics, Media, and Detergents

3-independent

channel

control

1. Imipenem/Cilastatin 500 mg/500 mg. 2. Ceftazidime 1 g. 3. Colistimethate 150 mg. 4. Fluconazole 400 mg/200 mL. 5. Cell culture grade water. 6. Dulbecco’s Modified Eagle’s medium high glucose glutamine. 7. Fetal bovine serum. 8. Gibco™ Antibiotic-Antimycotic (100).

L-

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9. Gibco™ Gentamicin (50 mg/mL). 10. Primocin™ Invivogen—Antimicrobial Agent. 11. Fungin™ Invivogen—Antifungal Reagent. 12. Sodium dodecyl sulfate. 13. Triton X-100. 14. Filtered deionized H2O. 15. PBS without Ca2+/Mg2+.

3

Methods

3.1 Preparation of Antibiotics, Media, and Detergents (See Notes 1 and 2) 3.1.1 Cocktail of Antibiotics

Prepare under sterile conditions inside the BSC using cell culture H2O: 1. Reconstitute one vial of Imipenem/Cilastatin 500 mg/ 500 mg with 8 mL of H2O, i.e., 67.5 mg/mL. 2. Reconstitute two vials of Ceftazidime 1 g by adding 5.2 mL of H2O to each vial, i.e., 192 mg/mL. 3. Reconstitute one vial of Colistimethate 150 mg with 6 mL of H2O, i.e., 25 mg/mL. 4. Measure out 340 mL of H2O and add: (a) 8 mL of Imipenem/Cilastatin. (b) 8 mL of Ceftazidime. (c) 4 mL of Colistimethate. (d) 40 mL of Fluconazole 400 mg/200 mL, i.e., 2 mg/mL. 5. Discard unused portions of Ceftazidime, Colistimethate, and Fluconazole. 6. Prepare 10 mL aliquots of the cocktail solution into 15-mL conical centrifuge tubes. 7. Store them at 20  C. Aliquots are stable at 20  C for up to 6 months.

3.1.2 DMEM + Antibiotics

Inside the BSC under sterile conditions: 1. Measure 430 mL of Dulbecco’s Modified Eagle’s Medium High Glucose L-Glutamine (DMEM). 2. Add: (a) 50 mL of fetal bovine serum. (b) 10 mL of cocktail of antibiotics. (c) 10 mL of Gibco™ Antibiotic-Antimycotic (100). (d) 0.5 mL of Gibco™ Gentamicin (50 mg/mL).

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(e) 0.5 mL of Primocin™ Invivogen; (Note that this is an antimicrobial agent). (f) 0.5 mL of Fungin™ Invivogen; (Note that this is antifungal reagent). 3.1.3 1% Sodium Dodecyl Sulfate

1. Weigh 5 g SDS and add to a 500-mL Duran bottle. Avoid inhalation using a fume mask or weigh the powder under a fume hood. 2. Measure 500 mL of deionized H2O and add to the Duran bottle. 3. Add a magnetic stir bar and place on a magnetic stirring plate to ensure mixing of the solution; use a low-to-medium speed and gently heat up the solution (60  C) using a hot plate. 4. Verify and adjust the pH to 7.4 (see Note 3). 5. Filter the solution inside the BSC and store at room temperature (RT).

3.1.4 1% Triton X-100

1. Measure 500 mL of deionized H2O and add to a 1000-mL Duran bottle. 2. Use a plastic pipette to add 5 mL of Triton X-100 to the Duran bottle. 3. Add a magnetic stir bar and place on a magnetic stirring plate to mix the solution; use a low-to-medium speed and gently heat up the solution (60  C) using a hot plate. 4. Verify and adjust the pH to 7.4 (see Note 3). 5. Filter the solution in the BSC and store the solution at RT.

3.2 Graft Procurement and Preparation

Use sterile surgical instruments (see Note 4). 1. At the Operating Room (OR): (a) Procure the trachea aseptically in a sterile fashion following standard surgical procedures (see Note 5). (b) Scrape the tracheal luminal surface with a sterile metal spatula to remove residual mucus. (c) Submerge the trachea in a sterile hybridization bottle with 100 mL DMEM + Antibiotics, then transport it to the laboratory. 2. At the laboratory: (a) Place the bottle with the trachea in a rocker at low-medium speed for 1 h at RT. (b) Inside the BSC, transfer the trachea to another sterile bottle with fresh DMEM + Antibiotics after scraping the lumen with another sterile spatula. (c) Repeat steps (a and b) twice to complete a 3-h wash period.

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1. Put on gloves, remove everything from the BSC, and disinfect it with 70% ethanol. 2. Disinfect tools, DMEM + Antibiotics, detergents, deionized filtered H2O, PBS, and other materials one by one (see Note 6).

3.3.1 Detergent Circuitries

1. Label the Erlenmeyer flasks with a marker, and fill each flask with the respective solution (a, b, c, or d) using one plastic pipette per flask as follows: (a) Add 250 mL of H2O to the 250 mL flask. (b) Add 140 mL of Triton X-100 to a 125 mL flask. (c) Add 140 mL of PBS to a 125 mL flask. (d) Add 75 mL of SDS to a 125 mL flask. 2. Insert two tubing into each rubber stopper as follows (see Note 7): (a) H2O flask: Place one 5 cm tubing in one stopper hole, and one 25 cm tubing in the other hole down to the bottom of the flask. (b) Triton X-100 flask: Place one 5 cm tubing in one stopper hole, and one 15 cm tubing in the other hole down to the bottom of the flask. (c) PBS flask: Place one 5 cm tubing in one stopper hole, and one 15 cm tubing in the other hole down to the bottom of the flask. (d) SDS flask: Place two 25 cm tubing in each stopper hole, both down to the bottom of the flask. 3. Introduce each stopper into the corresponding flask. 4. Attach one Tee connector to each of the following: H2O short tubing, both Triton X-100 tubing, and PBS long tubing (Fig. 2). 5. Attach one elbow connector to each of the following: H2O long tubing, PBS short tubing, and both SDS tubing. 6. Connect both H2O and Triton X-100 flask short tubing using a 15 cm tubing. 7. Connect the Triton X-100 and PBS flask short tubing using a 15 cm tubing. 8. Connect both H2O and Triton X-100 flask long tubing using a 15 cm tubing. 9. Connect the Triton X-100 and PBS flask long tubing using a 15 cm tubing. 10. Attach a 15 cm tubing to H2O short tubing tee connector. Then, connect it to a one-way valve (flow direction towards the H2O flask).

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Fig. 2 Schematic representation of the bioreactor system setup for tracheal de-epithelialization. Length of certain tubing is specified on the side in centimeters. Tube-A and Tube-B connect the SDS circuitry with the detergent-waste system. Electronic circuitry not shown

11. Attach a 5 cm tubing to the valve and a Tee connector to it. Then, (a) Attach two 5 cm silicone tubing to the Tee connector. (b) Connect a one-way valve to one tubing (flow direction towards the tee connector) and a 0.2 μm filter to it. (c) Attach another one-way valve to the second tubing and connect a 5 cm tubing to its other end. Connect a 60 mL syringe to it, this syringe will be positioned later onto the syringe pump. 12. Attach a 15 cm tubing (Tube-A) to the PBS long tubing tee connector. (a) Connect it to one SDS tubing using another tee connector. (b) Attach a 15 cm tubing to the third end of the tee connector. This tubing will be connected to the bioreactor LEFT side. 13. Attach a 15 cm tubing to the second SDS tubing. Connect it with a Tygon® 3-stop tubing using a fitting reducer. 14. Attach a 15 cm tubing to the Tygon® using a fitting reducer. Connect the tubing to another 15 cm tubing using a tee connector. This tubing will be connected on the RIGHT side of the bioreactor. 15. Attach a 15 cm tubing (Tube-B) to the last end of the tee connector and insert it into the waste bottle. 16. Put aside the flasks and tubing circuitries, while keeping them in the BSC.

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1. Open the sterile tray package inside the BSC (see Note 8). Then, open the autoclaved main and extra bioreactor components, and place them on the tray. 2. Position the motor bracket onto the bioreactor anterior wall. Place four washers between the bracket and the bioreactor. Using an Allen key, screw on the bracket with four screws (Fig. 1a). 3. Insert a barbed tube fitting adapter into the short anchor threaded end, and a swivel joint into the long anchor threaded end (see Note 9). 4. Place one O-ring on each end of the bioreactor body. Lubricate the O-rings with the autoclaved silicone-based lubricant using a sterile swab/applicator. 5. Place the two rotating anchors on opposite ends of the bioreactor’s body; the anchors should be in contact with the O-rings. Place the short anchor on the left end and the long one on the right end. 6. Adjust the position of the long anchor with respect to the bioreactor body to accommodate the trachea (note that the system would allow a trachea of any length from 3 to 10 cm). Insert the large pulley into the long rotating anchor, secure it by tightening the screw. 7. Wrap the thread portions of two plastic barbed tube fittings with Teflon tape to ensure water-tight threads. Insert one barbed tube fitting into the threaded hole at the anterior bioreactor wall, and another tube fitting at the posterior one. 8. Wrap the temperature sensor threaded section with Teflon tape. Screw the sensor long probe into the second hole at the bioreactor anterior wall. 9. Using the pliers, make sure the fittings and the sensor have a tight seal. 10. Connect a 5 cm silicone tube to each barbed tube fitting protruding from the bioreactor body (anterior and posterior walls). Place one plastic cap into each previously positioned tubing. 11. Introduce the 15 cm tubing (Subheading 3.3.1, step 15, i.e., bioreactor RIGHT side tubing) into the shaft placed in the shaft holder; secure the tubing by tightening the shaft screw. Place the shaft holder approximately 6–7 cm from the bioreactor using its rod; secure the shaft holder by tightening its screw at the bottom onto the rod. Attach the rod-shaft holder to the right side of the bioreactor; secure its position tightening the screw at the bottom of the bioreactor body.

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12. Introduce the 15 cm tubing into the short anchor fitting protruding at the left of the bioreactor (Subheading 3.3.1, step 12b, i.e., bioreactor LEFT side tubing). 3.3.3 Positioning the Trachea

1. Disinfect the hybridization bottle (with the trachea) before placing it inside the BSC (see Note 6). 2. Open a sterile package with clamps, forceps, and scissors (see Note 4). 3. Clamp the tubing attached to both anchors. 4. Fill the bioreactor with 225 mL of DMEM + Antibiotics using a plastic pipette. 5. Put on sterile gloves. 6. Place the trachea inside the bioreactor’s body and keep it submerged in DMEM + Antibiotics, using a forceps. 7. Fit the two ends of the trachea over each anchor with grooved tips to prevent slippage. Secure the trachea with one tight 400 -long cable tie, using the pliers and tweezers to make it tight. Then make a triple knot using a double-looped 20 cm silk suture on each end of the trachea. Place the suture adjacent to the cable tie on each end (see Note 10). 8. Connect the two anchors using two rods, specifically by placing the rods’ washers into the small anchor’s grooves (Fig. 1b). 9. Secure the rods with one tight 600 long cable tie at each end. 10. Position and secure the lid with the silicone gasket on top of the body by tightening two worm-drive hose clamps around the bioreactor body and lid. Use a screwdriver for this step. 11. Remove the clamps. 12. Place all flasks, tubing, and clamps on the tray; then carefully transfer the bioreactor system to the incubator.

3.3.4 Motor and Pumps Installation

1. Place the bioreactor system inside the incubator at 37  C. 2. Place the small pulley onto the motor and secure it by tightening its screw. 3. Attach the programmable gear motor to the motor bracket using four screws. 4. Make sure that the timing belt is connecting the large and small pulleys. 5. Place the Tygon® 3-stop tubing (SDS circuitry) onto the peristaltic pump. Set its flow to 7.5 mL/min. 6. Secure the 60 mL syringe onto the syringe pump. 7. Connect the motor to the Arduino motor shield. Then, connect the Arduino to the electric power using an electric coaxial cable and a 12 V 5 A switching power supply adapter.

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8. Program the Arduino for a speed rotation of 5 rpm. The bioreactor system is ready for de-epithelialization as shown in Fig. 2. 3.4 Tracheal Graft De-epithelialization

1. Start the tracheal rotation at 5 rpm inside an incubator at 37  C. 2. Clamp Tube-A and Tube-B from Subheading 3.3.1, steps 12 and 15. 3. Fill up the SDS closed-loop circuitry and tracheal luminal chamber with 1% SDS at a flow rate of 5 mL/min using the peristaltic pump (Fig. 3). 4. Tilt the bioreactor at 45 for 5 min to remove the air inside the trachea. Continue the SDS step for 3 h. 5. Stop the peristaltic pump, then remove the clamps from TubeA and Tube-B. The main one-way flow detergent-waste system is now open. 6. Close the SDS circuitry by placing a clamp onto both SDS tubing (see Note 11). Used SDS will remain in its reservoir until the end of the process. 7. Clamp both Triton X-100 tubing (see Note 11). 8. Clamp both PBS tubing (see Note 11). 9. Wash the tracheal lumen with 240 mL of filtered deionized H2O for 40 min using the syringe pump at flow rate of 6 mL/ min (Fig. 4). Used H2O should be seen in the waste bottle. 10. Remove the Triton X-100 tubing clamps.

Fig. 3 SDS step schematic representation. Two clamps are positioned on each side of the tubing main circuitry (Tube-A and Tube-B) to make an SDS closed system

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Fig. 4 H2O step schematic representation. SDS, Triton X-100, and PBS tubes are clamped on top of their respective reservoirs. H2O flows through the tubing main circuitry, to the trachea and ends in the waste bottle

Fig. 5 Triton X-100 step schematic representation. H2O and PBS tubes are clamped on top of their respective reservoirs. Triton X-100 washes the trachea and ends in the waste bottle. SDS circuitry remains closed until the end of the de-epithelialization

11. Clamp both H2O tubing (see Note 11). 12. Flow 140 mL of 1% Triton X-100 into the lumen for 30 min using the syringe pump at flow rate of 4.5 mL/min (Fig. 5). Used Triton X-100 should end up in the waste bottle. 13. Clamp Triton X-100 tubing (see Note 11). 14. Remove the PBS tubing clamp.

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Fig. 6 PBS step schematic representation. H2O and Triton X-100 tubes are clamped on top of their respective reservoirs. PBS flows through the tubing main circuitry and ends in the waste bottle. This is the final step of the tracheal de-epithelialization

15. Remove the detergents with 140 mL of PBS for 30 min using the syringe pump at a flow rate of 4.5 mL/min (Fig. 6). Now, the waste bottle should be filled with H2O, Triton X-100, and PBS. 16. Remove the motor and small pulley from the system using an Allen key. 17. Take the bioreactor system to the BSC using the tray, disinfect the surfaces with 70% ethanol. 18. Disinfect a sterile hybridization bottle with fresh DMEM + Antibiotics and place it inside the BSC. 19. Loosen and remove the two worm-drive hose clamps placed around the bioreactor body and lid using the screwdriver. 20. Remove the lid on top of the body and expose the trachea. 21. Cut the 600 long cable ties using the scissors and remove both rods. 22. Cut the trachea close to the anchors transversely. Use sterile forceps to determine the edge of each anchor. 23. Put the trachea into the bottle with DMEM + Antibiotics. 3.5 Decontamination of the Deepithelialized Graft

1. Place the bottle with the trachea in a rocker at low-medium speed for 24 h at RT. 2. Disinfect the bottle with 70% ethanol and place it inside the BSC. 3. Disinfect a new bottle of fresh DMEM + Antibiotics with 70% ethanol and place it in the BSC.

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4. Change the trachea from one bottle to the new one using sterile forceps. 5. Place the tracheal bottle on the rocker at low-medium speed for 24 h at RT. Please see Note 12 for the approximate time period expected for each of the described steps.

4

Notes 1. Make solutions, media, and detergents at least the day before setting up the bioreactor. Filter the deionized water as well. It may take 3 or 4 h. 2. Sterilization of biological scaffolds for tissue engineering applications carries the risk of unwanted changes in physical and chemical structure [11]. We designed an antimicrobial solution-based disinfection method for partially decellularized tracheal grafts. The concentrations of antibiotics and antimycotics were optimized for further in vitro and ex vivo airway epithelial cell differentiation protocols (human tracheal epithelial cells and human induced pluripotent derived airway progenitors). 3. Verify and adjust the pH to 7.4: If the pH is higher than 7.4, add a few drops of hydrochloric acid solution. If the pH is lower than 7.4, adjust it using sodium hydroxide solution. Use a plastic pipette with the correct solution and add a few drops to the solution in the beaker and wait at least 20 s before reading the pH on the meter. 4. Make different packages to autoclave for ease of handling. Packages of surgical instruments include the following: (a) For OR use, include one of each of the following: forceps, spatula, scissors, and scalpel. (b) For lab use and graft preparation, make two packages with each of the following instruments: forceps, spatulas. (c) For the tracheal positioning step, make one package with the following: three clamps, one forceps, one spatula, and a pair of scissors. (d) For the graft decontamination step, make a package with only one forceps. 5. Procure the trachea aseptically in a sterile fashion using standard surgical procedures [12]: (a) Place the pig in supine position. (b) Make a skin incision along the anterior neck and overlying the sternum.

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(c) Use a saw to open the sternum and expose the mediastinal contents. (d) Dissect the midline neck and strap muscles, connective tissue and mobilize the thyroid to expose the trachea. (e) Dissect circumferentially around the trachea as well as proximally and distally into the chest to expose it above the carina (note: there is variability in anatomy with pigs having a separate bronchus above the carina). (f) Use forceps to grasp the proximal portion of the trachea and using scissors make an incision in the trachea below the larynx. (g) Lift the trachea caudally and snip any attachments of the mediastinal tissue. (h) Make an incision in the trachea at the carina or above the bronchus to separate it from the lungs. 6. Wipe with 10% bleach, spray 70% ethanol, then dry with one paper towel at a time before putting any non-autoclaved objects inside the BSC. 7. Make the tubing circuitries on the bench and send them for autoclaving, including the rubber stoppers with the tubes in their holes. Replace the tubing with new ones when they start to wear off, perhaps every two or three experiments. 8. Make one package with the bioreactor main components and another one with the extra components. Package the tray separately and autoclave the silicone-based lubricant in a 50 mL glass bottle. Have extra packages with sterile surgical instruments, tubing, and tube fittings just in case they are needed. 9. Prevent leakage by using Teflon tape which may be needed for the barbed swivel adapter tube fitting and the barbed tube fitting adapter that goes into the anchors. 10. Position cable ties and silk sutures to prevent the communication between the inner and outer chambers; this will prevent detergents from being in contact with the outer wall of the trachea. 11. Place the clamp on the tubing protruding from the top of the stopper. 12. The estimated time for each of the steps is as follows: (a) Preparation of media, antibiotics, and detergents: 4 h. (b) Graft procurement and preparation: 4 h. (c) Bioreactor setup for de-epithelialization. l

Detergent circuitries: 30 min.

l

Bioreactor setup: 10 min.

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Trachea positioning: 5–8 min. This time may be variable, the trachea is submerged in medium, therefore it could take as long as needed.

(d) Motor and pump installation: 10 min. (e) Tracheal graft de-epithelialization: 5 h. (f)

Graft decontamination: 36–48 h.

Acknowledgments This work was supported in part of the University of Toronto’s Medicine by Design initiative, which receives funding from the Canada First Research Excellence Fund (CFREF; C1TPA-201618). References 1. Gilbert TW, Sellaro TL, Badylak SF (2006) Decellularization of tissues and organs. Biomaterials 27:3675–3683 2. Scarrit ME, Pachos NC, Bunnell BA (2015) A review of cellularization strategies for tissue engineering of whole organs. Front Bioeng Biotechnol 3:1–17 3. Remlinger NT, Czajka CA, Juhas ME et al (2010) Hydrated xenogeneic decellularized tracheal matrix as a scaffold for tracheal reconstruction. Biomaterials 31:3520–3526 4. Batioglu-Karaaltin A, Karaaltin MV, Ovali E et al (2015) In vivo tissue-engineered allogenic trachea transplantation in rabbits: a preliminary report. Stem Cell Rev Rep 11:347–356 5. Delaere P, Van Raemdonck D (2016) Tracheal replacement. J Thorac Dis 8:S186–S196 6. Delaere P, Vranckx J, Verleden G et al (2010) Tracheal allotransplantation after withdrawal of immunosuppressive therapy. N Engl J Med 362:138–145 7. Delaere PR (2012) Tracheal transplantation. Curr Opin Pulm Med 18:313–320 8. Aoki FG, Varma R, Marin-Araujo AE et al (2019) De-epithelialization of porcine tracheal

allografts in a bioreactor system: implications in tissue engineering approaches for tracheal replacement. Sci Rep 9:12034 9. Lee H, Marin-Araujo AE, Aoki FG et al (2021) Computational fluid dynamics for enhanced tracheal bioreactor design and long-segment graft recellularization. Sci Rep 11:1187 10. Haykal S, Salna M, Zhou Y et al (2014) Double-chamber rotating bioreactor for dynamic perfusion cell seeding of largesegment tracheal allografts: comparison to conventional static methods. Tissue Eng Part C Methods 20:8 11. Łopianiak I, Butruk-Raszeja BA (2020) Evaluation of sterilization/disinfection methods of fibrous polyurethane scaffolds designed for tissue engineering applications. Int J Mol Sci 21:8092 12. JoVE Science Education Database (2021) Lab animal research, sterile tissue harvest protocol. JoVE, Cambridge, MA. https://www.jove. com/v/10298/sterile-tissue-harvest. Accessed 12 Jun 2021

Methods in Molecular Biology (2022) 2436: 183–192 DOI 10.1007/7651_2021_413 © Springer Science+Business Media, LLC 2021 Published online: 07 September 2021

Production of Extracellular Vesicles Using a CELLine Adherent Bioreactor Flask Anastasiia Artuyants, Vanessa Chang, Gabrielle Reshef, Cherie Blenkiron, Lawrence W. Chamley, Euphemia Leung, and Colin L. Hisey Abstract The efficient production of extracellular vesicles (EVs) from adherent cells in vitro can be challenging when using conventional culture flasks. Issues such as low cell density leading to low EV yield, and the inability to completely remove bovine serum EVs without starvation contribute to this challenge. By comparison, the two-chamber CELLine adherent bioreactor can produce significantly more EVs with improved time, space, and resource efficiency. Furthermore, it is highly accessible and can continually produce EVs using long term cultures without the need for passaging. Lastly, the 10 kDa semipermeable, cellulose acetate membrane separating the cell and media chambers allows for the continual use of bovine serum in the media chamber while preventing bovine EVs from contaminating the conditioned media. Key words CELLine bioreactor, Exosomes, Extracellular vesicles, Microvesicles

1

Introduction To date, most researchers producing EVs from cell cultures rely on the use of multiple conventional culture flasks due to the low-density monolayers of cells on the surface of the flasks. To maintain multiple flasks, large volumes of culture media are required, which must then be concentrated using ultrafiltration or multiple ultracentrifuge spins for downstream characterization and use of the EVs. In addition, this method requires the use of significant amounts of single use polystyrene, user time, and incubator space. Importantly, bovine or other animal serum is routinely used in conventional cultures to support cell growth up to the point of conditioned media collection, but the endogenous EVs and other nanoparticles (e.g., protein/growth factor aggregates) within the serum are nearly impossible to fully deplete, with researchers often resorting to up to 72 h of serum starvation prior to media collection with confounding effects [1–3]. Lastly, these EV productions are typically done as one-off experiments, causing the final EV preparation to be seen as a precious material, ultimately discouraging more creative and high-risk experimental designs.

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Fig. 1 CELLine AD 1000 Bioreactor flask schematic illustration. (Figure created with BioRender.com)

Bioreactors originally designed for monoclonal antibody production have shown immense promise as a solution to many of the aforementioned issues when appropriately repurposed for EV production. These include hollow-fiber flow systems like the FiberCell bioreactor [4–6], and two-chambered static systems like the CELLine AD 1000 bioreactor flask [7–14]. Both approaches eliminate the risk of endogenous serum EV contamination by separating cell and media chambers using membranes with molecular cutoff sizes much smaller than EVs, while also eliminating the need for cell passaging. However, the CELLine AD 1000 system (Fig. 1) differs in that it is a static system which requires no powered pumps and takes up only slightly more space than a T-175 flask. Furthermore, the 15 mL cell chamber and 1 L media chamber provide a highly concentrated conditioned media product with minimal maintenance. In combination, these benefits will enable researchers to freely explore many other applications of EVs by producing much larger amounts of EVs from long-term cultures, with significantly less space and time requirements. In this protocol, we demonstrate a simple workflow to inoculate, maintain, and monitor the CELLine AD 1000 bioreactor flask for EV production, as well as the basic steps for isolation and characterization of the resulting EVs as defined by the MISEV guidelines [15]. Lastly, we present a method for imaging the 3D growth of cells on the bioreactor surface once EV production has ended.

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2 2.1

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Materials Cell Culture

1. Obtain cells, media, flasks, and CELLine AD 1000 bioreactor flask (see Notes 1 and 2). 2. Culture materials. (a) Sterile PBS. (b) DMEM, RPMI or other media. (c) Advanced media (optional). (d) Fetal bovine serum (FBS) or other appropriate serum. (e) Penicillin–streptomycin (PS). (f) CDM-HD serum replacement (see Note 3).

2.2 EV Isolation Materials

1. 35 nm qEV Original size exclusion chromatography (SEC) column or equivalent column 2. Eppendorf 1.5 mL LoBind tubes. 3. Ultracentrifuge tubes, ultracentrifuge, and fixed-angle rotor (Beckman Avanti J-30I or 100,000
g capable equivalent).

2.3 Characterization Materials

1. Bicinchoninic acid (BCA) kit or comparable protein assay. 2. NanoSight NS300 or comparable nanoparticle counting and sizing system. 3. Transmission electron microscopy (TEM) reagents (copper grids, 2% uranyl acetate). 4. Western blot membranes).

reagents

(blocking

solution,

antibodies,

5. 4% glutaraldehyde.

3

Methods

3.1 Preparation of CELLine AD 1000 Bioreactor Flask (Figs. 1 and 2)

1. Add 50 mL of prewarmed inoculation culture media in the media chamber and let the semipermeable membrane equilibrate for 5 min.

3.2 Inoculation of Cells (See Note 4)

1. Obtain at least 2.5
107 viable cells from a preculture in a growth phase and resuspend the cells in 15 mL media containing 10% FBS and 1% PS (this equates to approximately 5–8 T175 flasks of cells depending on the cell type and confluence). 2. Always loosen the media chamber cap when exchanging fluid in the cell chamber to prevent an air lock (Fig. 2a) which can damage the delicate semipermeable membrane (see Note 5).

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Fig. 2 Step-by-step process of continually harvesting EVs from the CELLine AD 1000 bioreactor. (a) Loosening the cell and media chamber caps, (b) removing media from the media chamber, (c) aspirating conditioned media from the cell chamber, (d) transferring conditioned media to a falcon tube for centrifugation, (e) washing the cell chamber with PBS, (f) adding fresh media to the cell chamber, (g) adding fresh media to the media chamber, and (h) returning bioreactor to incubator

3. Collect the 15 mL preculture cell suspension with a 25 mL serological pipette, open the cell chamber (lower compartment) and inoculate it by inserting the pipette into the black silicone cone (Fig. 2f). Close the cell chamber by completely tightening the cap. 4. Add 450 mL of additional media into the media chamber and then completely tighten the cap (Fig. 2g). 5. If bubbles are present in the cell chamber, tilt the reactor so that the bubbles move toward the black silicone cone and remove the bubbles using a 25 mL serological pipette. 6. If droplets of media are spilled near the black silicone interface, use an ethanol-soaked wipe to clean the area. 7. Gently place the bioreactor into a standard 5% CO2 and 37  C incubator (Fig. 2h). 3.3 Adapting Cells to Long-Term Bioreactor Media (See Note 6)

1. Every 3–4 days, collect 15 mL conditioned media (Fig. 2c) using a 25 mL serological pipette inserted into the black silicone cone, gently wash cell chamber 2–3
with 15 mL prewarmed sterile PBS (Fig. 2e), and refill with 15 mL prewarmed media based on adaptation schedule (Fig. 2f). Every 7 days discard the 500 mL media in the media chamber (Fig. 2b) and refill with prewarmed media (Fig. 2g). Continual use of 1% PS in both chambers is optional but encouraged.

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2. After 7 days postinoculation, reduce the percentage of FBS in both chambers to 5% and use 50% DMEM/50% Advanced DMEM with 1% CDM-HD or other appropriate media. 3. After 14 days post-inoculation, reduce the percentage of FBS in both chambers to 1% and use 25% DMEM/75% Advanced DMEM with 2% CDM-HD or other appropriate media. 4. After 21 days post-inoculation, use 100% Advanced DMEM and 2% CDM-HD in both chambers, 1% FBS in the media chamber, and exclude FBS from the cell chamber. 3.4 Continually Harvesting EVs and Monitoring Shed Cells from the Bioreactor (Fig. 2)

1. The CELLine AD 1000 bioreactor is opaque and must be monitored using the conditioned media that is collected (volume and cell count). 2. Start collecting EVs after media adaptation has finished and the cell chamber is serum-free. 3. Collection of conditioned media and replacement (cell chamber) is done twice a week (can be done once a week or less depending on cell type) and media replacement in the media chamber is done once a week. 4. Loosen the cap of the cell chamber to prevent an air lock (Fig. 2a). Transfer all media from the media chamber to a waste container (Fig. 2b). 5. Aspirate 15 mL from the cell chamber using a 25 mL serological pipette inserted into the black silicone cone (Fig. 2c–d). Make a note of the volume (see Note 7). 6. Count the cell concentration and viability in the collected conditioned media using Trypan Blue exclusion method and a hemocytometer (see Note 8) or other suitable method (Fig. 3a). 7. Immediately centrifuge the conditioned media at 2000
g to remove cells and debris, then transfer supernatant to an ultracentrifuge tube. A 200
g spin prior to the 2000
g is optional. 8. Gently wash the cell chamber 2–3
with 15 mL prewarmed sterile PBS using a 25 mL serological pipette inserted into the black silicone cone (Fig. 2e). 9. Add 15 mL of prewarmed media to the cell chamber using a 25 mL serological pipette inserted into the black silicone cone (Fig. 2f). Add 500 mL of prewarmed media to the media chamber (Fig. 2g) and return the bioreactor to a 5% CO2 and 37  C incubator (Fig. 2h). 10. Check the conditioned media for microbial contamination microscopically if excessive cloudiness or discoloration is present (see Note 9). Some fat producing cells may produce visible amounts of lipids that cause cloudiness due to the high cell density.

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Fig. 3 Example HDFa (human dermal fibroblast) bioreactor data showing monitoring of shed cells, EV characterization based on MISEV guidelines [15], and SEM bioreactor growth surface. (a) The number of shed cells and their viability over the bioreactor’s 5 month lifetime, (b) SEC purification profile showing protein and particles amounts in each collected fraction, (c) NTA size distribution of pooled HDFa small EVs, (d) TEM image of purified HDFa small EVs (scale bar ¼ 100 nm), (e) Western blot showing CD81, CD9, GRP94, and GAPDH for HDFa small EVs (sEV) and cell lysates, and (f) SEM images of a growth surface of the HDFa bioreactor with and without cells (scale bars ¼ 100 μm) 3.5 EV Isolation and Purification (See Note 10)

1. Ultracentrifuge the supernatant from the 2000
g spin at 10,000
g for 30 min at 4  C (or optionally at 20,000
g). Transfer the supernatant to a new ultracentrifuge tube and resuspend the 10,000
g pellet in PBS or other suitable buffer if interested in large EVs and store (Fig. 1). 2. Ultracentrifuge the 10,000
g supernatant at 100,000
g for 70 min at 4  C. 3. Discard the supernatant. Resuspend the resulting crude small EV pellet (in 500 μL buffer for SEC) and store (see Note 11). 4. Purify the crude small EV suspension with SEC using a 35 nm qEV Original or other appropriate column (see Note 12). 5. See Size Exclusion Chromatography: A Simple and Reliable Method for Exosome Purification for extensive details [16].

3.6 Characterizing EVs

EVs should be characterized according to the MISEV2018 guidelines [15] prior to downstream molecular profiling or functional use in order to enhance rigor and reproducibility in EV studies. 1. Quantify protein using BCA (high sensitivity standards are recommended) or other appropriate assay.

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2. Measure particle size and a concentration with nanoparticle tracking analysis (NTA), tunable resistive pulse sensing (TRPS), or other counting and sizing system. (a) Settings will be sample-dependent, but setting consistency is important to be able to compare between different EV preparations. (b) If doing SEC for the first time, validate which fractions contain EVs (Fig. 3b) and pool EV-rich fractions for downstream experimentation (Fig. 3c). If necessary, EV-rich fractions can be concentrated using ultracentrifugation or 10–100 kDa cutoff filters. 3. Visualization of EV morphology using TEM (Fig. 3D). (a) See Detection and Characterization of Extracellular Vesicles by Transmission and Cryo-Transmission Electron Microscopy for extensive details [17]. 4. Western Blotting for EV and contaminant markers (see Note 13). (a) To prepare EV lysates, mix EV samples with Protease Inhibitor and radioimmunoprecipitation assay buffer (RIPA buffer). Some antibodies require reduction of protein using DTT or 2-mercaptoethanol. (b) At least two proteins from a category of EV-enriched proteins (CD9, CD63, CD81, Alix, TSG101, etc.) and one from the category of EV-depleted proteins (calnexin, GRP94, etc.) are recommended to be analyzed to demonstrate the EV nature and the degree of purity of an EV preparation [15] (Fig. 3e). (c) See Extracellular Vesicle Isolation and Analysis by Western Blotting for extensive details [18]. 3.7 Imaging the Bioreactor Growth Surface

1. Before terminating the bioreactor culture, wash 3
with prewarmed PBS. 2. Add 15 mL 4% glutaraldehyde or Karnovsky’s fixative to the cell chamber using a 25 mL serological pipette and 30 mL to the media chamber and leave at 4  C overnight. 3. Aspirate fixative from cell chamber and pour out from media chamber, then cut the bioreactor open from the bottom using a Dremel™ circular blade or similar tool using the outline of the growth surface as a guide. Carefully cut out regions of the growth surface of approximately 25 mm2 using a scalpel or scissors. 4. Transfer the regions of interest to the wells of a 6 well culture plate and gradually dehydrate from 100% PBS to 100% ethanol

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(0%, 30%, 50%, 70%, 80%, 90%, 100%) by submerging samples (2
5 min) at each progressive concentration. 5. Air-dry, sputter-coat with 10 nm gold, and image bioreactor surfaces using scanning electron microscopy (SEM) (Fig. 3f). We have found that 3D morphology differs greatly with cell type. 3.8 EV-Associated RNA

4

EV-associated RNA can be isolated by resuspending large or small EV pellets in 50–100 μL PBS and adding TRIzol, or by using other RNA purification reagents [9].

Notes 1. There is a variant of the CELLine flask available for suspension cells which requires passaging to maintain an appropriate cell number. This protocol has been optimized for adherent cells only. 2. Media choices will depend on the type of cells used. We recommend using commercially available defined “advanced media” formulations that contain serum replacements when possible to support long-term, high density growth with minimal serum usage, but some cells may perform better in standard media. Media formulations should be tested in conventional cultures using prior to inoculation. 3. The use of CDM-HD or other serum replacement is optional, and cells should be tested prior to bioreactor inoculation to determine if it affects their viability. We have found that CDM-HD offers great benefits with several different cell lines in maintaining cell viability and reducing serum usage. 4. Initially seeding adherent cells with 10% serum then weaning them off serum gradually seems to help them initially adhere to the fibrous polyethylene terephthalate (PET) growth surface. 5. When handling the bioreactor, it is important to do everything gently as the 10 kDa membrane is only 8 μm thick and is the most frequent cause of the bioreactor’s failure. 6. Media adaptation will be highly cell-dependent, with some cells able to be adapted to their final media prior to inoculation or adapted more quickly once inoculated. We recommend slow adaptation in most cases. 7. It is common to retrieve less than 15 mL from the cell chamber due to osmotic gradients, do not apply excessive suction with a serological pipette if less than 15 mL is collected because there is a risk of rupturing the membrane. Repeatedly retrieving more than 20 mL from the cell chamber is a sign that the

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membrane may have ruptured and one should consider terminating the bioreactor. 8. Once established, live and dead cells are shed into the cell chamber culture media at a variable rate throughout the life of the bioreactor culture. Do not misinterpret a large number of shed cells as a “dead” reactor. Often, these shed cells can be plated to demonstrate their viability or used as cell lysates for comparative western blots. The rate at which cells are shed into the conditioned media is highly dependent upon the cell type. 9. Testing for mycoplasma is recommended prior to plating as well as an agar growth of cell media during the bioreactor culture. 10. EVs can be isolated from conditioned media using several other methods, depending on the application. For instance, a second 100,000
g spin can be performed by resuspending the pellet from the first 100,000
g spin to further reduce contaminating proteins, or ultracentrifugation can be avoided altogether by using ultrafiltration and/or a larger size exclusion column. 11. EVs should be resuspended in minimal volume and used fresh when possible as all concentration processes have significant inherent EV-loss. However, they can be stored at 4  C short term, or 80  C for long term. Multiple freeze–thaw cycles are not recommended. 12. We recommend using an automated fraction collector for SEC. Fractions can be collected manually, but this will potentially introduce more variability in volume between fractions. Density gradient ultracentrifugation is a suitable alternative to SEC [15]. Other SEC columns are also suitable for use. 13. The use of biotinylated secondary antibodies and streptavidinconjugated horseradish peroxidase (HRP) can drastically reduce the amount of required EV protein loading (1–5 μg/ lane). We recommend this approach for most EV workflows. References 1. Lehrich BM et al (2018) Fetal bovine serumderived extracellular vesicles persist within vesicle-depleted culture media. Int J Mol Sci 19(11):3538 2. Shelke GV et al (2014) Importance of exosome depletion protocols to eliminate functional and RNA-containing extracellular vesicles from fetal bovine serum. J Extracell Vesicles 3 3. Lehrich BM, Liang Y, Fiandaca MS (2021) Foetal bovine serum influence on in vitro extracellular vesicle analyses. J Extracell Vesicles 10 (3):e12061

4. Yan IS, Shukla N, Borrelli DA, Patel T (2018) Use of a hollow fiber bioreactor to collect evs from cells in culture. In: Walker J (ed) Extracellular RNA: methods in molecular biology, vol 1740. Springer Protocols, New York 5. Watson DC et al (2018) Scalable, cGMPcompatible purification of extracellular vesicles carrying bioactive human heterodimeric IL-15/lactadherin complexes. J Extracell Vesicles 7(1):1442088 6. Watson DC et al (2016) Efficient production and enhanced tumor delivery of engineered

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extracellular vesicles. Biomaterials 105:195–205 7. Mitchell JP et al (2008) Increased exosome production from tumour cell cultures using the Integra CELLine culture system. J Immunol Methods 335(1–2):98–105 8. Guerreiro EM et al (2018) Efficient extracellular vesicle isolation by combining cell media modifications, ultrafiltration, and sizeexclusion chromatography. PLoS One 13(9): e0204276 9. Hisey CL et al (2020) Towards establishing extracellular vesicle-associated RNAs as biomarkers for HER2+ breast cancer. F1000Res 9:1362 10. Palviainen M et al (2019) Metabolic signature of extracellular vesicles depends on the cell culture conditions. J Extracell Vesicles 8 (1):1596669 11. Jeppesen DK et al (2014) Quantitative proteomics of fractionated membrane and lumen exosome proteins from isogenic metastatic and nonmetastatic bladder cancer cells reveal differential expression of EMT factors. Proteomics 14(6):699–712 12. Griffiths SG et al (2017) Differential proteome analysis of extracellular vesicles from breast cancer cell lines by chaperone affinity enrichment. Proteomes 5(4):25 13. Ji H et al (2014) Deep sequencing of RNA from three different extracellular vesicle (EV) subtypes released from the human

LIM1863 colon cancer cell line uncovers distinct miRNA-enrichment signatures. PLoS One 9(10):e110314 14. Chen M et al (2019) Distinct shed microvesicle and exosome microRNA signatures reveal diagnostic markers for colorectal cancer. PLoS One 14(1):e0210003 15. Thery C et al (2018) Minimal information for studies of extracellular vesicles 2018 (MISEV2018): a position statement of the International Society for Extracellular Vesicles and update of the MISEV2014 guidelines. J Extracell Vesicles 7(1):1535750 16. Lobb R, Moller A (2017) Size exclusion chromatography: a simple and reliable method for exosome purification. In: Kuo WP, Shidong J (eds) Extracellular vesicles: methods and protocols. Humana Press, Totowa, New Jersey, pp 105–110 17. Cizmar P, Yuana Y (2017) Detection and characterization of extracellular vesicles by transmission and cryo-transmission electron microscopy. In: Kuo WP, Shidong J (eds) Extracellular vesicles: methods and protocols. Humana Press, Totowa, New Jersey, pp 221–232 18. Kowal EJK et al (2017) Extracellular vesicle isolation and analysis by western blotting. In: Kuo WP, Shidong J (eds) Extracellular vesicles: methods and protocols. Humana Press, Totowa, New Jersey, pp 143–152

Methods in Molecular Biology (2022) 2436: 193–204 DOI 10.1007/7651_2021_416 © Springer Science+Business Media, LLC 2021 Published online: 07 September 2021

Extracellular Vesicle Collection from Human Stem Cells Grown in Suspension Bioreactors Xuegang Yuan, Xingchi Chen, Changchun Zeng, David G. Meckes Jr, and Yan Li Abstract Extracellular vesicles (EVs) are particles with 100–1000 nm sizes which are secreted by cells for intercellular communication. Meanwhile, studies have found that EVs secreted by human stem cells carry similar characteristics (microRNAs, proteins, metabolites, etc.) from their cell counterpart. Thus, EVs derived from stem cells, especially human induced pluripotent stem cells (hiPSCs) and human mesenchymal stromal/stem cells (hMSCs) are promising candidates for cell-free therapy. However, conventional planar culture is insufficient to produce a large amount of cells or EVs to satisfy clinical requirements. In this chapter, we described feasible approaches to harvest EVs secreted by lineage-specific hiPSCs and undifferentiated hMSCs in suspension bioreactors. Differentiation of hiPSCs to cortical organoids can be performed in suspension bioreactors and the corresponding EVs can be isolated and purified. This scale-up protocol can be applied to a majority of stem cell types with EV collection thus provides useful information for both experimental and biomanufacturing purposes. Key words Biomanufacturing, Differential centrifugation, Extracellular vesicles, Human stem cells, Suspension bioreactors

1

Introduction In past decades, human stem cells including pluripotent stem cells and adult multipotent stem cells have drawn significant attentions pre-clinically and clinically [1]. Human induced pluripotent stem cells (hiPSCs) exhibit robust differentiation potential for modeling of disease pathology, drug discovery, and act as potential cell sources for therapeutic applications [2]. On the other hand, human mesenchymal stromal/stem cells (hMSCs) have been widely acknowledged for their regenerative potentials mediated by their secretome and paracrine effects. Studies have demonstrated promising strategies by using both types of stem cells to understand disease progression and therapeutic mechanisms, as well as in preclinical/clinical trials following biomanufacturing regulations [3, 4]. Recently, extracellular vesicles (EVs) or exosomes, the nano-

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particles secreted by cells, have been shown to carry specific cellular characteristics from the original cells and thus could represent lineage commitment or exhibit potential therapeutic effects [5]. EVs generated from undifferentiated hiPSCs carry specific cargos indicating the difference from the original cells being reprogrammed [6]. Potentially, EVs from hiPSCs exert homeostatic regulation on stressed human umbilical vein endothelial cells (HUVECs) to maintain cell viability and reduce senescence [7]. Moreover, EVs from hiPSC-derived neural progenitor cells (hiPSC-NPCs) not only carry lineage-specific information but also exhibit pro-neurogenesis and circuit assembly potentials [8, 9]. For hMSCs, there is an increasing body of evidence indicating the secreted EVs capture the therapeutic potentials of hMSCs and remain biologically active after transplantation [10– 12]. Human umbilical cord MSC-secreted EVs have been shown to exert liver protection under culture stress and ameliorate autoimmune symptoms via immunomodulation in rodent uveoretinitis models [13, 14]. Clinical trials have approached to study steroid refractory graft-versus host disease and grade III-IV chronic kidney disease patients with hMSC-EV administrations [15, 16]. Two-dimension (2D) conventional culture of human stem cells is sufficient and robust in general laboratory scale. However, industrial or clinical applications require a large amount of cells as well as stem cell derivatives beyond what the 2D culture can provide. For instance, 2–8  106 cells/kg patient weight would be needed for graft-versus-host diseases in stem cell based therapy [17]. To fulfill the clinical requirement, three-dimensional (3D) suspension bioreactors are designed for biomanufacturing of human stem cells and their derivatives. Microcarriers have been applied for anchoragedependent cells such as hMSCs. Studies have reported up to 43-fold increase of hMSCs in a 50 L stir-tank bioreactor [18]. Moreover, advanced bioreactors, such as wave bioreactor and PBS vertical wheel (PBS-VW) bioreactors have been designed to reduce shear stress to maintain hMSC homeostasis while still provide rapid expansion [19]. With the success of cell expansion in bioreactors, stem cell derivatives such as EVs should also be able to be scaled up for biomanufacturing purpose, though not many cases have been published. In this protocol chapter, detailed procedures and handling notes are described for EV isolation from hiPSC-NPCs and human umbilical cord-derived MSCs (hUC-MSCs) grown in suspension bioreactors. For hiPSC-NPC differentiation and expansion, a 50 mL spinner flask is selected. For hUC-MSC expansion, Cytodex-1 microcarrier and a 100 mL PBS-VW bioreactor is selected. EV-free medium preparation and the downstream EV isolation is modified and described based on our previous studies [20–22].

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Materials

2.1 Materials for hiPSC Differentiation in Planar and Bioreactor Cultures

1. A frozen hiPSC line, iPSK3 cells, was kindly provided by Dr. Stephen Duncan from Medical College of Wisconsin [23]. Briefly, the cell line was derived from human foreskin fibroblasts with transfection of plasmid DNA encoding reprogramming factors: OCT4, NANOG, SOX2, and LIN28 (see Note 1). 2. Prior to hiPSC culture, Lactose Dehydrogenase Elevating Virus (LDEV)-Free Reduced Growth Factor Basement Membrane Matrix Geltrex (Life Technologies, #A1413202) is thawed under 4  C overnight, then aliquoted and stored under 20  C. Geltrex is diluted in cold Dulbecco’s Modified Eagle’s medium (DMEM, Life Technologies, Carlsbad, CA) at 1:100 dilution (1%) and stored under 20  C. Tissue culture surface is coated with 1% Geltrex solution for overnight in 37  C incubator (see Note 2). 3. hiPSC complete culture medium (hiPSC-CCM) is prepared by adding 20% serum-free mTeSR™1 5 supplement (StemCell™ Technologies Inc., #85852) in mTeSR™1 basal medium (StemCell™ Technologies Inc., #05850). The hiPSC-CCM can be stored under 4  C. To dissociate hiPSCs for passaging, Accutase solution (VWR International, Radnor, PA., #10210-214) is used for obtaining single cell suspension. Aliquoted Accutase solution is stored under 20  C. Rho-associated kinase (ROCK) inhibitor Y27632, 2 mg or 10 mM in 624.4 μL dimethyl sulfoxide (DMSO, #MD0025), is purchased from iXCells Biotechnologies (San Diego, CA). The final working concentration is 10 μM by adding 1 μL of the stock solution per mL of hiPSC-CCM (see Note 3). 4. Human NPC differentiation medium is composed of DMEMF12 (Gibco™, #125000-062), with 2% B27 serum-free supplement (50, Life Technologies, #17504044) and stored under 4  C. Dual-SMAD inhibition for neural differentiation of hiPSCs is achieved by two small molecules: SB431542 (Sigma, #S4317) and LDN193189 (Sigma, #SML0559), both dissolved in DMSO and stored under 20  C. The working concentrations in differentiation medium are 10 μM SB431542 and 100 nM LDN193189 (see Note 4).

2.2 Materials for hMSC Expansion in Planar and Bioreactor Cultures

1. Frozen hMSC lines derived from human umbilical cords are provided by SynergyBiologics (Tallahassee, FL). hMSCs at passage 0 (P0) are cryopreserved and stored in liquid nitrogen. 2. hMSC complete culture medium (hMSC-CCM) is prepared by dissolving 10.08 g Minimum Essential Medium Alpha Medium

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(Life Technologies, #12000-063), 2.2 g sodium bicarbonate (Sigma-Aldrich, #S5761), 10 mL penicillin–streptomycin (Life Technologies, #15070-063), and 100 mL fetal bovine serum (FBS, Atlanta Biologicals, Inc. #S11110) in 1 L deionized water; then filtered by 0.2 μm pore size filter in sterile bottle and stored under 4  C for future use. The 150 mm diameter tissue culture petri dishes are obtained from VWR International (#25382-442). 0.25% trypsin/EDTA solution is from ThermoFisher Scientific (#25200056). 2.3 Bioreactor Preparation

1. A lab scale of 50 mL glass spinner flask bioreactor (Wheaton, #356875) is used for hiPSC-NPC differentiation, as well as medium collection for EV isolation. Prior to culture, the glass vessel is coated with 1 mL sigmacote (Sigma-Aldrich, #SL2) and then dried overnight at room temperature. The spinner flask is autoclaved for future use. 2. For PBS-VW bioreactor, PBS mini system is purchased from PBS Biotech™, Inc. (Camarillo, CA). This system includes magnetic agitation base and a sterile single-use vessel in 100 mL. The system is assembled and placed in standard cell culture 37  C incubator with 5% CO2. Cytodex 1 microcarriers (GE Healthcare Life Sciences, #17-0448-01) are prepared by hydrating the microcarriers in phosphate buffered saline (PBS) for overnight and washed twice with PBS before autoclave. Then the microcarriers are washed again with PBS and ready to use.

2.4 Materials for EV Isolation from Human Stem Cells Grown in Bioreactor Cultures

3

1. EV-free FBS is prepared by ultracentrifuge. FBS is spun at 4  C, 100,000  g for 20 h. The supernatant is carefully collected as EV-free FBS for EV collection. 2. All centrifuges are pre-cooled to 4  C during EV isolation and purification. Polyethylene glycol 6000 (PEG 6000, VWR International, Radnor, PA., #80503) solution is prepared by mixing 160 g PEG6000 with 1 M sodium chloride (NaCl, VWR International, #470302) in 1 L milli-Q water, then filtered with 0.2 μm pore size filter, resulting in 16% PEG6000 solution. EV-free PBS is prepared by double-filter of sterile PBS for future use (see Note 5).

Methods

3.1 Culture and Expansion of hiPSCs in Planar Culture

1. Frozen iPSK3 cells are recovered by immediately thawing in a 37  C water bath for 30 s until a small piece of ice remains. Spray the cryopreserve vial with 70% ethanol and open the vial in biological safety cabinet. Transfer the cell suspension carefully into at least ten times of volume of hiPSC-CCM (for

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instance, 1 mL cell suspension in 10 mL CCM) in a centrifuge tube. Gently pipette the mixture and then centrifuge at 300  g for 5 min (see Note 6). 2. After centrifugation, carefully remove the supernatant and do not disturb the cell pellet. Resuspend the cell pellet with 1–3 mL hiPSC-CCM carefully and distribute the cell suspension onto Geltrex-coated culture surface at 1–2  105 cells/ cm2. ROCK inhibitor Y27632 is added at 10 μM in the media. The recovered hiPSCs are cultured in a standard incubator (37  C, 5% CO2). 3. First medium change is performed 24-h later to remove ROCK inhibitor. Cell attachment can be visualized under microscope. hiPSC culture is maintained in CCM and medium is replaced every 2–4 days. When cells are compacted and reach a high density, medium change is performed daily. 4. For passaging hiPSCs, culture medium is removed, and the cells are washed with sterile PBS. Then the cells are incubated with Accutase solution at 37  C for 5 min. Gently pipet the mixture to acquire single cell suspension. Transfer the mixture to a 15 mL centrifuge tube, wash the culture surface once with hiPSC-CCM, and transfer the medium to the centrifuge tube (see Note 7). 5. Spin down the cells at 300  g for 5 min and remove the supernatant. Resuspend the cell pellet with hiPSC-CCM and determine the cell number by hemocytometer. Passaged hiPSCs can be expanded on Geltrex-coated surface or prepared for NPC differentiation in bioreactors. 3.2 Differentiation of NPC Organoid from hiPSCs in Spinner Flasks

1. hiPSC suspension is collected and seeded in the 15 mL spinner flask at 4–5  105 cells/mL in NPC differentiation medium containing 10 μM Y27632. The bioreactor is set up on a programmable magnetic stirrer (Wheaton, #900701) and the whole system is placed in a standard culture incubator (37  C, 5% CO2). 2. In initial aggregation phase (day 0), intermittent agitation is used after cell seeding in the bioreactor. The stirrer is set to 80 rpm for 15 min and off for 15 min for a total of 10 cycles. Then the agitation speed is set to 80 rpm for the rest of the culture. 3. At day 1, stop the agitation and let the hiPSC aggregates to settle down at the bottom; carefully remove the medium by pipette and resuspend the aggregates with fresh NPC differentiation medium containing 10 μM SB431542 and 100 nM LDN193189 to induce neural lineage commitment. Restart the agitation and culture.

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4. Medium sample can be collected every day for the analysis of glucose and lactate concentration by YSI2950 biochemical analyzer. Half medium change is performed every 2 days with fresh NPC differentiation medium containing 10 μM SB431542 and 100 nM LDN193189. 5. On day 8, NPC aggregates or spheres can be collected for the evaluation of neural progenitor markers. Both SB431542 and LDN193189 are removed by medium change and fresh NPC differentiation medium is added to the bioreactor for continuous agitation culture (see Note 8). 6. For EV collection, conditioned medium collection can be performed by defined time interval, for example, every 2 days. Cultures can be maintained in this manner for a very long time (25–70 days) (Fig. 1a). Collected media are preserved under 4  C for further processing. A

DAY 25

B

DAY 27

C hiPSC-NPC EV size 250

EV size (nm)

200 150 100 50 0 Mean

Mode

Fig. 1 (a) hiPSC-NPC organoid growth in spinner flask at day 25 and day 27 under optical light microscope. Scale bar: 400 μm. (b) Transmission electron microscopy (TEM) images of EVs from hiPSC-NPC bioreactor culture. Scale bar: 200 nm. (c) The size of hiPSC-NPC secreted EVs determined by nanoparticle tracking analysis (NTA)

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1. One vial of frozen hMSCs (generally contains 1  106 cells) is recovered by immediately thawing in a 37  C water bath for 30 s until a small piece of ice remains. Spray the vial with 70% ethanol and open the vial in biological safety cabinet. Transfer the cell suspension carefully into at least ten times of volume of hMSC-CCM (for instance, 1 mL cell suspension in 10 mL CCM) in centrifuge tube. Gently pipette the mixture and then centrifuge at 400  g for 5 min. 2. After centrifugation, carefully remove the supernatant and do not disturb the cell pellet. Resuspend the cell pellet with 1–3 mL hMSC-CCM carefully and distribute the cell suspension into 150 mm diameter petri dish at 1200–1500 cells/cm2. The culture is maintained in a standard incubator (37  C, 5% CO2). 3. First medium change is performed after 12 h to remove debris and unattached cells. Then medium is changed every 2 days with fresh hMSC-CCM. 4. For passaging hMSCs, culture medium is removed, and the cells are washed with sterile PBS twice. Then the hMSCs are treated with 0.25% trypsin solution at 37  C for 5–7 min. Trypsin is neutralized by adding hMSC-CCM and detached cells are collected in a 15 mL centrifuge tube. The petri dish is washed with fresh hMSC-CCM once to collect residue cells for maximized yield. 5. Spin down the hMSCs at 500  g for 5 min and remove the supernatant. Resuspend the cell pellet with hMSC-CCM and determine the cell number by hemocytometer. hMSCs can be expanded on tissue culture surface or on microcarriers in PBS-VW bioreactors for EV collection (see Note 9).

3.4 Expansion of hMSCs in PBS-VW Bioreactors

1. hMSCs in planar cultures are harvested and resuspended with hMSC-CCM containing EV-free FBS. Then hMSCs are mixed with sterile Cytodex-1 microcarriers (0.25–0.5 g) at the density of 1000–1500 cells/cm2. The mixture is added to the PBS-VW bioreactors and the volume is brought to 60 mL with hMSCCCM containing EV-free FBS (see Note 10). 2. On day 0, intermittent agitation is used after cells and microcarriers are transferred into the bioreactor. The agitation base is set to 25 rpm for 5 min and off for 15 min for a total of 12 cycles (roughly 4 h). Then the agitation speed is set to 25 rpm for the rest of the culture period. The total volume is remained at 60 mL for seeding phase. After seeding, the total volume is brought to 100 mL with fresh hMSC-CCM containing EV-free FBS. The culture is maintained in a standard incubator (37  C, 5% CO2).

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A

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Fig. 2 (a) hUC-MSCs growth in PBS-VW bioreactors at day 1 and day 7, stained by Hoechst 33342 for cell nuclei and visualized under fluorescent microscope. Scale bar: 200 μm. (b) Transmission electron microscopy (TEM) images of EVs from hUC-MSC bioreactor culture. Scale bar: 100 nm. (c) The size of hUC-MSC secreted EVs determined by nanoparticle tracking analysis (NTA)

3. Sampling of the bioreactor can be performed every day by taking 1 mL of mixture from PBS-VW bioreactor under agitation. Cells on microcarriers can be visualized by nucleus staining (Fig. 2a) and culture medium supernatants can be used for glucose/lactate measurement. 4. Medium collection is performed every 2 days with 50% fresh medium change. The agitation is stopped to let cells/microcarriers to settle down. 50 mL media are collected from the PBS-VW bioreactor. Then 50 mL of pre-warmed EV-free hMSC-CCM is added to the bioreactor and agitation is started again. Collected medium is ready for EV isolation.

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1. Collected media can be stored at 4  C for 1 week before processing, otherwise they should be stored at 50  C. Collected media, both from hiPSC-NPC organoid culture and hMSC bioreactor culture, undergo modified differential centrifugation at 4  C: 500  g for 5 min, 2000  g for 10 min, and 10,000  g for 30 min. Supernatants are collected from each step (see Note 11). 2. For hiPSC-NPC organoid conditioned media, 10% EV-free FBS and 16% PEG6000 solution are added to the medium to a final concentration of 8% PEG6000. For hMSC conditioned media, only 16% PEG6000 solution is added to the media at a final concentration of 8% PEG6000. All samples are well mixed and stored at 4  C for 12–24 h. 3. Next, medium samples are centrifuged at 3200  g for 60 min at 4  C to collect PEG-enriched EV pellets. The supernatants are discarded, and the pellets are resuspended in sterile, EV-free PBS for washing out PEG 6000, then ultracentrifuge is performed at 120,000  g for 70 min at 4  C. 4. After ultracentrifuge, supernatants are discarded and the EV pellets are resuspended in sterile, EV-free PBS (usually at 100 μL, but the volume can vary). The morphology of the isolated EVs can be examined by transmission electron microscopy (Figs. 1b and 2b). EV size distribution and concentrations can be determined by Nanoparticle Tracking Analysis (NTA) using a Malvern NanoSight LM10 instrument (Figs. 1c and 2c) (see Note 12). 5. Stem cell-secreted EVs are aliquoted and stored at 80  C for further downstream analysis, culture experiments, or animal transplantation.

4

Notes 1. For long time storage, cryopreserved hiPSCs should be stored in liquid nitrogen. 2. Proper Geltrex coating is critical for hiPSC recovery and expansion. Geltrex coating solution should be well mixed in cold medium before applied to culture surface. 3. mTeSR™1 medium should be properly prepared and stored following the manufacturer’s instruction. The hiPSCs are very sensitive to the quality of the medium. Degradation in both basal medium and 5 supplement would result in cell detachment and spontaneous differentiation. 4. Differentiation protocols can be modified to reflect the purpose of neural lineage commitment. Small molecules treatment can be modified to generate neural cells with specific brain

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region identity or specific neuronal subtypes. For example, Wnt inhibition can be applied for generating dorsal forebrain neural cells and sonic hedgehog activation can be used for enriching ventral forebrain cells. 5. PEG6000 solution is critical for EV isolation via the modified extraPEG enrichment method, and thus should be prepared properly. Besides 16% PEG6000 solution, 24% PEG6000 solution can also be prepared and then the medium:24% PEG6000 solution would be 2:1 to reach a final concentration of 8% PEG6000 medium mixture. 6. DMSO is a general cryopreservation agent, and is harmful for cells at room temperature. Thus, the thawing and recovery process of hiPSCs should be rapid. Moreover, hiPSCs are very fragile post-thaw and should be taken care in caution, e.g., pipetting of cells should be gentle. 7. Passaging hiPSCs with Accutase should be closely monitored. hiPSC colonies are detached during this process and gentle pipetting may be required to achieve a single cell suspension. Treatment time should be minimized to reduce the damage to the cells. 8. hiPSC-NPC organoids can be collected and replate on the Geltrex-coated surface for further culture. Neuronal network can be observed after replating and culturing. Meanwhile, aggregates or spheroids can be dissociated by enzyme to generate single cell suspension for further analysis by flow cytometry. 9. It is recommended to wash hMSCs with PBS several times before the bioreactor culture using EV-free medium to minimize the EV contamination from the previous culture. 10. During initial seeding phase, the medium volume is reduced to increase the possibility of hMSCs contacting with microcarriers for better attachment. The volume can be adjusted as long as it covers the vertical wheel. 11. It is recommended that collected media should be stored at 4  C and processed within 1 week. If immediate processing is not available, media should be centrifuged at 4  C by 500  g for 5 min, 2000  g for 10 min, and then stored at 50  C. 12. After ultracentrifugation, EV pellets can be visualized at the bottom of the centrifuge tube. It is recommended to circle the pellets with marker and resuspend the EV pellets to visualize their complete dissolution. Alternatively, EV pellets can be directly lysed for protein or microRNA cargo analysis. Sterile, EV-free milli-Q water can be used to resuspend EVs as well.

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Acknowledgments This work was supported by National Science Foundation (Nos. 1652992 and 1743426) as well as partially by the National Institutes of Health (NIH; R01NS102395). The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH. References 1. Zomer HD, Vidane AS, Gonc¸alves NN, Ambro´sio CE (2015) Mesenchymal and induced pluripotent stem cells: general insights and clinical perspectives. Stem Cells Cloning 8:125–134 2. Rowe RG, Daley GQ (2019) Induced pluripotent stem cells in disease modelling and drug discovery. Nat Rev Genet 20:377–388 3. Yuan X, Rosenberg JT, Liu Y, Grant SC, Ma T (2019) Aggregation of human mesenchymal stem cells enhances survival and efficacy in stroke treatment. Cytotherapy 21:1033–1048 4. Yuan X, Logan TM, Ma T (2019) Metabolism in human mesenchymal stromal cells: a missing link between hMSC biomanufacturing and therapy? Front Immunol 10:977 5. The´ry C, Witwer KW, Aikawa E, Alcaraz MJ, Anderson JD et al (2018) Minimal information for studies of extracellular vesicles 2018: a position statement of the International Society for Extracellular Vesicles and update of the MISEV2014 guidelines. J Extracell Vesicles 7:1535750 6. Ding Q, Sun R, Wang P, Zhang H, Xiang M et al (2018) Protective effects of human induced pluripotent stem cell-derived exosomes on high glucose-induced injury in human endothelial cells. Exp Ther Med 15:4791–4797 7. Skamagki M, Zhang C, Ross CA, Ananthanarayanan A, Liu Z et al (2017) RNA exosome complex-mediated control of redox status in pluripotent stem cells. Stem Cell Rep 9:1053–1061 8. Hicks DA, Jones AC, Corbett NJ, Fisher K, Pickering-Brown SM et al (2020) Extracellular vesicles isolated from human induced pluripotent stem cell-derived neurons contain a transcriptional network. Neurochem Res 45:1711–1728 9. Park K-S, Bandeira E, Shelke GV, L€asser C, Lo¨tvall J (2019) Enhancement of therapeutic potential of mesenchymal stem cell-derived extracellular vesicles. Stem Cell Res Ther 10:1–15

10. Sharma P, Mesci P, Carromeu C, McClatchy DR, Schiapparelli L et al (2019) Exosomes regulate neurogenesis and circuit assembly. Proc Natl Acad Sci U S A 116:16086–16094 11. Clark K, Zhang S, Barthe S, Kumar P, Pivetti C et al (2019) Placental mesenchymal stem cellderived extracellular vesicles promote myelin regeneration in an animal model of multiple sclerosis. Cell 8:1497 12. Li T, Yan Y, Wang B, Qian H, Zhang X et al (2013) Exosomes derived from human umbilical cord mesenchymal stem cells alleviate liver fibrosis. Stem Cells Dev 22:845–854 13. Witwer KW, Van Balkom BW, Bruno S, Choo A, Dominici M et al (2019) Defining mesenchymal stromal cell (MSC)-derived small extracellular vesicles for therapeutic applications. J Extracell Vesicles 8:1609206 14. Bai L, Shao H, Wang H, Zhang Z, Su C et al (2017) Effects of mesenchymal stem cellderived exosomes on experimental autoimmune uveitis. Sci Rep 7(1):4323 15. Kordelas L, Rebmann V, Ludwig A, Radtke S, Ruesing J et al (2014) MSC-derived exosomes: a novel tool to treat therapy-refractory graftversus-host disease. Leukemia 28:970–973 16. Nassar W, El-Ansary M, Sabry D, Mostafa MA, Fayad T et al (2016) Umbilical cord mesenchymal stem cells derived extracellular vesicles can safely ameliorate the progression of chronic kidney diseases. Biomater Res 20:21 17. Kebriaei P, Isola L, Bahceci E, Holland K, Rowley S et al (2009) Adult human mesenchymal stem cells added to corticosteroid therapy for the treatment of acute graft-versus-host disease. Biol Blood Marrow Transplant 15:804–811 18. Lawson T, Kehoe DE, Schnitzler AC, Rapiejko PJ, Der KA et al (2017) Process development for expansion of human mesenchymal stromal cells in a 50 L single-use stirred tank bioreactor. Biochem Eng J 120:49–62 19. de Sousa PD, Bandeiras C, de Almeida FM, Rodrigues CA, Jung S et al (2019) Scalable manufacturing of human mesenchymal stromal

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cells in the vertical-wheel bioreactor system: an experimental and economic approach. Biotechnol J 14:1800716 20. Rider MA, Hurwitz SN, Meckes DG Jr (2016) ExtraPEG: a polyethylene glycol-based method for enrichment of extracellular vesicles. Sci Rep 6:23978 21. Cone AS, Hurwitz SN, Lee GS, Yuan X, Zhou Y et al (2020) Alix and Syntenin-1 direct amyloid precursor protein trafficking into extracellular vesicles. BMC Mol Cell Biol 21:1–20

22. Marzano M, Bou-Dargham MJ, Cone AS, York S, Helsper S et al (2021) Biogenesis of extracellular vesicles produced from humanstem-cell-derived cortical spheroids exposed to iron oxides. ACS Biomater Sci Eng 7:1111–1122 23. Si-Tayeb K, Noto FK, Sepac A, Sedlic F, Bosnjak ZJ et al (2010) Generation of human induced pluripotent stem cells by simple transient transfection of plasmid DNA encoding reprogramming factors. BMC Dev Biol 10:1–10

Methods in Molecular Biology (2022) 2436: 205–222 DOI 10.1007/7651_2021_417 © Springer Science+Business Media, LLC 2021 Published online: 10 September 2021

Bacterial Nanocellulose-Based Grafts for Cell Colonization Studies: An In Vitro Bioreactor Perfusion Model Max Wacker, Jan Riedel, Priya Veluswamy, Maximilian Scherner, Jens Wippermann, Heike Walles, and Jo¨rn Hu¨lsmann Abstract With the aging population, the demand for artificial small diameter vascular grafts is constantly increasing, as the availability of autologous grafts is limited due to vascular diseases. A confluent lining with endothelial cells is considered to be a cornerstone for long-term patency of artificial small diameter grafts. We use bacterial nanocellulose off-the-shelf grafts and describe a detailed methodology to study the ability of these grafts to re-colonize with endothelial cells in an in vitro bioreactor model. The viability of the constructs generated in this process was investigated using established cell culture and tissue engineering methods, which includes WST-1 proliferation assay, AcLDL uptake assay, lactate balancing and histological characterization. The data generated this straight forward methodology allow an initial assessment of the principal prospects of success in forming a stable endothelium in artificial vascular prostheses. Keywords Bacterial cellulose, Cell seeding, Endothelialization, Perfusion bioreactor, Small diameter vascular grafts, Vascular tissue engineering

1

Introduction Due to the aging population and the resulting limited availability of autologous vessels, there is an increasing demand for small caliber artificial grafts in the fields of cardiac and vascular surgery [1]. The development of small caliber vascular grafts poses a challenge to prosthesis developers. The patency rate of the grafts remains to be the main criterion for their successful application in vivo, as this directly influences morbidity and survival in the use, for e.g., coronary artery bypass graft [2]. The failure of grafts when implanted in vivo is especially characterized by two mechanisms; (1) narrowing of the lumen by intimal hyperplasia and (2) thrombus formation as a consequence of graft surface and blood interactions [3]. In vivo, the local regulation of these processes is mainly regulated by the vascular endothelium [4]. Therefore, the process of endothelialization is subject of current research in vascular tissue engineering and

Max Wacker and Jan Riedel contributed equally to this work.

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is considered to be pivotal for acceptable long-term patency rates of small diameter vascular grafts [5, 6]. Especially for acellular vascular grafts that exhibit on-site endothelialization after implantation, the preclinical evaluation of their re-endothelialization potential is of utmost importance to avoid graft failure related complications. But while well-established small and large animal models for vascular prostheses are already available for preclinical studies [7], further establishment of in vitro methodology that targets to estimate endothelialization processes for vascular grafts is inevitable, as evidenced by the Three Rs principles for the ethical use of animals for scientific investigations [8]. For the evaluation of acellular grafts, which are directly introduced into a complex physiological environment after implantation, dynamic cell culture models are more plausible than the static cell culture models, because cultivation under flow generated shear stress simulates in vivo conditions resulting in a more functional endothelium [9]. To provide such physiological stimulations, state of the art bioreactor systems are available to study small caliber vascular grafts under perfusion and pulsatile flow in vitro [10]. They usually consist of two chambers, providing intraluminal pulsatile flow to provide culture medium and shear stress for the endothelial cells, and a surrounding container, that can be filled with smooth muscle cells and cell culture medium, if necessary [11]. More complex bioreactor systems include pressure and flow measurement, biosensors for surveillance of nutrient content and metabolites in the medium and rotational parts to overcome gravitational effects during the cultivation period [12]. However, the whole complexity of the in vivo situation for the aspired clinical application cannot be mimicked at once in bioreactor setups in vitro. Especially if artificial biomaterials are used for the scaffolds, additional questions, e.g., regarding the initial cell adhesion need to be addressed in more simplified setups. Accordingly, approaches like static or low flow seeding and pre-cultivation techniques that were developed for the generation of cellular grafts needs to be considered in preparative steps prior to bioreactor-assisted cultivation [9–11]. The comparability between the many different approaches existing today might therefore be impacted and most importantly, the scientific question to be answered might differ with regard to acellular scaffolds. Acellular scaffolds are thought to be repopulated by host cells onsite after implantation in vivo. Regarding the initial adherence, it should be noted that even hours after seeding of the cells, only low flow rates should be applied to prevent a washout of the seeded cells [13]. But, the medium flow in the system should be high enough to wash off non-adherent cells, while applying detectable shear stress to the cells. Here, we present a protocol to estimate the homing of endothelial cells in artificial bacterial cellulose based acellular vascular grafts and their physiological performance in a perfusion bioreactor

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system after rotational cell seeding, followed by low flow mediaperfusion for 5 days. The setup allows medium sampling and describes the implementation of intraluminal cell proliferation and functional assays to determine endothelial cell function and the histological characterization of the seeded cell constructs. Next to the 3D printed bioreactors, the system consists of common laboratory equipment such as pumps, filters and Luer Lock connectors and is therefore easy to set up by common lab equipment. It should be noted that today 3D printing has become an affordable methodology that can be applied in many laboratories. The above-mentioned methods can easily be adopted to tubular vascular grafts with other dimensions by printing customized bioreactors. Thus, we describe an efficient, reliable, and affordable method for evaluating the initial adherence and proliferation of endothelial cells on tubular, acellular vascular grafts under in vitro dynamic conditions.

2 2.1

Materials Bioreactor Setup

The applied bioreactor was designed as a straight perfusion chamber with two barb connectors for direct connection of the tubing (from the outside) to the perfused grafts (on the inside) and recirculation of leaking medium. 1. Bioreactor chamber (see Note 1). 2. Tubing: (a) 1 pump hose (PharMed BPT). (b) Silicon tubing (inner diameter 3 mm). (c) Female Luer Lock Style to barb connector (according to used tubing diameters). (d) Male Luer Lock Style to barb connector (according to used tubing diameters). (e) Y-connector 3–5 mm. (f) Three-way stopcock. (g) Sealing caps. 3. Syringes: (a) 10 ml syringe with Luer Lock fitting. (b) 50 ml syringe with Luer Lock fitting. 4. Reservoir bottle with three inlets and outlets for media transport (see Note 2). We used a total of 50 ml of media for approximately 2.5 cm2 of reseeded luminal surface area and chose a 100 ml Schott Duran flask for reservoir.

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5. Filter. (a) 0.2 μm injection filter (e.g., B. Braun, Intrapur®Neonat). (b) 0.2 μm sterile filter for gas exchange and pressure compensation, connectable to Luer lock fittings. 6. Small diameter vascular grafts to be studied, inner diameter 5.0 mm. Here, we used bacterial nanocellulose small diameter vascular grafts with and without fibronectin coating. 2.2 Assays and Staining Solutions

1. WST-1 reagent (Roche, Mannheim, Germany, cat. nr.: 05 015 944 001). 2. Alexa Fluor 488 AcLDL (Thermofisher, Carlsbad, USA, cat. nr.: L23380). 3. Rhodamine Phalloidin Reagent (Abcam, Cambridge, USA, cat. nr.: ab235138). 4. NucBlue™ Live Ready Probes™ Reagent (Invitrogen, Carlsbad, USA, cat. nr.: R37605). 5. Acridine orange (Invitrogen, Carlsbad, USA, cat. nr.: A1301).

2.3

Cell Culture

1. Endothelial cell line (here, we used rabbit endothelial progenitor cells (rbEPC), isolated from Chinchilla bastard rabbit blood or human saphenous vein endothelial cells (HSVEC)). 2. Cell line specific cell culture medium containing 10% fetal calf serum. 3. Consumables for cell culture (flasks, conical tubes, 6-well plates, serological pipettes, detachment reagent).

2.4

Antibodies

1. CD31/PECAM-1 antibody that fits to your cell line (here, we used CD31/PECAM-1 mouse anti-human antibody [R&D Systems, Minneapolis, USA, cat. nr.: BBA7]). 2. Secondary antibody that fits to your primary antibody (here, we used Donkey anti-mouse Alexa Fluro 488 [Jackson ImmunoResearch, West Grove, USA, cat. nr.: 715-545-152]).

2.5 Chemicals and Solutions

1. Phosphate buffered saline without calcium and magnesium (PBS). 2. PBS with calcium and magnesium (PBS+). 3. PBS with 1% penicillin-streptomycin cell culture supplement. 4. Penicillin-streptomycin cell culture supplement. 5. Normal donkey serum. 6. Paraformaldehyde (PFA) solution, 4%. 7. Bovine serum albumin (BSA).

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Consumables

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1. Micro reaction tubes (2 ml). 2. Reaction tubes (15 and 50 ml). 3. Pipettes and tips (0.1–2.5, 2–20, 10–100, 100–100 μl). 4. 96-Well plates, flat bottom, clear and 12-well plates with clear bottom, suitable for cell culture. 5. Microscope slides (e.g., SuperFrost plus microscope slides, Fisher Scientific, Schwerte, Germany, cat. nr.: 10149870).

2.7

Hardware

1. Bioreactor perfusion platform as described by Schuerlein et al. [14] (Customized from University Hospital Wu¨rzburg, Department for Tissue Engineering and Regenerative Medicine, Wu¨rzburg, Germany). As an alternative, a CO2 cell culture incubator with accessibility to a peristaltic pump and space for the bioreactor perfusion system could be used. 2. Set for bioreactor assembly: Sterile surgical gloves, coat and sheets, non-absorbable braided suture material (strength 2-0), surgical tweezers and needle holder. 3. Cell culture incubator with controlled CO2 atmosphere (5–10% according to requirements of used media) and temperature (37
C). 4. Sterile bench, large enough to assemble the perfusion system (see Fig. 1).

Fig. 1 Complete perfusion setup placed inside the incubator platform with controlled atmosphere and temperature. 1: Medium reservoir; 2: Medium sample syringes; 3: Roller pump; 4: 0.2 μm sterile filter for adding WST-1 cell proliferation assay reagent; 5: 3D printed reactor containing vascular grafts; 6: Attached syringes to collect WST-1 cell proliferation assay reagent

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5. Benchtop centrifuge with temperature control and swinging bucket rotor e.g., Allegra X-12R, Beckmann Coulter, Brea (CA), USA. 6. Roller mixer capable of slow rotation (here, we used 2 revolutions/30 min). 7. Fluorescence microscope (EVOS Auto 2, ThermoFisher Scientific, MA, USA). 8. Confocal microscope (Leica TCS SP8, Leica Microsystems GmbH, Germany). 9. Multimode plate reader (such as Tecan Infinite® 200 PRO).

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Methods

3.1 General Considerations

3.2

Bioreactor Setup

l

The presented method is used to determine the success of cell homing after seeding in artificial small diameter vascular grafts and subsequent evaluation of metabolic activity and preservation of basic endothelial cell functions over a short period of 5 days.

l

The needed solutions should be prepared freshly on the day of their usage.

l

All autoclavable parts should be autoclaved in advance to allow them to cool down before usage on the first day of the experiments.

l

The cells used for the experiments were harvested on the day of cell seeding, thus cultivation has to be started 3–5 days in advance, depending on the cell type.

l

In our experiments we used cells cultivated under static conditions as positive controls to determine the seeding efficiency on the grafts. Therefore, the cells were seeded out on day 1 of the experiment in a 12-well plate with the same density compared to cells seeded on the luminal side of the vascular grafts and cultivated for 24 h until the first WST-1 assay.

Prior to conducting any experiments, set up the bioreactor system as shown in Fig. 1 in an unsterile fashion to make sure that all parts are fitting and to practice the handling. Adjust the length of the silicon tubes to ensure that the system fits inside the bioreactor perfusion platform. When the setup has been established, autoclave the silicone tubings with the respective Luer Lock connectors, the medium reservoir bottles and the pump tubing. Disconnect the bioreactor and autoclave separately. Do not autoclave syringes, 3-way stopcocks and filters, as these are intended for single use and not autoclavable.

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As it is essential to maintain sterility, all following steps need to be carried out under a sterile cell culture bench (Laminar air flow). Wear sterile, surgical gloves and gown during experiments. Additionally, we spread out a sterile surgical sheet inside the cell culture bench. 3.3 Connection of Small Diameter Vascular Grafts to Bioreactor Chamber

1. Open the autoclaved bioreactor chamber. 2. Pull the first graft over the barb connectors inside the bioreactor chamber using tweezers. 3. Use non resorbable surgical sutures to tie in the tube (see Note 3). 4. Attach one 3-way stopcock to each graft outlet of the bioreactor chamber. 5. Attach a syringe filled with 5 ml of PBS to one 3-way stopcock. 6. Carefully fill tube with PBS, holding the bioreactor chamber in an upright position with the syringe connected to the bottom 3-way stopcock to allow the air to evacuate. Determine the volume needed to completely fill the lumen between the two 3-way stopcocks upstream and downstream of the graft with fluid. This is the volume needed for endothelial cell seeding and WST-1 assay. 7. Close the 3-way stopcock on the opposite side. 8. Carefully apply some pressure in order to check for leakage. 9. Repeat with the second graft following steps 2–8. 10. Close all open Luer Lock ports with sterile sealing cabs, close the bioreactor chamber and store it in an incubator until seeding of endothelial cells.

3.4

Cell Seeding

On the first day of the experiments, the endothelial cells of the respective endothelial cell line will be seeded to the luminal surface of the grafts. In advance, the cells should be pre-cultivated to a confluency level of about 80% before the commencement of experiment and harvesting for cell seeding. They can be harvested following standard cell culture protocols. 1. Prepare endothelial the cell suspension for seeding in a required concentration. Adjust the working concentration to a cell number per luminal surface area that fits your testing hypothesis as well as corresponding targeted cultivation time. It should be noted that the same normalized cell count should also be applicable to your static 2D controls (see Note 4). In our specific setup we needed 3 ml of volume for each graft and seeded with a density of 5  104 cells per cm2, thus the concentration of the cell suspension for seeding was set to 4  105 cells per ml.

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2. Aspirate cell suspension in a 10 ml syringe. 3. Carefully fill up the tubes inside the bioreactor with the cell suspension (see Note 5). 4. Place the bioreactor chamber on a roller mixer inside a cell culture incubator at 37
C. 5. Rotate for a total time that is suitable for first attachment of the used cells (e.g., in this setup, we rotated at 2.0 revolutions/ 30 min). 6. In the meantime, seed the positive controls in 12 well plates in triplicates, using the same normalized cell count as the cells seeded on the grafts. The positive controls will be used to determine the seeding efficiency as explained in Subheading 3.6. 7. Stop the rotation and incubate the bioreactor under static conditions for the time that affirms safe attachment of the used cells (in this setup, we chose 3.5 h) (see Note 6). 8. Subsequently, after 3.5 h of static cultivation, take the bioreactor chamber under the sterile bench. 9. Discard the remaining suspension (see Note 7). 10. Gently flush once with PBS to remove non-adherent cells. 11. Replace all 3-way stopcocks by new sterile ones. 12. Attach the chamber to the perfusion setup. 13. Assemble all remaining syringes, 3-way-connectors and filters. 14. Fill up the perfusion setup with the required volume of media for cultivation. Here, we used 50 ml of cell culture medium containing 10% fetal calf serum and 1% penicillin/streptomycin supplement. 15. Connect the system to the roller pump inside the perfusion platform. 16. Start the perfusion and adjust flow conditions according to the planned approach. Here, we used a flow rate of 1.5 ml/min with pulsatile flow, 0.5 Hz. 17. Make sure all remaining air bubbles are flushed towards the reservoir bottle. 18. Continue perfusion for the intended cultivation time. Here, we applied a total length of 5 days (i.e., 96 h). 3.5 Lactate Monitoring

An evaluation of lactate production was considered standard to estimate the cell metabolism in tissue engineered constructs, which is also commonly applied for the process accompanying analytics in bioreactor assisted cultivations since the emergence of tissue engineering field in the 1990s [15]. In particular, by a simple batchwise media perfusion, it can be easily integrated by sequential

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media sampling. As the metabolism of endothelial cells mainly relies on glycolysis and consequent lactate production [16], lactate was measured in the cell culture medium in order to gain insight into metabolic activity of seeded cells. Henceforth, 1 ml of cell culture medium is collected through the syringes attached to the medium reservoir, to measure the lactate concentration everyday. 1. Start with the first medium collection 30 min after starting media perfusion. 2. Take up 1 ml of media into the last 10 ml syringe connected to the media reservoir. 3. Close the corresponding 3-way stopcock so that the syringe is disengaged from the medium reservoir. 4. Flush 5 ml of air from the second last syringe connected to the media reservoir through the tubing inside the medium reservoir. This is done to ensure that no media is left in the sampling tube. 5. Disconnect the last syringe with the medium and store the sample at 80
C. 6. Repeat steps 2–5 each day of the experiment. 7. After completion of the experiment, thaw the media samples and measure the lactate levels. In this setup, a large scale analyzer was used for medical laboratories in collaboration with the local institute for clinical chemistry. Alternatively, e.g., colorimetric or fluorometric assays are commercially available. 8. The acquired concentration was converted into an absolute molar mass in consideration of the media volume at the time of sampling. Exemplary results are shown in Fig. 2. 3.6 WST-1 Proliferation Assay

In recent past, WST-1 proliferation assay has been applied to quantitatively assess the cell proliferation in perfused tissue engineered constructs [17–19]. Another beneficial application comes by the ease to determine the seeding efficiency in the 3D artificial grafts in direct comparison to the optimal conditions of vertical sedimentation onto a two-dimensional cell culture surface in the positive controls. Especially as low efficiency rates for adhesion of seeded cells seems to be a problem in colonization of 3D scaffolds [20], we prefer a direct evaluation of the resulting vitalization over more laborious cell counting. We could see from preliminary experiments, that almost 100% of cells would attach and proliferate when seeded in a well plate. Therefore, the estimated seeding efficiency will be used as a measure for a ratio of cells that could attach to the tested grafts after 24 h relative to the positive controls in the wells.

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Fig. 2 Exemplary data of growth kinetics over the course of 96 h. Rabbit endothelial progenitor cells (rbEPC) were seeded on day one on either uncoated vs. fibronectin coated grafts made from bacterial nanocellulose. The proposed methods enabled us to observe distinct differences between the proliferative and metabolic activity of rbEPCs cultivated on the different graft types. (a) Cumulative lactate production over the course of 96 h. The data show that the cells seeded on fibronectin coated grafts produced more lactate compared to the uncoated grafts, indicating enhanced metabolic activity of the endothelial cells in the fibronectin group. (b) Metabolic activity as indicated by WST-1 assay. For both graft types used, the cells show growth over the time course of 96 h, but in the fibronectin group, the considerably higher growth activity indicates higher cell numbers attached to the graft in this group. (c) Calculated seeding efficiency from WST-1 proliferation assay data after 24 h. Compared to the uncoated group, the fibronectin group shows higher adherence of the seeded cells, reaching approximately 50% of the controls seeded under optimal conditions in a 12-well plate

The comparison between measured absolute metabolic activity after 24 and 96 h will allow for an estimation of the growth in the course of 96 h.

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The WST-1 assay solution has to be prepared in advance of the experiment. We used 12.5% of standard WST-1 working solution diluted in cell culture medium containing 10% FCS (see Note 8). 3.6.1 Perform WST-1 Assay (Modified from Manufacturer’s Instructions)

1. Take up 3 ml of WST-1 assay solution for each graft inside a 10 ml Luer Lock syringe. 2. Stop the perfusion. 3. Attach the syringe to one of the 0.2 μm infusion filters upstream to the graft. 4. Set all corresponding 3-way stopcocks to a position so that the assay solution can only flow inside the examined graft. 5. Carefully fill the lumen of the opened perfusion path with the assay solution. Take care about the remaining air, so that it will function as a watershed to push remaining medium out of the graft, and fill each graft with WST-1 solution until the watershed reaches the downstream 3-way stopcock. 6. Now close the upstream and distal 3-way-stopcock. 7. Let the assay incubate as recommended by the manufacturer and adjust the incubation time to the used cell type for up to 4 h. 8. Perform the WST-1 assay for the positive controls in the well plate. In order to norm WST-1 proliferation data to a positive control, the WST-1 assay has to be carried out with the same volume per seeded cells per surface area ratio as in the examined tubes. In our case we seeded 175,000 cells per well in a 12-well plate. The WST-1 assay was therefore carried out with 437 μl. 9. After 4 h remove the WST-1 solution out of the tubes by opening all 3-way-connectors so that you will be able to push out the assay solution with air out of the upstream attached syringe and collect the assay solution in the already attached downstream sampling syringe. 10. Open all 3-way connectors toward the perfusion system and restart the perfusion. 11. Measure the assay solution in a 96-well plate in duplicates for absorption at 440 nm and use 600 nm as reference wave length. 12. Calculate the WST-1 data for seeding efficiency as shown below. Exemplary results of seeding efficiency and cell proliferation are shown in Fig. 2.

Seeding efficiency ¼

ðAbsorbance at 440 nm  Absorbance at 600 nmÞsamples ðAbsorbance at 440 nm  Absorbance at 600 nmÞpositive controls

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3.7 Termination of Experiment

1. After 96 h of cultivation, stop the perfusion, take out the grafts and divide them into five equal samples by equidistant crosssectional cutting in order to perform the immune stainings and AcLDL assay. 2. Store the graft sections in PBS and immediately proceed with the assay and staining procedures. 3. The samples which are not used for AcLDL uptake assay should be washed in PBS (in 50 ml falcon tube) for 5 min, followed by fixation with in 4% PFA for 5 min (in 15 ml Ficoll tube). Wash them twice afterwards in an excess volume of PBS in a 50 ml falcon tube.

3.8 AcLDL Uptake Assay (Modified from Manufacturer’s Instructions)

The AcLDL assay is an easy and direct method to verify the intact functionality of the seeded endothelial cells by the uptake of acetylated low density lipoprotein (LDL) particles via scavenger receptors expressed on an functionality intact endothelial cell [21]. As a result, positive signals would indicate an unimpaired function of the endothelial cells, while the lack of LDL signals would indicate either impaired and degenerated endothelial cells or might represent a possible overgrowth by non-endothelial cells. 1. Place one of the samples from 3.7.2 in a 2 ml microreaction tube. 2. Wash the designated sample two times with PBS+. 3. Cover the sample in an excess of AcLDL working solution. The working solution consists of the respective cell culture medium with 10% FCS and 1.25% (v/v) AcLDL supplied stock solution. For 1 cm of sample, we used 400 μl of AcLDL working solution. 4. Incubate for 3.5 h at 37
C in a cell culture incubator. 5. Add one drop of NucBlue reagent. 6. Incubate for another 30 min. After incubation, the samples can be processed by routine protocols for fixation and visualization of immune-fluorescent stained tissue constructs. 7. Wash three times with PBS+ containing 1% BSA. 8. Fixate with 4% PFA for 5 min at room temperature. 9. Wash three times with PBS. 10. Carefully cut the circular graft open and spread out flat on a microscope slide. 11. Acquire images or use the section for further staining (e.g., Phalloidin). Exemplary images are shown in Fig. 3.

3.9 Phalloidin—FActin Staining

Visualization of the cytoskeleton by widely used f-actin fiber staining allows for a quick and easy evaluation of morphological characteristics regarding the spreading and orientation of distributed

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Fig. 3 Exemplary images of rabbit endothelial progenitor cells (rbEPC) and human saphenous vein endothelial cells (HSVEC) cultivated for 96 h on fibronectin coated bacterial nanocellulose vascular grafts. By histological examination of the cell constructs, we found highly colonized areas with apparently preserved endothelial functionality as indicated by AcLDL uptake. (a) Confocal microscopical image of CD31 stained HSVEC indicating tight cell-cell junctions. (b) Confocal microscopical image showing the merge of AcLDL uptake assay (green), Phalloidin (red), and NucBlue (blue) of rbEPC, indicating that the cells were functional intact and expressed a distinct cytoskeleton. (c) Conventional fluorescence microscopy image of rbEPC after subjection to AcLDL uptake assay. The presence of intracellular green AcLDL particles reflects a functional state of the endothelial cells. (d) 3D-reconstruction of confocal microscopical image stack, displayed is the merge of AcLDL uptake assay (green), Phalloidin (red), NucBlue (blue) of rbEPC. By 3D reconstruction, the morphology of the EC layer can be estimated i.e., by the thickness and arrangement of stress fibers in the cytoskeleton. (e, f) Images of acridine orange staining of rbEPC. (e) Magnification of whole cross-section (see rectangular region of interest in f). (f) Cross-section through fibronectin coated BNC vascular graft. The acridine orange staining allows to detect an overall graphical impression of the cell distribution on the luminal side of the graft, showing more and less confluent areas

cell bodies as well as their arrangement in colonies [18, 22]. Here, we used rhodamine conjugated phalloidin according to the manufacturer’s manual for phallotoxins to characterize the colonized scaffolds with respect to potentially formed endothelial layers. 1. Prepare the rhodamine staining solution freshly (0.1% rhodamine phalloidin supplied stock diluted in PBS containing 1% BSA and one drop per ml NucBlue reagent). The staining

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solution should be prepared in an excess of volume to ensure that the sample is completely immersed. We used 1.0 ml of staining solution in a 2 ml microreaction tube for 1 cm of sample (see Note 9). 2. Incubate the dedicated sample for 45 min at room temperature in the dark. 3. Wash three times with PBS (see Note 10). 4. Cut open the circular sample in longitudinal orientation and spread the colonized luminal surface flat onto a microscope slide. 5. Acquire images according to specifications as described in the manufacturers’ manual. Exemplary images are shown in Fig. 3. 3.10 Acridine Orange Staining

A quick and easy option to get a visual impression of cell-survival, cell-morphology and its distribution on the artificial scaffold can be achieved by Acridine Orange Staining, a fluorescent dye that reports both, cytoplasmatic RNA as well as DNA in the nuceli [18, 23]. 1. Prepare the staining solution freshly (0.1% acridine orange supplied stock solution in PBS). The staining solution should be prepared in an excess of volume to ensure that the sample is completely immersed. We used 1.0 ml of staining solution in a 2 ml microreaction tube for 1 cm of sample. 2. Place the section in a 2 ml microreaction tube and incubate for 45 min at room temperature in the dark. 3. Wash three times with PBS. 4. Cut open the section and spread it out flat on a microscope slide. 5. Acquire images according to specifications as described in the manufacturers manual or data sheet. Exemplary images are shown in Fig. 3.

3.11 CD31 Immunofluorescence Staining

Complementary to the cytoskeleton staining as described in Subheading 3.9, staining of the samples for endothelial cell-cell junctions allows investigating the cell-lining in formed colonies and layers. For this purpose, immune staining of CD31 or PECAM-1, which is expressed on endothelial cell-cell junctions and involved in platelet and leukocyte transmigration, thus also representing a marker for endothelial functionality, is widely used and accepted in literature [24, 25]. Prepare the antibody dilutions of primary and secondary antibodies with concentrations recommended by the manufacturer in PBS with 3% donkey serum in advance. We used CD31/PECAM-1

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mouse anti-human antibody in a concentration of 8 μg per ml as primary antibody and donkey anti-human Alexa Fluor 488 at the dilution of 1:500 as secondary antibody. 1. Block the dedicated graft sample in PBS containing 3% donkey serum for 45 min at room temperature in a 2 ml microreaction tube. 2. Transfer sample in a 2 ml microreaction tube containing an excess of primary antibody solution to ensure that the sample is completely immersed. 3. Incubate over night at 4
C. 4. Wash samples three times with PBS by transferring to 2 ml eppendorf tubes filled with PBS. 5. Incubate the section in a 2 ml microreaction tube containing an excess of secondary antibody solution with one drop per ml Nucblue reagent to ensure that the sample is completely immersed and incubate for 1 h in the dark at room temperature. 6. Wash three times with PBS. 7. Cut the circular sample longitudinally and spread the luminal, colonized surface flat on a microscope slide. 8. Acquire images in accordance to the specifications as described in the manufacturers manual or data sheet.

4

Notes 1. Our bioreactor chamber was customized using a 3D printer. It features four barb connectors on the inside and six on the outside. Two grafts could be perfused simultaneously. 2. The glass flask used as medium reservoir was connected to the perfusion circuit through two of the three outlets in the flask. Additionally, for sterile gas exchange, a 22 μm gas filter was connected to the reservoir. One inlet in the bottle cab was used to acquire media for substrate balancing. Another one was connected with a silicon tube to the bioreactor chamber in order to guarantee pressure release. 3. Make two loops around the tubes over the barb connectors and secure the loose ends with a surgical knot on top of the tube. Make sure, to avoid making air knots. 4. If you chose to run controls under 2D conditions for your experiments, the growth kinetics of your respective cell line should be tested in advance to ensure that the cell number normalized to surface area similar to the tested grafts does allow for cell growth without reaching high confluency levels

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or contact inhibition before the planned termination of the experiment. 5. To seed the cells, hold the bioreactor in an upright position. Attach the syringe at the lower 3-way stopcock which is now attached to the grafts at the bottom of the reactor chamber. Carefully fill the tubes until the cell suspension reaches the upper 3-way stopcocks attached to the tubes on top of the bioreactor. Close all 3-way stopcocks. 6. To determine the optimal timing for initial attachment of the cell type used in the experiments, it is advisable to test the time required for attachment in advance by 2D seeding experiments in well plates under optimal conditions, e.g., coatings that are recommended for the respective cell type. 7. Attach two syringes to the 3-way stopcocks at both sides of chamber. One syringe must be empty, the other should be filled with sterile air. Hold the chamber in an upright position again, with the air-filled syringe attached to the top of the chamber. Flush out the seeding medium while replacing it with air out of the upper syringe. Replace both syringes. This time one syringe should be filled with warm PBS. Flush 2 ml of warm PBS through each tube. Collect perfused PBS with an empty syringe on the other side. If necessary, the washout can be stored for further analysis of the non-adherent cells or e.g., cell count from washout, which can provide first insights to the number of adherent cells. 8. As cell metabolism differs between cell lines and high concentration of WST-1 assay might impact cell viability, the optimal concentration of WST-1 assay solution should be tested in advance under static conditions to evaluate the lowest concentration that still allows detecting cell proliferation. 9. If not described otherwise, all staining procedures of tube sections were carried out in 2 ml microreaction tubes. 10. If not described otherwise, all washing steps were performed for 5 min at room temperature.

Acknowledgments This project was funded by the German Research Foundation (grant number WA 4489/1-1).

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10. Jafarihaghighi F, Ardjmand M, Mirzadeh A, Hassani MS, Parizi SS (2020) Current challenges and future trends in manufacturing small diameter artificial vascular grafts in bioreactors. Cell Tissue Bank 21(3):377–403. https://doi.org/10.1007/s10561-02009837-0 11. Mertsching H, Hansmann J (2009) Bioreactor technology in cardiovascular tissue engineering. Adv Biochem Eng Biotechnol 112:29–37. https://doi.org/10.1007/978-3540-69357-4_2 12. Maschhoff P, Heene S, Lavrentieva A, Hentrop T, Leibold C, Wahalla M-N, Stanislawski N, Blume H, Scheper T, Blume C (2017) An intelligent bioreactor system for the cultivation of a bioartificial vascular graft. Eng Life Sci 17(5):567–578. https://doi.org/10. 1002/elsc.201600138 13. McFetridge PS, Bodamyali T, Horrocks M, Chaudhuri JB (2004) Endothelial and smooth muscle cell seeding onto processed ex vivo arterial scaffolds using 3D vascular bioreactors. ASAIO J 50(6):591–600. https://doi.org/ 10.1097/01.mat.0000144365.22025.9b 14. Schuerlein S, Schwarz T, Krziminski S, Gatzner S, Hoppensack A, Schwedhelm I, Schweinlin M, Walles H, Hansmann J (2017) A versatile modular bioreactor platform for Tissue Engineering. Biotechnol J 12(2). https:// doi.org/10.1002/biot.201600326 15. Freed LE, Vunjak-Novakovic G (2000) Chapter 13: Tissue engineering bioreactors. In: Lanza RP, Langer R, Vacanti J (eds) Principles of tissue engineering, 2nd edn. Academic Press, San Diego, pp 143–156. https://doi. org/10.1016/B978-012436630-5/50017-9 16. De Bock K, Georgiadou M, Schoors S, Kuchnio A, Wong Brian W, Cantelmo Anna R, Quaegebeur A, Ghesquie`re B, Cauwenberghs S, Eelen G, Phng L-K, Betz I, Tembuyser B, Brepoels K, Welti J, Geudens I, Segura I, Cruys B, Bifari F, Decimo I, Blanco R, Wyns S, Vangindertael J, Rocha S, Collins Russel T, Munck S, Daelemans D, Imamura H, Devlieger R, Rider M, Van Veldhoven PP, Schuit F, Bartrons R, Hofkens J, Fraisl P, Telang S, DeBerardinis Ralph J, Schoonjans L, Vinckier S, Chesney J, Gerhardt H, Dewerchin M, Carmeliet P (2013) Role of PFKFB3-driven glycolysis in vessel sprouting. Cell 154(3):651–663. https://doi.org/10.1016/j.cell.2013.06.037 17. Hulsmann J, Aubin H, Wehrmann A, Jenke A, Lichtenberg A, Akhyari P (2016) Whole-heart construct cultivation under 3D mechanical

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acetylated-low density lipoprotein. J Cell Biol 99(6):2034–2040. https://doi.org/10.1083/ jcb.99.6.2034 22. Avari H, Rogers KA, Savory E (2019) Quantification of morphological modulation, F-actin remodeling and PECAM-1 (CD-31) re-distribution in endothelial cells in response to fluid-induced shear stress under various flow conditions. J Biomech Eng. https://doi.org/ 10.1115/1.4042601 23. Zhang K, Li JA, Deng K, Liu T, Chen JY, Huang N (2013) The endothelialization and hemocompatibility of the functional multilayer on titanium surface constructed with type IV collagen and heparin. Colloids Surf B Biointerfaces 108:295–304. https://doi.org/10. 1016/j.colsurfb.2012.12.053 24. Privratsky JR, Newman PJ (2014) PECAM-1: regulator of endothelial junctional integrity. Cell Tissue Res 355(3):607–619. https://doi. org/10.1007/s00441-013-1779-3 25. Lertkiatmongkol P, Liao D, Mei H, Hu Y, Newman PJ (2016) Endothelial functions of platelet/endothelial cell adhesion molecule-1 (CD31). Curr Opin Hematol 23(3):253–259. https://doi.org/10.1097/MOH. 0000000000000239

Methods in Molecular Biology (2022) 2436: 223–240 DOI 10.1007/7651_2021_424 © Springer Science+Business Media, LLC 2021 Published online: 15 September 2021

A Guideline to Set Up Cascaded Continuous Cultivation with E. coli Bl21 (DE3) Julian Kopp and Oliver Spadiut Abstract Continuous processing allows to maximize space-time yields and is implemented in many industrial branches. However, in manufacturing of value added compounds produced with microbial hosts, continuous processing is not state-of-the-art yet. This is because fluctuating productivity causes unwanted process deviations. Cascaded continuous bioprocessing, unlike conventional continuous process modes, was found to result in stable productivity. This manuscript serves as a guideline how to set up a cascaded continuous cultivation with Escherichia coli BL21 DE(3). Key words Cascaded continuous cultivation, Escherichia coli BL21(DE3), Long-term stable processes, Microbial continuous cultivation, Stable productivity

1

Introduction Continuous bioprocessing has been implemented in many industrial branches [1]. This is because continuous production allows a major improvement in volumetric throughput by facilitating manufacturing in smaller production scales [1–3]. In respect to biotechnological applications, Herbert et al. demonstrated the economic feasibility of continuous bioprocesses over batch cultivations back in 1956 [4]. Continuous cultivation was already introduced by Monod, Novick, and Szilard in the early 1950s [5–8]. The so-called chemostat cultivation was originally developed to investigate cell physiology allowing prediction of environmental conditions on host cells [7–9]. Biotechnological approaches in industry nowadays try to follow the principles of Herbert et al. [4], to boost the space-time yield of conventional processes. Unfortunately, this approach proved to be more tricky than expected [10]. Many recombinantly produced components are toxic for host cells at required industrial product titers [11], potentially causing process deviations. As host cells try to escape the formation of the target product, effects might lead to subpopulation diversification [11–13]. The formation of subpopulations (either phenotypic or genotypic) [14–16] leads to unwanted process deviations.

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As the likelihood of subpopulation formation is increasing with elongated generation times [11], continuous processes especially suffer from subpopulation formation. Diverse microbial continuous processes and their effects on productivity have been summarized in literature already [17, 18]: Productivity over process time resulted in a bell shaped curve: upon induction of inducible promotors, productivity increased up to a stable level; however, ongoing subpopulation diversification is believed to promote a fast decrease of recombinant protein formation. Unlike microbial continuous cultivation, continuous bioprocessing has already been implemented for mammalian host cell lines, and the first products have been commercialized [19– 22]. As mammalian cell lines are known to propagate at slower growth rates than microbial cells, subpopulation effects might not be visible during “conventional continuous process times” (i.e., 3–6 weeks process duration) [10, 11]. As industry accomplished stable continuous processes with mammalian cell lines, the microbial production sector is aiming at realizing stable continuous processes as well. Results indicate cascaded continuous cultivation to outperform (conventional) chemostat cultivation in regard to long-term stable productivity [23–25]. Chemostat cultivation is limited in its biomass production, as dilution rate and fed substrate have to be adapted in order to avoid host cell washout [26, 27]. As cascaded continuous cultivations uses two sequentially operated chemostat processes, higher biomass can be achieved compared to conventional chemostat processes [25]. Cascaded continuous cultivation comprises two sequentially continuously operated reactors without additional requirements for cell retention [10, 25]. Thereby a spatial separation of biomass growth and target protein formation can be achieved. Reactor one (i.e., stage one) is conventionally used for biomass growth only. Noninduced cells are transferred to the second stage where an additional feed is applied for induction. Recombinant product can be harvested as a bleed stream from the second stage. For E. coli BL21(DE3), pET plasmids are frequently employed controlling target gene expression under control of the lac promotor [28, 29]. Induction is thus restricted to either isopropyl-β-D-1thiogalactopyranoside (IPTG) or the natural inducer allolactose, formed by fed lactose [30, 31]. IPTG has shown beneficial results when used for short induction times (i.e., fed-batch cultivations) [28, 32]. On the other hand side, IPTG induction was described to exhibit toxic effects on host cells, especially visible at higher generation times [33]. Lactose induction facilitated a more stable productivity than IPTG induction for both chemostat and cascaded continuous cultivation. Still, when feeding lactose, carbon catabolite repression (CCR) is a well-known phenomenon occurring in substrate co-feeding [34, 35]. The glucose-lactose diauxic growth causes decreased lactose uptake rates when glucose is present in

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excess [34, 35]. The effect of glucose and glycerol as carbon source in combination with lactose as inducer was investigated in diverse pre-studies [35, 36]. In contrast to glucose, higher lactose uptake rates could be monitored at higher feeding rates for glycerol feeding. Higher inducer uptake rates are believed to boost heterologous protein production [31, 37, 38]. Chemostat cultivations on glycerol–lactose systems indicated long-term productivity can be boosted compared to glucose-lactose co-feeding [36]. Based on our previous investigations and stated arguments in literature, glycerol and lactose show a suitable combination to enable stable longterm cultivations. Therefore, we focus the method workflow given in this protocol to the utilization of glycerol and lactose. In the following sections, a guideline is given on how to set up a cascaded continuous cultivation with E. coli BL21(DE3).

2

Materials

2.1

Host Cells

Competent E. coli BL21(DE3) host cells (Life technologies, Carlsbad, CA, USA) are used. Transformation of target plasmid in E. coli host cells is described in detail elsewhere [39–41]. Depending on the plasmid backbone, the antibiotic resistance has to be adapted for cultivation.

2.2

Required Media

A defined medium referred to DeLisa et al. is used for all cultivations [42]. Medium composition is summarized in Table 1.

2.3 Required Devices for Cultivation

1. For preculture cultivation, an incubation shaker is required, allowing shaking possibilities up to 250 rpm and a temperature control in the range of 30–40
C (e.g., Multitron shaker, Infors, Bottmingen, Switzerland). 2. For cascaded continuous cultivations (at least) two continuously operated stirred-tank reactors are required: (a) Both reactors require separate control systems. (b) Reactors need to be capable of stirring to at least 1400 rpm. (c) Reactors need to be capable of supplying pressurized air at minimum 2 vvm. (d) Additional oxygen supply must be present to mix gas flows to a desired ratio. (e) A PI- or PID-controller needs be integrated in the process control system to keep dO2 > 30%. (f) Devices must have a working volume larger than 250 mL; for example, Minifors 2 reactor systems, Labfors 4 or 5 reactor systems or DASGip cultivation systems could be used. Smaller devices showed process deviations due to sample volume taken.

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Table 1 Composition of the defined medium according to DeLisa et al., which is used for all performed cultivations [42]. Stock solutions for trace elements are sterilized separately. All DeLisa medium components can be sterilized together at 121 ˚C (see Note 1) DeLisa medium

Final conc. (g/L)

Glycerol KH2PO4 (NH4)2HPO4 Citric acid

Adjust to desired amount 13.30 4.00 1.70

Stock solutions

Final conc. (g/L)

MgSO4 · 7 H2O Fe(III) citrate EDTA Zn(CH3COO)2 ·2 H2O CoCl2 · 6 H2O MnCl2 · 4 H2O CuCl2 · 2 H2O H3BO3 Na2MoO4 · 2 H2O Thiamine HCl

1.20 0.1000 0.0084 0.0130 0.0025 0.0150 0.0012 0.0030 0.0025 0.0045

(g) Reactors need to be equipped with immersion tubes adjusted to the desired volume and controlled via a peristaltic pump. (h) Each bioreactor requires a suitable PI-controller connected to a peristaltic pump to control the pH value. (i) All process parameters applied during cultivation need to be logged. We recommend to use a process control system to alter diverse set-points. 3. Each reactor system needs to be equipped with at least four pumps: (a) Pump for feeding. (b) Pump for transferring/harvesting cultivation broth depending on the reactor. (c) Pump for acid control. (d) Pump for base control. 4. All bioreactors need to be equipped with a pH probe (e.g., EasyFerm plus pH-sensor (Hamilton, Reno, NV, USA)). pH probes need to be calibrated within a pH range of 4.01–7. pH needs to be controlled for acidic deviations with 12.5% NH4OH solution. If alkaline process deviations occur, pH can be adjusted with 5% H3PO4.

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5. All bioreactors need to be equipped with a dissolved oxygen (dO2) probe to monitor dO2 in cultivation broth; e.g., a fluorescence-dissolved oxygen electrode Visiferm DO (Hamilton, Reno, NV, USA). Probes must be calibrated prior use. In case dilution rates/residence times are adjusted via an immersion tube and the fermentation broth volume, dO2 control is only performed by adding oxygen as a stirrer-based control would lead to deviations in the dilution rate. 6. All feeds and base/acids dosage bottles need to be placed on data-recording scales. 7. Residual offgas from reactors needs to be quantified with required offgas sensors (e.g., Blue Sens Gas analytics, Herten, Germany). (a) Exhausting oxygen is quantified by zirconium oxide sensors. (b) Exhausting carbon dioxide is quantified by near-infrared spectra analysis. 2.4 Required Equipment for Process Analysis

1. Any photometric device/plate reader quantifying the absorption in liquid broth can be used, e.g., a Genesys 20 photometer (Thermo Scientific, Waltham, MA, USA).

2.4.1 Biomass Determination

2. Cuvettes for OD600 determination. 3. 2-mL Eppendorf Safe-Lock Tubes (Eppendorf, Hamburg, Germany). 4. Oven at 105
C. 5. Small-scale centrifuge capable of cooling and spinning down 2-mL safe lock tubes (Eppendorf, Hamburg, Germany). 6. Filtered 0.9% NaCl solution for washing steps.

2.4.2 Viable Biomass Determination

1. Flow cytometry device with integrated 488 nm laser and detector cell (e.g., Partec, Sysmex Germany). 2. Fluorescent stains for live–dead determination, e.g., DiBAC4 (3) (bis-(1,3-dibutylbarbituricacid-trimethineoxonol). 3. Filtered 0.9% NaCl solution for dilution steps.

2.4.3 Metabolite Determination

1. Liquid chromatography device including refractory index (RI) detector (e.g., Dionex Ultimate 3000 SD, Thermo Scientific, Waltham, MA, USA); the device must include: (a) Pump module. (b) Auto-sampler module. (c) Oven module. (d) Detector module. (e) RI detector module.

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2. HPLC (high performance liquid chromatography)-grade eluents (i.e., purified water). 3. Anion exchange chromatography column (e.g., Supelcogelcolumn (Agilent, Vienna, Austria)). 4. All fed carbon sources and all residual metabolites in solid/ liquid form for quantification. (a) In this case, feeding is conducted with lactose and glycerol. In addition, glucose and galactose, resulting from hydrolyzed lactose, need to be quantified. Furthermore, any metabolite potentially accumulating (e.g., acetate) needs to be prepared as a standard reference. 5. Syringes and 0.2-μm syringe filters. 2.4.4 Product Determination

3 3.1

This section is highly dependent on the expressed target protein and must be conducted according to established analytics. In case of intracellular product formation, a high-pressure homogenizer is required (e.g., PANDA+ 2000, GEA, Biberach, Germany).

Methods Cultivation Setup

1. All bioreactor and preculture cultivations are carried out using the same medium composition to avoid deviations. We recommend a defined chemical medium, to reduce process distributions (e.g., DeLisa et al., 1999, [42]). 2. Cultivation setup depicted in Fig. 1. 3. For the adjustment of the correct dilution rate, dip-pipes need to be adjusted to the required height in the bioreactors: (a) Calculate the desired residence time (or dilution rate) given the applied volumetric flow in the reactor. Dilution rate can then be adjusted either (i) via volumetric flow or (ii) via volume in the reactor; e.g., 100 mL/h is the volumetric flow in the reactor and the residence time is desired at 10 h. Hence, 1 L operating volume is required for continuous cultivation. (b) Fill reactor with desired volume prior to sterilization and apply process parameters (e.g., 1,400 rpm and 2 vvm). (c) Make sure reactor contains all probes and pipes at this stage, which could influence the liquid level in the reactor. (d) Set immersion pipe to adequate height and mark at the reactor side in case deviations might occur during sterilization. 4. For continuous cultivation, constant volume can be adjusted in two ways:

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Fig. 1 Process overview of a continuous cascaded cultivation. Reactor 1 (i.e., stage 1) is used for biomass formation. Biomass stream is transferred to reactor 2 (i.e., stage 2); recombinant protein production does only take place in the second stage; Di ¼ dilution rate (1/h), Fi ¼ feed rate (L/h), Xi ¼ biomass flow (L/h), Pi ¼ product flow (L/h), Vi ¼ volume (L)

(a) Via an immersion pipe set to certain reactor volume, which is connected via a tube to a peristaltic pump. For continuous cultivation, pump settings given beneath need to be adapted at all times: pump rate flow in < pump rate flow out. (b) Via gravimetric control of the reactor weight, which can be monitored and controlled via a PI-controller in combination with a pneumatic valve. The valve opens once the weight deviation exceeds a certain set-point and close afterwards again. 3.2 Cultivation Scheme 3.2.1 Preculture and Batch Phase

Cultivation scheme depicted in Fig. 2 represents a guidance to cascaded continuous cultivation (see Note 2). 1. All required media need to be sterilized at previously mentioned conditions (i.e., sterilization at 121
C or filtered at 0.2 μm in sterile environment). 2. Preculture is conducted as an overnight culture, i.e., 20 h, at 37
C, and 230 rpm. 3. In addition to Table 1, the pH for the preculture needs to be set to 7.2 prior to sterilization with a 5 M NaOH stock solution.

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Fig. 2 The process overview of the continuous cascaded cultivation on a time-dependent manner for seed reactor (reactor 1) and recombinant production reactors (reactors 2 and 3); down-times consist of steam in place (SIP) and cleaning in place (CIP) phases, whereas cultivation is separated in a batch phase, a continuous adaptation phase, and a continuous induced cultivation phase

4. Depending on the targeted biomass for preculture, noninducing carbon source (i.e., glycerol) needs to be adapted in the medium composition; preculture volume(s) need to be adjusted to the desired reactor size; i.e., a preculture consisting of 8 g/L glycerol will result in a biomass concentration of approximately 4 g/L, assuming a biomass substrate yield of 0.5 g/g. In case reactor volume consist of 1 L, the preculture volume needs to be 100 mL to allow inoculation with 1/10th of fermentation broth (see Note 3). 5. Reactors are adapted to batch culture conditions prior to inoculation: 37
C, 1400 rpm stirring, 2 vvm aeration, and a set pH of 6.7. Media pH is set to desired value with 12.5% NH4OH after sterilization. 6. Once set process parameters are achieved, dO2 probes are calibrated for 100% dissolved oxygen. 7. After 20 h of cultivation, the preculture is quantified in its optical density (OD600). Depending on the host strain and the used amount of noninducing carbon source, the targeted biomass should be achieved. 8. Batch media is inoculated with 10% Preculture; dO2 in cultivation must never be beneath 30% at all stages of the cultivation ! adjustment via PI controller is necessary. 9. End of batch phase is indicated by either a dO2 peak or a drop in the residual CO2. 3.2.2 Continuous Adaptation Phase

1. To get the cells in equilibrium state prior to induction, the continuous cultivation is started; however, only noninducing feed is applied at this phase (Fig. 3). Cell equilibrium state can be obtained by a constant offgas signal in the biomass reactor.

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Fig. 3 Process overview of the continuous cascaded cultivation. All lines in green are opened during continuous adaptation phase, whereas black lines are closed. Reactor 1 ¼ stage 1 is used for biomass formation. Biomass streams are separated equivalent into reactor 2 and 3 ¼ stage 2; Stage 2 is thus adapted for continuous recombinant protein expression during the continuous adaptation phase. Di ¼ dilution rate (1/h), Fi ¼ feed rate (L/h), Xi ¼ biomass flow (L/h), Pi ¼ product flow (L/h), Vi ¼ volume (L)

Hence, the continuous adaptation phase only consists of feeding in stage 1; however, no feed is added to stage 2, as this feed contains of inducer. 2. For the cascaded continuous cultivations, stirred-tank reactors are connected by transfer tubes coupled to peristaltic pumps (e.g., Ismatec, Wertheim, Germany). 3. At the end of the batch phase, reactor temperature is adjusted to desired values: (a) Stage 1: for biomass formation. (b) Stage 2: for recombinant protein production. 4. Continuous adaptation phase lasts for 4–5 residence times for media exchange at least. 5. For screening of ideal process conditions, a higher amount of induced bioreactors would reduce screening times; as biomass

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stream of one seed reactor can be adapted to multiple induced reactor systems, as shown for two induced reactors in Fig. 3 exemplarily. 6. For continuous adaptation phase, only lines marked in green are opened at this level of cultivation (Fig. 3); all lines marked in black (i.e., induction feed lines) are closed at this stage. 7. For continuous adaptation phase, only lines marked in green are opened (Fig. 3); all lines marked in black (i.e., induction feed lines) are closed at this stage. 3.2.3 Induction Phase

1. Once stage 1 is in equilibrium state (constant CO2 signals for 4 residence times at least), induction feed can be added. 2. Pumps need to be adjusted to desired volumetric rates. 3. All lines are opened at this stage (Fig. 4). 4. Efflux of stage 1 thus contains solely biomass, whereas efflux of stage 2 contains cells including product (see Note 4).

Fig. 4 Process overview of the continuous cascaded cultivation. Reactor 1 is used for biomass formation. Biomass streams are separated equivalent into reactors 2 and 3; Reactors 2 and 3 are used for recombinant protein production. For screening of recombinant protein production, only process parameters in reactors 2 and 3 are varied; all lines (marked in green) are opened in this figure; Di ¼ dilution rate (1/h), Fi ¼ feed rate (L/h), Xi ¼ biomass flow (L/h), Pi ¼ product flow (L/h), Vi ¼ volume (L)

A Guideline to Set Up Cascaded Continuous Cultivation with E. coli Bl21 (DE3)

3.3 Sampling and Analysis

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Prior to induction, samples after batch are taken to determine viable biomass growth and potential metabolite accumulation. An additional sample after the continuous adaptation phase must be taken to again determine viable biomass and metabolite accumulation. Sampling frequency during induction is highly dependent on: l

The doubling time of the host organism.

l

Expected deviations in productivity.

For E. coli BL21 (DE3), samples are taken once to twice a day. In addition to previously stated analysis, product formation needs to be monitored additionally after induction. 3.3.1 Biomass Determination

Biomass is quantified via optical density (OD600) and gravimetrically by weighing the dry cell weight (DCW) while flow cytometry analysis (FCM) was used for the determination of cell death. OD600 Determination

1. Any photometric device/plate reader quantifying the absorption in liquid broth can be used, e.g., a Genesys 20 photometer (Thermo Scientific, Waltham, MA, USA). 2. Vortex fermentation broth after sampling to get homogenous solution for 5–10 s. 3. Determine OD600 using photometric cuvettes within the linear range of the system (i.e., 0.2–0.8 [AU]). 4. In case the sample is not measured within linear range, dilute sample after vortexing with deionized water; make sure to never exceed 1:10 dilution steps due to accuracy. 5. Conduct at least duplicate measurements. DCW Determination

1. Preheat 2-mL Safe-Lock Tubes (e.g., Eppendorf, Hamburg, Germany) overnight at 105
C to free volatile compounds prior to gravimetric determination. 2. Cool down tubes to room temperature, label reaction tubes, and weigh the pre-tared amounts. 3. Vortex fermentation broth for 5–10 s after sampling to get homogenous solution. 4. Pipette 1 mL in a pre-tared 2-mL reaction tube. 5. Centrifuge the sample in a small-scale centrifuge for 10 min at 11,180  g at 4
C. 6. Separate the supernatant from the pellet by decanting it (see Note 5). 7. Immediately resuspend the pellet in 0.9% NaCl solution. 8. Centrifuge the dissolved pellet solution at the previously given conditions (10 min at 11,180  g at 4
C).

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9. Triplicates must be performed for DCW determination per sample. Cell Viability Determination

1. Fermentation broth is diluted with pre-filtered 0.9% NaCl solution to an approximate OD of 0.01–0.05 (see Note 6). 2. Stock solution of 0.5 mM DiBAC4(3) (bis-(1,3-dibutylbarbituricacid-trimethineoxonol) is dissolved and diluted with DMSO (dimethylsulfoxide) (see Note 7). 3. Diluted sample is fused with 1.5 μL of 0.5 mM DiBAC4(3)stock solution. 4. Additionally, the stain RH 414 (N-(3-triethylammoniumpropyl)-4-(4-(4-(diethylamino)phenyl)butadienyl)pyridinium dibromide) could be added (1.5 μL of 2 mM stock solution) to reduce background signals. 5. Incubate the sample for approximately 5 min in the dark. 6. Measure the sample with a suitable flow cytometry device (e.g., Partec, Cube 8, Sysmex, Germany). 7. Treat data with a suitable evaluation software (e.g., FACS express 5.0, Sysmex, Germany); more details to this very method can be found here [43]. 3.3.2 Determination of Metabolite Accumulation

Supernatant samples from sugar concentrations of feed and fermentation broth are quantified via liquid chromatography. 1. Vortex fermentation broth after sampling to get homogenous solution for 5–10 s. 2. Pipette 1 mL in a pre-tared 2-mL reaction tube. 3. Centrifuge the sample in a table centrifuge for 10 min at 11,180  g at 4
C. 4. Separate the supernatant from the pellet by decanting it. 5. Supernatant at this purification step contains of accumulated sugars, required for quantification. 6. Filter supernatant samples at 0.2 μm prior to HPLC analysis. 7. Prepare an eluent at 0.1% H3PO4. 8. Prepare an anion exchange chromatography (e.g., SupelcogelAEX) for analysis at 30
C and a flow of 0.5 mL/min; Chromatogram recording should be carried out for 30 min. 9. Prepare standards at 1, 5, 10, 20, 30, 40, and 50 g/L of the respective used carbon sources and accumulating metabolites (see Note 8). 10. Chromatograms must be analyzed with a suitable processing software (e.g., Chromeleon; Thermo Scientific, Waltham, MA, USA).

A Guideline to Set Up Cascaded Continuous Cultivation with E. coli Bl21 (DE3) 3.3.3 Product Determination

3.4 Calculation of Flow and Substrate Uptake Rates 3.4.1 Calculation of Flow Rates

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Product determination is highly dependent on the given target protein and subsequent analytic procedures; hence, no general protocol can be given. 1. Dilution rate in the first stage needs to be calculated according to chemostat principles rendering to Monod, Novick, and Szilard with μ ¼ qs  YX/S ¼ D; if μ < Dcrit [5–7]; dilution rates beyond Dcrit must be avoided. 2. Dilution rate is calculated via monitored feed volumes over time (Eq. 1). Reactor volume must be kept constant. As feeding rates are monitored gravimetrically and feed density is determined, volumetric rates were calculated as follows: D Ind: Reactor ¼

F_ indFeed þ F_ biomass V reactor

ð1Þ

Dind. Reactor . . . dilution rate in the reactor for protein expression (1/h). Find. Feed . . . applied feeding rate of the induction feed (mL/h). Fbiomass . . . applied biomass flow to the protein expression reactor (mL/h). Vreactor . . . volume in the protein expression reactor (L). 3.4.2 Calculation of Substrate Uptake Rates

1. Feed composition in the induction reactor can be adjusted by: (a) The ratio of transferred cells to the amount of feed solution. (b) The concentration in the induction feed. 2. Specific substrate uptake rate is calculated according to Eq. 2. 3. Real sugar uptake rates need to be used, evaluated by the accumulated component subtracted from the fed substrate. q s,c ¼

c Feed  V Feed,in  c acc  V reactor Δt  X Δt

ð2Þ

qs,c . . . specific uptake rate of desired component (g/g/h). cFeed . . . concentration of corresponding component in feed (g/L). VFeed,in . . . feed volume in reactor in timespan Δt (L). cacc . . . concentration of the corresponding component which accumulated in the reactor (HPLC measurement) (g/L). Vreactor . . . volume in the protein expression reactor (L). Δt . . . timespan for calculated uptake rate (h). XΔt . . . average biomass in the protein expression reactor during the timespan Δt (g). 4. In case mixed feeds are used, a careful evaluation of the carbon source to inducer ratio must be done prior to utilization. Screening approaches therefore can be found in given citations [35, 38].

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Notes 1. If glycerol is exchanged by glucose, carbon source needs to be sterilized separately in split volumes. MgSO4 · 7H2O, Fe(III) citrate, and EDTA were sterilized at 121
C for 20 min. All other chemicals were filtered at 0.2 μm under sterile conditions in pre-sterilized bottles. 2. Please note that process times are highly dependent on the host organism and are given in this case for E. coli BL21(DE3). 3. Initial substrate concentrations beyond 50 g/L might lead to high metabolite accumulation (i.e., acetate) and must be avoided. 4. Even though productivity might be constant in stage 2, the offgas signal might deviate due to subpopulation diversification, potentially resulting in different sugar uptake rates; still offgas signals in the seed reactor should be constant at all time. 5. Supernatant at this stage contains accumulated sugars/proteins and requires storage at 20
C for further analysis. 6. Dilutions with deionized water might lead to deviations in quantification (due to osmotic pressure and subsequent cell lysis); dilution steps should thus not exceed 1:50 dilutions. 7. Flow cytometer stain is light sensitive and thus requires adequate storage. DiBAC4(3) can only penetrate cells, if the cell wall is damaged. Due to an increase in the fluorescent gain, a differentiation between viable and non-viable cells can be obtained. 8. Each metabolite can result in an altering RI-fingerprint; hence, calibration is only valid with respective standard substance.

5

General Notes on Process Intensification for Cascaded Continuous Cultivation For E. coli BL21 (DE3), a careful optimization strategy has to be carried out in microbial continuous cultivation regarding: (a) Dilution rates. (b) Substrate uptake rate. (c) Cultivation temperature.

5.1

Dilution Rate

1. Maximum growth rate is determined in a batch culture. (a) A fit of residual carbon dioxide signals can be performed. (b) Frequent sampling and determination of biomass growth and carbon reduction has to be done. (c) A 96-well plate reader screening optical density at controlled environments (e.g., Tecan, M€annedorf, Switzerland) can be used to determine μmax.

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2. According to determined maximum growth rate, turbidostat experiments can be laid out, to determine washout of host cells. (a) This can be done in a single experiment increasing the dilution rate in a stepwise approach. (b) Interval is adjusted according to μmax in 5–10% steps. (c) Each step needs to be held for 4–5 residence times until biomass washout can be obtained. 3. Effect of dilution rate must be correlated to productivity: This approach is confronted with a high experimental load where effects of product stability need to be correlated to the dilution rates. (a) Parallelization increases throughput induced reactors (Figs. 3 and 4)).

(i.e.,

multiple

(b) Design of experiment approaches can be used in case a second factor influences productivity to reduce experimental workload. 5.2 Substrate Uptake Rate

1. In the first stage of cascaded continuous cultivation, μ can be directly correlated to the specific substrate uptake rate. 2. In the second stage, substrate uptake rate can be either adjusted via the ratio of inducer feed to the ratio of transferred cells or the feed concentration; hence, Eq. 2 must be used for specific substrate uptake rate calculation. 3. For experimental approaches on specific uptake rate determination, the following approach is utilized: (a) Inducer feed is set to lower values at first, whereas biomass stream requires settings to a higher level to avoid washout of host cells. (b) Desired ratios can be adjusted in a turibodstat-like approach; i.e., start at 90% Biomass-stream (X1) and at 10% F2 or F3 of the overall dilution rate in stage 2. 4. When altering the overall dilution rate in stage 2 of cascaded continuous cultivation, the approach shown in point 3 has to be repeated until the desired specific substrate uptake rate is adjusted. This is required as the average residence time influences substrate uptake rates.

5.3 Cultivation Temperature

1. Effects of productivity in microbials is often correlated to expression temperature. Temperature dependency thus needs to be investigated. 2. As continuous processes suffer from a high experimental load, screening for optimized production temperature is recommend to be performed in fed-batch cultivations.

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3. In case a second parameter is believed to influence productivity, a design of experiment (DoE) reduces experimental load. 4. Obtained effects of productivity at given temperature can then be applied for continuous cultivation.

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Methods in Molecular Biology (2022) 2436: 241–256 DOI 10.1007/7651_2021_442 © Springer Science+Business Media, LLC 2021 Published online: 02 November 2021

Applying Stirred Perfusion to 3D Tissue Equivalents to Mimic the Dynamic In Vivo Microenvironment Henry W. Hoyle, Claire L. Mobbs, and Stefan A. Przyborski Abstract Complex three-dimensional (3D) tissue equivalents have been widely developed with applications with a multitude of organs and tissues. While these systems lead to significant improvements over conventional two-dimensional culture, the static conditions of the surrounding medium still present a limitation to the physiological relevance of these models. Medium perfusion and convective mixing can be introduced to these models through a variety of techniques using equipment such as pumps and rockers. These systems can easily become very complex or suffer from limited control over the fluid flow properties. We have developed a bioreactor enabling controlled perfusion of 3D tissue equivalents utilizing a magnetic stirrer–based system, allowing for scalability and ease of use. This system has demonstrated potential applications in a range of tissues such as the liver, intestine, and skin, with many other potential applications yet to be tested. Our solution provides users with a low cost and easy to use alternative to complex bioreactor systems while still providing high levels of control over fluid flow and structural properties of the tissue constructs. Key words Bioreactor, Perfusion, Organotypic culture, Alvetex®, Tissue engineering, Three-dimensional cell culture

1

Introduction Cells and tissues in vivo exist in a highly complex and dynamic microenvironment. This has a wide range of factors influencing their structure and function. These include but are not limited to: high levels of cell–cell and cell–matrix contact, facilitating communication and the response of tissues to stimuli; the presence of an extracellular matrix (ECM) composed of a variety of different ECM proteins and supporting three-dimensional (3D) growth of cells; fluid flow from the bloodstream and other fluids present in tissues, providing low levels of shear stress and allowing adequate turnover of nutrients and cellular waste. These factors are all important for the healthy function of tissues, however only a subset of these are usually present in tissue models in vitro [1, 2]. A variety of methods have been developed to overcome this limitation and therefore maximize the physiological relevance of in vitro tissue equivalents. One of the major areas of study for this purpose is the use of 3D growth substrates. These can range from

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Fig. 1 Different functions that fluid flow can perform in cells or tissues. (a) Nutrients and signaling molecules such as hormones are delivered to the cells through the bloodstream. (b) Secreted products and metabolites from the cells are removed, preventing a buildup of toxic molecules and maintaining osmotic balance. (c) Flow disrupts unstirred layers which build up close to the cell surface, increasing turnover of nutrients and waste products in this region. (d) Shear stress provides a mechanical stimulus to cells which can induce specific functionality or polarization

simple micropatterned surfaces to complex matrices with tunable parameters to be adapted for the cells of interest. These can be further improved through the incorporation of extracellular matrix proteins in forms such as hydrogels or matrix coating [3]. The composition of the ECM in vivo is variable in different tissues as well as in disease states [4], and this variability can be recapitulated with in vitro models [5]. Fluid flow is one factor which is less frequently used in cell and tissue culture, though can also have a major impact on tissue structure and function. There are many properties which are modulated by tissue perfusion and therefore lacking in static culture methods, as shown by Fig. 1. The fluid flow causes shear stress along the membranes of cells which can then impact their function through mechanotransduction. The exact levels of shear stress can often be difficult to elucidate in tissue, though it is typically at microdyne/cm2 levels due to the protective function of endothelial cells around blood vessels [6]. Further improvements brought about by perfusion are due to the replenishment of medium around the cells. This ensures a consistent delivery of nutrients, both constituents of the medium such as proteins and growth factors, as well as components such as dissolved oxygen which are maintained at consistent levels with the convective mixing in perfusion systems. This transport is also effective for removal of metabolic waste products such as lactic acid, as well as other secreted compounds such as albumin, which can otherwise build up close to the cell surface to the detriment of the cells. The disruption of unstirred layers close to the cell surface is also beneficial for this. The variable concentrations found in static conditions near to membranes are disrupted with perfusion, resulting in bulk medium concentrations being the same for regions closer to the cells. This is also beneficial

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for applications such as drug toxicology, with the reduction in unstirred layers leading to an improved, more physiologically relevant transfer of drugs to the cells [7]. There are many techniques which can be used to introduce media perfusion into cell culture systems. These can range from very simple systems such as placing standard culture ware onto an electronic shaker or rocker, to complex systems consisting of multiple components such as pumps and medium reservoirs. The higher complexity typically comes with a greater level of control over the fluid flow properties such as shear stress and mass transfer, however, can also lead to greater spatial requirements, increased levels of user expertise required, and higher cost. The system utilized for this protocol uses a magnetic stirrer to induce perfusion in a 3D cell culture system based around porous Alvetex® membranes. This allows for a simplistic setup, which can be used with minimal expertise required while still giving control over properties of the fluid across the cells. A completely selfcontained vessel with a large volume of medium means that cells and tissues can be cultured for prolonged periods without the need for media changing, leading to more consistent culture conditions over time. As shown by Fig. 2, this system consists of a glass beaker to contain the culture, a polytetrafluoroethylene (PTFE)-coated magnetic stir bar and PTFE stand which holds an Alvetex® insert as well as providing control over the fluid flow, and a PTFE lid.

Fig. 2 The stirred bioreactor system. This design incorporates includes the conical stand to hold a 6- or 12-well Alvetex insert. A stir bar placed at the bottom provides continuous fluid recirculation. (a) Exploded diagram of the major components of the bioreactor system. (b) The system when fully assembled. The vessel can hold up to 120 mL of cell culture medium

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Fig. 3 Schematics for the Alvetex insert. The insert holder has a conical baffle for the lower portion to help direct fluid recirculation within the vessel while the top of the holder features a notched upper surface to allow the Alvetex insert to be held in place. Scale bar: 30 mm

The entire system, apart from the Alvetex® insert, is repeatedly autoclavable, allowing for sterilization and therefore a high level of reusability. This bioreactor vessel can be placed on a commercial magnetic stirrer; however, a custom system is utilized for the experiments described herein due to the ability to stir a number of vessels simultaneously and using stir speeds down to 50 rpm. Figure 3 shows the design of the custom stand to hold the Alvetex® inserts. This was designed to promote fluid recirculation throughout the vessel while protecting the cultures from high levels of shear stress, however the exact design of this stand can be varied depending on the desired flow properties. The preferred material for the Alvetex® insert holder is PTFE due to its minimal chemical and biological activity and the ability to withstand multiple autoclave cycles. The outer vessel was made from glass due to the ease of production and low costs on top of its biological compatibility and autoclavability. While a custom lid can be used, a standard 60 mm petri dish lid may be adequate for successful culture with this system.

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Fig. 4 H&E staining and metabolic function of HepG2 grown on Alvetex Strata in static and perfusion. (a) HepG2 cells were grown on Alvetex Strata in static and perfusion with the formation of a thicker cell layer and greater invasion in perfusion as shown by the H&E stain. Scale bars are 100 μm. (b) After carrying out an MTT assay the membranes show the even distribution of viable cells across the membrane. (c) Results of the MTT assay for HepG2 on Alvetex Strata in static and perfused conditions show evidence of increased cell metabolism in perfusion. Values are mean
SEM, *** ¼ p < 0.05

Two applications have been used to demonstrate this technology. A liver model based around the HepG2 hepatocellular carcinoma cell line was initially developed due to the highly vascularized nature of the liver suggesting it would benefit from such a system. Further work demonstrated the utility of the system for the culture of an intestinal model, using the Caco-2 cell line cocultured with fibroblasts [8]. For the liver model a thicker tissue-like layer of HepG2 cells is formed on the surface of the Alvetex® Strata in perfused conditions, as can be visualized using Hematoxylin and eosin (H&E) staining of tissue sections (Fig. 4a). The high viability of this tissue is supported by the results of an MTT assay (Fig. 4b, c). Immunofluorescent analysis of these models demonstrates that the levels of key

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Fig. 5 Immunofluorescence staining of static and perfused 3D HepG2 models. The transporters MRP2 and MRP3 show distinct membrane staining, with an increase in perfusion. The tight junction protein claudin 2 exhibits a uniform staining of the cell membranes in both 3D static and 3D perfused culture methods. Scale bars: 50 μm

hepatic transporter proteins can be increased with the addition of perfusion, while key junctional proteins such as claudin 2 are retained with specific membrane staining in perfusion (Fig. 5). This demonstrates the potential benefits of using such a system for the culture of HepG2 hepatic models. The intestine represents another organ subjected to mechanical cues in vivo. The intestinal mucosa is exposed to peristaltic movement of food substances at the apical lumen as well as basolateral vascularization. Intestine models perfused within the bioreactor system exhibit physiological parameters that more accurately represent the in vivo microenvironment of the small intestine compared to that of static culture. Under dynamic conditions, the epithelial cells appear to form villus–crypt-like architecture. Longer-term perfusion results in a greater degree of cellular projection, as shown by the histological data (Fig. 6). Visualization of Ki67positive cells by immunofluorescence staining reveals proliferative cells are restricted to the basal area of the villus-like structures like that of the in vivo environment (Fig. 7). Epithelial polarization at the surface of the villus-like structures is evident and junctional complexes form, as demonstrated by E-cadherin expression (Fig. 7). Fibroblasts within the Alvetex® Scaffold adopt a myofibroblast phenotype as a result of perfusion, as shown by positive alpha smooth muscle actin staining. Perfusion of intestine models illustrates the potential of the bioreactor system in simulating an improved growth environment enabling more physiologically relevant tissue morphogenesis.

Fig. 6 H&E staining of in vitro intestine models grown in static and perfusion culture for 7 and 14 days. HDFn fibroblasts were cultured in Alvetex Scaffold to create a lamina propria-like compartment. Caco-2 cells were then cultured on the Scaffold for 21 days until a mature epithelial layer formed. These mature intestinal models were cultured for a further 7–14 days in static and perfused conditions. Perfused intestine models exhibit crypt–villus morphogenesis. Caco-2 cells form a polarized layer on the surface of the villus structures. Larger villus-like structures form following 14 days of perfusion compared to 7 days. Scale bars: 100 μm

Fig. 7 Immunofluorescence staining of static and perfused 3D intestine models. Static intestine models show no Ki67 staining in the epithelial layer. Perfusion of intestine models results in Ki67 positive staining restricted to the basal layer of the crypt structures, suggesting proliferation is occurring. E-cadherin staining is evident throughout the epithelial layer in both static and perfused models, indicating mature junctional complexes form between epithelial cells. Alpha smooth muscle actin staining in fibroblasts increases within the laminapropria-like compartment as a result of perfusion. Scale bars: 50 μm

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Materials Cell Culture

2.1.1 HepG2 Tissue Model

HepG2 hepatocellular carcinoma cells (ECACC). 6-well Alvetex® insert (REPROCELL). 6-well plate. Dulbecco’s Modified Eagle’s Medium (DMEM) (Thermo Fisher Scientific). Fetal bovine serum (FBS) (Thermo Fisher Scientific). Penicillin–streptomycin (Thermo Fisher Scientific). L-Glutamine

(Thermo Fisher Scientific).

Trypsin-EDTA (Thermo Fisher Scientific). Phosphate Buffered Saline. Sterile Forceps. Stirred Bioreactor Apparatus. 2.1.2 Intestinal Tissue Model

Human neonatal dermal fibroblasts (HDFn, Thermo Fisher Scientific).

Stromal Compartment

Dulbecco’s Modified Eagle Medium (DMEM, Thermo Fisher Scientific). Fetal bovine serum (FBS, Thermo Fisher Scientific). L-glutamine

(Thermo Fisher Scientific).

Nonessential amino acids (NEAA, Thermo Fisher Scientific). Penicillin–streptomycin solution (Thermo Fisher Scientific). Transforming growth factor β1 (TGFβ1, PeproTech). Ascorbic acid (Sigma-Aldrich). Trypsin-EDTA (Thermo Fisher Scientific). Phosphate buffered saline. Sterile forceps. Epithelial Compartment

Human epithelial colorectal adenocarcinoma cells (Caco-2, The European Collection for Authenticated Cell Cultures). Dulbecco’s Modified Eagle Medium (DMEM, Thermo Fisher Scientific). Fetal bovine serum (FBS, Thermo Fisher Scientific). L-Glutamine

(Thermo Fisher Scientific).

Nonessential amino acids (NEAA, Thermo Fisher Scientific). Penicillin–streptomycin solution (Thermo Fisher Scientific). Sterile forceps. Stirred bioreactor apparatus.

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100% ethanol. Serial dilutions of ethanol: 95%, 90%, 80%, 70%, 50%, 30%. Histo-Clear II. Paraffin wax. Tissue embedding cassettes. Tissue embedding molds. Microtome. Positively charged microscope slides. Distilled water. Mayer’s hematoxylin (Sigma). Alkaline ethanol (3% ammonia in 70% ethanol). Eosin. Neonatal calf serum. Triton X-100. Primary antibodies. Fluorophore-conjugated secondary antibody. Vectashield HardSet with DAPI. Glass coverslips. Thiazolyl blue tetrazolium bromide powder. Phenol-free DMEM. Isopropanol. Acidified isopropanol (2% hydrochloric acid in isopropanol). 96-well plate. Plate rocker. Absorbance microplate reader.

3

Methods

3.1 Growth of HepG2 Liver Models for Drug Toxicity Testing

3.1.1 Revival of HepG2 Cells

Culture of HepG2 in the perfusion system provides a simple method to create physiologically relevant culture conditions while maintaining ease of use and low cost. The culture of the models in this system can be summarized in a simple 3 step protocol, as shown in Fig. 8. 1. Prepare a bottle of complete Dulbecco’s modified essential medium (DMEM) made up of DMEM supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, and 100 U/mL of penicillin–streptomycin.

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Fig. 8 Protocol for setting up HepG2 bioreactor cultures in in static and perfusion. HepG2 cells are initially seeded into Alvetex inserts in a 6-well plate and cultured for a week to allow the cells to adhere and migrate across the membrane. The samples are then moved into bioreactors and placed on the stirrer unit with the perfused samples on a lane set to 100 rpm and the static samples placed on a lane which is turned off. The samples are then cultured for a further 7 days before fixation

2. Revive and culture HepG2 cells in in complete DMEM, as per the manufacturer’s instructions. 3.1.2 Preparation of Alvetex® Strata

The Alvetex® Strata must be treated with ethanol before use, to render the scaffold hydrophilic. (a) Remove the Alvetex® Strata inserts from the packaging using sterile forceps and place into a sterile petri dish or beaker containing 70% ethanol. (b) Submerge the Alvetex® Strata inserts in the 70% ethanol for a minimum of 15 min. (c) Move the Alvetex® insert into a fresh 6-well plate and add 10 mL of sterile PBS to remove all ethanol. (d) Aspirate the sterile PBS and add 5 mL of complete DMEM, bringing the liquid to the level of the Alvetex® Strata membrane.

3.1.3 Seeding HepG2 Cells onto the Alvetex® Strata

1. Trypsinize HepG2 cells using trypsin–EDTA until cell have detached. 2. Neutralize the trypsin–EDTA with complete DMEM. 3. Centrifuge the cells at 1000  g for 3 min. 4. Perform cell counts using a trypan blue exclusion assay. 5. Seed 2  106 HepG2 cells onto the prepared Alvetex® Strata inserts. 6. Incubate at 37  C in a 5% CO2 humidified incubator for 7 days with a complete medium change every 2 days.

3.1.4 Preparation of the Perfusion Bioreactors

Perfusion bioreactors need to be autoclaved prior to use to maintain sterility. (a) Take a complete bioreactor system and place an autoclavable lid to cover the top, or alternatively wrap aluminum foil over the top of the vessel.

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Fig. 9 Protocol for setting up intestinal bioreactor cultures in in static and perfusion. HepG2 cells are initially seeded into Alvetex inserts in a 6-well plate and cultured for a week to allow the cells to adhere and migrate across the membrane. The samples are then moved into bioreactors and placed on the stirrer unit with the perfused samples on a lane set to 100 rpm and the static samples placed on a lane which is turned off. The samples are then cultured for a further 7–14 days before fixation

(b) Autoclave the bioreactor for 20 min at 121  C using a standard clean cycle. (c) Store the bioreactor in a clean area until required (see Note 1). 3.1.5 Moving HepG2 Models into the Perfusion System

1. Take an autoclaved bioreactor and place in a laminar flood hood. 2. Add 100 mL of prewarmed complete DMEM to the system. 3. Aspirate the medium from the HepG2 models after 7 days of culture. 4. Using sterile forceps, lift the Alvetex® insert from the plate and place it into the holder in the bioreactor system (see Note 2). 5. Place a sterile 60 mm petri dish lid over the vessel and incubate it on a magnetic stirrer set to 100 rpm for a further 7 days in a 37  C, 5% CO2 humidified incubator (see Note 3).

3.2 Creation of Perfused Intestinal Models

3.2.1 Revival of Cells

For the intestinal models an Alvetex® Scaffold 12-well insert is used. These require lower quantities of cells compared to 6-well inserts while the use of Alvetex® Scaffold instead of Strata creates a more beneficial environment for the growth of the fibroblast compartment. The production of this model requires more user input compared to the previous HepG2 model, as shown by the diagrammatical summary of the protocol in Fig. 9. 1. Prepare a bottle of complete media by supplementing DMEM with 10% FBS, 1% NEAA, and 2 mM L-glutamine. Additional antimicrobial agents (100 U/mL penicillin and 100 μg/mL streptomycin) is also recommended. 2. Cryopreserved Caco-2 cells should be revived and cultured in complete DMEM as per the manufacturer’s instructions.

3.2.2 Preparation of Alvetex® Scaffold

The Alvetex® Scaffold must be treated with ethanol before use, to render the scaffold hydrophilic. (a) Remove the Alvetex® Scaffold inserts using sterile forceps and place into a sterile petri dish or beaker.

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(b) Submerge the Alvetex® Scaffold inserts in 70% ethanol for a minimum of 5 min. (c) Move the Alvetex® insert into a fresh 6-well plate and add sterile PBS to remove all ethanol. (d) Aspirate the sterile PBS and add 5 mL of complete DMEM supplemented with 5 ng/mL TGF-β1 and 100 μg/mL ascorbic acid, bringing the liquid to the level of the Alvetex® membrane. 3.2.3 Seeding HDFn Cells onto the Alvetex® Scaffold

1. Trypsinize HDFn cells at 80% confluency using 0.25% Trypsin– EDTA until cells have detached. 2. Neutralize the trypsin–EDTA with complete DMEM. 3. Centrifuge the cells at 200  g for 3 min. 4. Perform cell counts using a trypan blue exclusion assay. 5. Seed 5  105 HDFn cells onto the prepared Alvetex® Scaffold inserts. 6. Incubate at 37  C in a 5% CO2 humidified incubator in complete DMEM supplemented with 5 ng/mL TGF-β1 and 100 μg/mL ascorbic acid. Culture these for 14 days with a complete medium change twice weekly.

3.2.4 Seeding Caco2 Cells onto the Stromal Compartments

1. Trypsinize Caco-2 cells at 70–90% confluency using 0.25% trypsin–EDTA until cells have detached. 2. Neutralize the trypsin–EDTA with complete DMEM. 3. Centrifuge the cells at 200  g for 3 min. 4. Perform cell counts using a trypan blue exclusion assay. 5. Seed 0.4  106 Caco-2 cells onto the Alvetex® Scaffold inserts previously cultured with HDFn for 14 days. 6. Incubate at 37  C in a 5% CO2 humidified incubator for 21 days with a complete medium change every 2 days.

3.2.5 Preparation of the Perfusion Bioreactors

Perfusion bioreactors need to be autoclaved prior to use to maintain sterility. (a) Take a complete bioreactor system and place the lid in the vented orientation. (b) Autoclave the bioreactor for 20 min at 121  C using a standard clean cycle. (c) Immediately upon finish, reverse the lid of the bioreactor to the sealed orientation and store it until use.

3.2.6 Moving Intestinal Models into the Perfusion System

1. Take an autoclaved bioreactor and place in a laminar flood hood.

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2. Thoroughly clean the lid of the bioreactor with 70% ethanol prior to inverting it to the vented orientation. 3. Add 100 mL of prewarmed complete DMEM to the system. 4. Aspirate the medium from the HepG2 models after 7 days of culture. 5. Using sterile forceps, lift the Alvetex® insert from the plate and place it into the holder in the bioreactor system (see Note 2). 6. Replace the bioreactor lid and incubate it on a magnetic stirrer set to 100 rpm for a further 7–14 days in a 37  C, 5% CO2 humidified incubator. Perform a full media change on day 7 if perfusing for 14 days. 3.3 Processing and Analysis of the Perfused Models

To investigate the function and morphology of in vitro tissue equivalents, a wide range of analytical techniques can be used. The figures shown in this chapter were generated through histological and immunofluorescent techniques for visual analysis of gross morphology and expression of individual proteins, or through the use of an MTT assay to measure cellular metabolism.

3.3.1 Processing Samples for Paraffin Wax Embedding

1. Carefully unclip the Alvetex® insert using blunt forceps and place the membrane into a 12-well plate containing 2 mL of PBS. 2. Aspirate the PBS and wash a further two times to remove residual culture medium. 3. Fix in 4% paraformaldehyde at room temperature for 2 h, or overnight at 4  C. 4. Wash the fixed models three times in PBS to remove residual paraformaldehyde. 5. Dehydrate the samples through a gradient of ethanol concentrations (30%, 50%, 70%, 80%, 90%, 95%) for 15 min each before finally in 100% ethanol for 30 min. 6. Transfer the dehydrated samples into tissue processing cassettes and submerge in Histo-Clear II for 30 min. 7. Move the samples into a 1:1 mixture of Histo-Clear II and molten paraffin wax, and incubate at 65  C for 30 min. 8. Transfer the samples to pure molten paraffin wax and incubate at 65  C for 1 h. 9. Cut the tissue models in half across their diameter using a surgical scalpel and embed in molten wax with the flat edge orientated to the base of the plastic embedding mold. 10. Allow embedded models to completely set overnight at room temperature prior to sectioning and downstream analysis.

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3.3.2 Generation of Transverse Sections of Tissue Models

1. Section the wax blocks containing the tissue models at a thickness of 5–7 μm using a rotary microtome. 2. Lift the sections onto a water bath at 37  C and mount onto positively charged slides. 3. Place slides onto a heated slide rack at 30  C and leave for a minimum of 1 h prior to further processing.

3.3.3 Histological Analysis

1. Deparaffinize the tissue sections in Histo-Clear for 5 min. 2. Rehydrate the samples through sequential ethanol concentrations: 100% ethanol for 2 min followed by 95% ethanol, 70% ethanol, and distilled water for 1 min each. 3. Stain the slides with Mayer’s hematoxylin for 5 min. 4. Wash slides in distilled water for 30 s. 5. Blue the nuclei with alkaline alcohol for 30 s. 6. Dehydrate the samples through 70% ethanol and 95% ethanol for 30 s each. 7. Stain the samples with eosin for 1 min. 8. Dehydrate the samples with two washes in 95% ethanol for 10 s each followed by two washes in 100% ethanol for 15 and 30 s respectively. 9. Move the slides through two washes with Histo-Clear for 3 min each. 10. Mount a clean coverslip onto the slides using Omnimount. 11. Leave slides to dry at room temperature prior to imaging.

3.3.4 Immunofluorescent Analysis

1. Deparaffinize the tissue sections in Histo-Clear for 5 min. 2. Rehydrate the slides sequentially through 100% ethanol, 70% ethanol, and PBS for 5 min each. 3. Place the slides into citrate buffer at 95  C for 20 min to perform antigen retrieval. 4. Cool the slides slowly then perform blocking and permeabilization for 1 h using 100 μL of PBS containing 20% neonatal calf serum and 0.4% Triton X-100. 5. Wash the samples three times in PBS for 5 min each. 6. Incubate the samples for 2 h at room temperature with 100 μL of blocking solution containing the primary antibody at the relevant dilution (Table 1). 7. Wash the samples three times in PBS for 5 min each. 8. Incubate the samples for 1 h at room temperature with 100 μL of blocking solution containing the secondary antibody at the relevant dilution (Table 2). 9. Wash the samples three times in PBS for 5 min each.

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Table 1 Primary antibodies for use with immunofluorescent staining Primary antibody

Dilution

Species

Supplier

Product code

MRP2

1 in 100

Mouse

Santa Cruz

cc-71603

MRP3

1 in 100

Rabbit

Abcam

ab204322

Claudin 2

1 in 100

Rabbit

Abcam

ab53032

α-Smooth muscle actin

1 in 100

Mouse

Abcam

ab7817

E-cadherin

1 in 100

Mouse

BD Biosciences

6011082

Ki67

1 in 100

Rabbit

Abcam

ab16667

Table 2 Secondary antibodies for use with immunofluorescent staining Secondary antibody

Dilution

Supplier

Product code

Donkey anti-mouse Alexa Fluor 488

1 in 1000

Invitrogen

A21203

Donkey anti-rabbit Alexa Fluor 594

1 in 1000

Invitrogen

A21205

10. Mount a clean coverslip onto the slides using Vectashield HardSet with DAPI. 11. Allow the slides to dry in the dark at room temperature for 15 min prior to imaging with a confocal microscope (see Note 4). 3.3.5 MTT Assay

1. Make up the MTT solution with 1 mg/mL solution of thiazolyl blue tetrazolium bromide in phenol-free DMEM. 2. Carefully unclip the Alvetex® membranes and move them to a fresh 12-well plate. 3. Wash the membranes twice with PBS. 4. Add 1 mL of the MTT solution to each well. 5. Incubate the membranes for 1 h at 37  C. 6. Aspirate the MTT solution and add 1 mL of acidified isopropanol to each well. 7. Place the plates on a shaker at 120 rpm for 15 min. 8. Add 180 μL of isopropanol to wells in a 96-well plate. 9. Add 20 μL of the acidified isopropanol to the wells in the 96-well plate. 10. Read the absorbance of the wells at 570 nm using a plate reader.

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Notes 1. It is recommended to autoclave the bioreactor vessels as close as possible to the time they will be used to ensure sterility is maintained. 2. The arms of the Alvetex® insert line up with slots in the PTFE holder to ensure a stable fit with no rotation. When the insert is correctly positioned it should sit flat with minimal movement possible. 3. Growth of HepG2 models for greater time points can be carried out, with a further week of culture possible without further media changes. For longer periods than this the quantity of cells can lead to deleterious effects of the large tissue mass such as the formation of a necrotic core. 4. If samples are not going to be imaged immediately, they can be stored for several weeks in the dark at 4  C. For optimum results, it is recommended to image the samples as soon as possible.

References 1. Jensen C, Teng Y (2020) Is it time to start transitioning from 2D to 3D cell culture? Front Mol Biosci 7:1–15 2. Hansmann J, Egger D, Kasper C (2018) Advanced dynamic cell and tissue culture. Bioengineering 5(3):5–7 3. Knight E, Przyborski S (2015) Advances in 3D cell culture technologies enabling tissue-like structures to be created in vitro. J Anat 227 (6):746–756 4. Sonbol HS (2018) Extracellular matrix remodeling in human disease. J Microsc Ultrastruct 6 (3):123–128 5. Nicolas J, Magli S, Rabbachin L, Sampaolesi S, Nicotra F, Russo L (2020) 3D Extracellular

matrix mimics: fundamental concepts and role of materials chemistry to influence stem cell fate. Biomacromolecules 21(6):1968–1994 6. Ballermann BJ, Dardik A, Eng E, Liu A (1998) Shear stress and the endothelium. Kidney Int Suppl 54(67):100–108 7. Verkman AS, Dix JA (1984) Effect of unstirred layers on binding and reaction kinetics at a membrane surface. Anal Biochem 142(1):109–116 8. Darling NJ, Mobbs CL, Gonza´lez-Hau AL, Freer M, Przyborski S (2020) Bioengineering novel in vitro co-culture models that represent the human intestinal mucosa with improved Caco-2 structure and barrier function. Front Bioeng Biotechnol 8:1–15

Methods in Molecular Biology (2022) 2436: 257–266 DOI 10.1007/7651_2021_444 © Springer Science+Business Media, LLC 2021 Published online: 02 November 2021

Bioreactor-Based Adherent Cells Harvesting from Microcarriers with 3D Printed Inertial Microfluidics Lin Ding, Reza Moloudi, and Majid Ebrahimi Warkiani Abstract Harvesting adherent cells from microcarriers has become one of the major challenges of the downstream bioprocessing at large scale the current method has high maintenance and operation cost, which are the results of frequent clogging, due to cell lysing effect and microcarrier cake formation on membrane-based technology. These problems hugely impede the adaptation of microcarriers technologies in large-scale cell culture and hampered the supply of cells to the clinical need. Here, we describe two 3D printing-based methods to fabricate inertial microfluidic devices for separating adherent cells from microcarriers which overcome the above-mentioned limitations. The spiral devices are employed to separate mesenchymal stem cells from the microcarriers with 99% microcarrier removal rate and 77% cell recovery rate in one round of separation. Key words Stem cells, Inertial microfluidics, Cell harvesting, Microcarrier-based culture, Cell therapy industry

1

Introduction Microcarrier-based cell culture is considered as the future standard of adherent cells culture in large-scale [1, 2]. They have large surface area-to-volume ratio for the cells to attach and provide a better environment for cells interaction and secretion [1]. However, multilayer flasks are still the most common way of adherent cells production. The slow adaptation of new technology can be mainly attributed to the complicated cell harvesting procedure of microcarrier-based culture method. Harvesting cells from microcarriers heavily relies on membrane-based technologies Cells and microcarriers solution is passed through the physical filters which are frequently blocked by cell clumps, results in cell lysis and are expensive to change and operate [2]. One of the potential substitutions of membrane-based technologies is inertial microfluidic devices. Inertial microfluidics has been used widely for separation of cells from a heterogeneous population recently. It focuses particles with different sizes at different cross-sectional positions inside the channel, allowing the particles to be collected from different outlets [3, 4]. In a straight

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channel, this is achieved by the balance of two forces: (1) sheargradient inertial lift force induced by the velocity gradient of the fluid, and (2) wall-induced lift force originated from wall lubrication effect and an asymmetry in pressure distribution around the particle adjacent to the wall [5]. The balance between these two forces develops particle equilibrium positions in the channel, depending on the channel cross-sectional shape and the corresponding cross-laterally wall-effect lift force. Figure 1a illustrates four equilibrium positions of particles with a particle size a, for a microchannel with a square cross section. The net inertial lift force is a power-law function of particle size (FL ~ an, n > 1). When the particle size relative to the channel hydraulic diameter (a/DH) is greater than 0.07, the inertial focusing of neutrally buoyant particles can be guaranteed [6]. Inertial microfluidics is divided into four types according to their geometries: straight channels, serpentine channels, contraction–expansion channels, and spiral channels [7]. Adding curvature to the channel generates a secondary flow perpendicular to the main flow due to the centrifugal force. The secondary drag force scales linearly with particle size (FD ~ a) [8], where coupling it with the net inertial lift force leads to particle size-based differentially equilibriums (Fig. 1b). Among these structured channels, spiral microfluidic channels with trapezoidal cross-sections have the highest throughput in a single unit, with simple settings and high recovery rate. These features enable the spiral channels as potential candidates for large-volume liquid processing through multiplexing [9, 10]. It has been demonstrated for cell retention in perfusion bioreactors [11], as well as microcarrier-based adherent cells harvesting and microcarrier-cell complex retention in a perfusion condition [8, 12]. It was showed that the sorted cells express their normal surface markers, maintained the spindle morphology with uncompromised growth kinetics, differentiation potency and therapeutic properties after being processed by these devices. Compared to the traditional harvesting technologies, the membrane technology, spiral inertial microfluidic devices possess low-cost attributes due to ease of fabrication and maintenance. There is no need for frequent replacement due to clogging, which reduces the cost and the risk of contaminations. The throughput can be scaled out significantly by paralleling the spiral channels, while the footprint of the whole setup remains relatively small. Here, we presented the 3D printing technology-based method for rapid and low-cost fabrication of spiral microfluidic devices through (1) direct printing of the microchannels and (2) printing the master mold for soft lithography-based channel fabrication. We demonstrated that this method is capable of fabricating microfluidic devices within 24 h and subsequently separate adherent cells from microcarriers under high-throughput and scalable manner. We have achieved 77% cell recovery rate with 99% microcarriers

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Fig. 1 The inertial focusing of microparticles inside (a) straight microfluidic channels with square cross sections and (b) spiral microchannels with trapezoidal cross sections. The particle equilibrium positions are highly affected by the geometry of the channels. (c) The two methods of making the microfluidic channels for MSCs, microcarriers separation

removal rate in one round of separation. This method enables a low-cost and rapid fabrication process for the bioprocessing filtration devices and is capable of separating microcarriers in size range of 100–300 μm.

2 2.1

Materials 3D Printing

1. High resolution Digital Light Processing (DLP)/Stereolithography (SLR) 3D printer (see Note 1), filled with corresponding light-sensitive resin. 2. Scalpel for removing printed chip from the platform. 3. 100% isopropyl alcohol (IPA) (see Note 2). 4. UV curing machine.

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5. Double-sided adhesive tape (AR clear, adhesive biotech). 6. Polymethyl methacrylate (PMMA) sheet (see Note 3). 7. Scotch tape for cleaning the printed chip. 8. Air nozzle or hairdryer. 2.2 PDMS Device Making (in Replacement of DirectPrinted Device)

1. Trichloro(1H, 1H, 2H, 2H-perfluoro-octyl)silane to treat the surface of the mold, for easier device peel off. 2. Polydimethylsiloxane (PDMS), a biocompatible material for chip fabrication. 3. 45
C oven for PDMS solidification. 4. 3 mm Biopsy punches for making holes in the PDMS devices. 5. 1/1600 Rubber tubing (Tygon ND-100-65, USA) for introducing samples into and out of the device.

2.3

Cell Harvesting

1. 70% ethanol. 2. Dulbecco’s phosphate-buffered saline (DPBS). 3. Digestive enzyme (TrypLE express, Gibco, Invitrogen). 4. Syringe pump (see Note 4). 5. 50 mL disposable syringe. 6. Silicone tubing (inner diameter: 0.5000 , outer diameter: 0.9000 ). 7. Syringe tips (0.5000 Long Tip, Adhesive Dispensing Ltd., match the size of tubes). 8. Biosafety cabinet. 9. 50 mL falcon tubes. 10. Peristaltic pump (in replacement of syringe pump). 11. Rubber tube (in replacement of Tygon tubes).

3 3.1

Methods Device Design

Design the spiral microfluidic channels with CAD software (e.g., AutoCAD, Solidworks), export the “.Stl” file. Proceed the file in the printer software for slicing and covert to printing file and set to print (see Note 5). For direct printed device, the spiral design is consisted of 6 revolutions, 1.7 pitches, 550 μm inner wall height, 620 μm outer wall height and 1100 μm width inward trapezoidal spiral microchannel (see Note 6). For mold fabrication, the mold of the spiral was designed as an outward spiral with 5 revolutions, 500 μm inner wall height, 650 μm outer wall height and 4000 μm width trapezoidal spiral, the pitch of the Archimedean spiral was 7 mm (3 mm interval between successive spiral loops) [12]. Depending

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on the printing area of the printer, different designs can be chosen (see Note 7). The fabrication methods were summarized in Fig. 1c. 3.2 Direct Fabrication

1. The detail of the direct fabrication method was described in [13]. 2. Remove the printed spiral chip carefully from the pick-up platform with scalpel. 3. Wash the chip with IPA and dried with an air nozzle or hairdryer. Repeat the washing process for at least three times until there is no visible uncured resin. 4. Following washing, cure the spiral chip in the UV curing chamber for 2 min to further solidify the resin. 5. Tape the chip with Scotch tape to remove potentially available dust (see Note 8). 6. Bind the chip on PMMA-base with double-sided adhesive tape. 7. Press the chip and substrate together with force, make sure the chips are well attached to the substrate (see Note 9). 8. Place the silicone tubing inside the inlet (see Note 10) and outlets (see Note 11) of the spiral chip.

3.3

Mold Fabrication

1. The detail of the mold fabrication method was described in [14]. 2. Remove the printed spiral mold from the pick-up platform by scalpel. 3. Wash carefully with IPA and dried with an air nozzle or hairdryer. Repeat the washing process for at least three times until there is no visible uncured resin. 4. Cure the spiral mold in the UV curing chamber for 2 min to further solidify the resin. 5. Dip the mold in the IPA for at least 6 h to clean up the surface of the mold. 6. Put the spiral mold in the plasma machine, vacuum for 4 min, and plasma treat for 2 min. 7. After plasma treatment, Place the treated mold immediately in vacuum chamber. Add 100 μL of Trichloro (1H, 1H, 2H, 2H-perfluoro-octyl) silane in a small container inside the vacuum chamber, vacuum for 2 min and leave the mold inside for at least 8 h. These treatments allow a layer of silane to coat on the surface of the mold, helping PDMS device to be peeled off from the mold easier. 8. Prepare complete PDMS mixture by mixing the PDMS reagents and curing agent at 10:1 ratio, mix thoroughly for 5 min (see Note 12). Degas the PDMS in a vacuum pot until there is no bubble left.

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9. Cast PDMS mixture on the mold and degas the mold with PDMS in the vacuum pot until there is no bubble left in the PDMS. Cast PDMS on a petri dish to create a layer of PDMS substrate for device binding as well. 10. Leave the mold in 45
C oven until the PDMS device is cured (see Note 13). 11. Peel off the device from the mold. 12. Punch the inlet and outlet holes with the 3 mm biopsy punch, carefully clean the punching holes by flushing IPA through it and dry with an air nozzle. 13. Tape the chip with scotch tape to remove dust and debris. 14. Cut the PDMS substrate out of the petri dish, use the bottom side as substrate and leave it in the plasma machine with the PDMS device (channel side facing up). Vacuum for 4 min and plasma treat for 2 min in the machine (see Note 14) and bind them together. 15. Place the silicone tubing inside the inlet and outlets of the spiral chip. 3.4

Operation

Carry out all processes of separation/harvesting in a biosafety cabinet to reduce pathogen contamination risks (Fig. 2). 1. Spray 70% ethanol on all equipment and devices, move them into the biosafety cabinet and leave them under UV light for 30 min. 2. Use one syringe to inject 70% ethanol into the device for sterilization of the system for 5 min. 3. Use another syringe to inject DPBS into the device, wash away the 70% ethanol and perfuse for 10 min. Keep the perfused, sterile DPBS inside the channel. 4. Detach the cells from microcarriers by removing the supernatant as much as possible with a serological pipette, and add enough digestive enzymes (e.g., TrypLE express, Gibco) to cover all microcarriers and incubate for 10–15 min in the incubator, depending on the concentration and cell density. 5. Resuspend the cells and microcarriers for 10 times with a serological pipette. 6. Dilute the cells to certain concentration with DPBS: up to 0.75% v/v ratio for direct-printed chip design and 1.68% v/v ratio for PDMS chip design. 7. Load the cell-microcarrier solution into the syringe when using a syringe pump or leave it inside the bioreactor/spinner flask or a reservoir with a magnetic stirrer if the solution is delivered by a peristaltic pump.

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Fig. 2 Running the spiral microfluidic chips with different pumping systems. (a) The schematic illustration of the simple system setup. (b) The direct printed resin chip with a Syringe pump and (c) the PDMS chip with peristaltic pump. Direct-printed chip and PDMS chip can be operated with both pumps’ settings. The scalebars in (b) and (c) are 250 μm and 1 mm, respectively. (Reproduced from [12] with permission from Nature)

8. Pump the solution into the spiral microfluidic chip and collect the separated cells/microcarriers from the respective outlets in 50-mL tubes (Fig. 3b, c). The operation flow rate is 3 mL/ min and 30 mL/min for the direct-printed and PDMS chip, respectively (see Note 14). 9. Extra rounds of separation can be performed to increase the desired yield. Inner outlet solution can be reintroduced into the device to harvest more cells. The yield of cells depends on the inlet-outlet ratio. In this work, the cell recovery rate in one round can be as high as 77% and reach ~95% in two rounds of separation with the cell properties preserved (Fig. 3).

4

Notes 1. The printing resolution of the printer affects the geometry and surface roughness of the mold and channel, thus altering the inertial focusing attribute. 2. Washing the device/mold by 100% ethanol if the remaining resin is hard to clean. The printed part can also be cleaned by sonicating the part in a flask of IPA/ethanol for 5 min. 3. PMMA sheet is a transparent and rigid plastic sheet used as a base for binding. It can be replaced with any flat surface material. Depending on the printer, protocol and settings, some

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Fig. 3 The separation results of the PDMS-made device. (a) The separated microcarriers from the inner outlet and (b) harvested cells from the outer outlet respectively. The scale bar is 200 μm. (c) The viability of cells after running through the device under the high flow rate of 30 mL/min shows no significant drop even after two rounds of separation. (d) The cells were showed to preserve their differentiation potential by trilineage differentiation. (Reproduced from [12] with permission from Nature)

printers might be able to print the microfluidic channel directly without binding the channel to another substrate [15]. The method we described here has lower requirement of expertise and printer resolution, making the device more accessible to general laboratories. 4. In general, syringe pump is less preferred for this application due to the fast deposition of microcarriers and noncontinuous manner of separation in large-scale. 5. The SLA printer recommended for this purpose is Formlab form 3, and the DLP printer recommended is MIICRAFT. Higher printing resolution gives smoother geometry. Rough surface of the device and mold will disrupt the flow and affect the separation efficiency of the device. 6. The inlet of the direct-printed chip was placed at a 60
angle to prevent blockage of microcarriers. The inlet and outlet sizes of

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the spiral were designed according to the size of tubings. In our case, we used 2.278 mm diameter inlet and outlet holes. The inner outlet has a width of 450 μm, and the outer outlet has a width of 650 μm. 7. There dimensions of the spirals are not limited to the two designs we described here. Users can customize and test their own designs easily with this protocol. The two designs we suggested here were tested in our previous works. Due to the flexibility of PDMS, the PDMS chip and direct printed chip with same dimension might not work at the same flow rate, which need to be calibrated. 8. Fabrication in a cleanroom environment reduces the risk of having device defects caused by particulates and dust. Printing quality also directly affect the integrity of device. Make sure the resin tank of the printer is always free of debris and dust. 9. Only peel off the protective layer on PMMA sheet right before binding the chip to avoid dust in the channel. 10. Glue resin around the inlet tube and cure it to prevent leakage. 11. The length of outlets tubings need to be the same. Otherwise, it causes an imbalanced pressure between the two outlets and consequently affects the focusing position and separation efficiency of the device. 12. Incomplete mixing of PDMS with the curing agent results in uncured PDMS left on the surface of the mold during casting. The mold will be hard to clean afterward as well. Therefore, sufficient PDMS mixing is critical. 13. Resin mold might crack under high temperature, depending on the type of resin. So it is recommended to cure the PDMS chips in a low-temperature oven. However, after the first cure in the 45
C oven, the mold will be able to resist at least 65
C temperature without cracking (heat treatment). 14. PDMS-made channels fabricated from resin mold are weaker in plasma bonding compared to micromilling or silicon wafer mold. Binding PDMS with other materials, such as glasses, can improve the bonding strength if leakage happens. 15. The inertial focusing of particles takes a few seconds to reach steady state and get focused in the desired positions in the channel. During the transition period when pumps start and stop the particles are randomly dispersed in the channel.

Acknowledgments M. E. W. would like to acknowledge the support of the Australian Research Council through Discovery Project Grants

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(DP170103704 and DP180103003) and the National Health and Medical Research Council through the Career Development Fellowship (APP1143377). The authors declare no known competing interest. References 1. Tavassoli H et al (2018) Large-scale production of stem cells utilizing microcarriers: a biomaterials engineering perspective from academic research to commercialized products. Biomaterials 181:333–346 2. Jossen V et al (2018) Manufacturing human mesenchymal stem cells at clinical scale: process and regulatory challenges. Appl Microbiol Biotechnol 102(9):3981–3994 3. Zhang J et al (2016) Fundamentals and applications of inertial microfluidics: a review. Lab Chip 16(1):10–34 4. Amini H, Lee W, Di Carlo D (2014) Inertial microfluidic physics. Lab Chip 14 (15):2739–2761 5. Moloudi R et al (2018) Inertial particle focusing dynamics in a trapezoidal straight microchannel: application to particle filtration. Microfluid Nanofluid 22(3):33 6. Di Carlo D et al (2007) Continuous inertial focusing, ordering, and separation of particles in microchannels. Proc Natl Acad Sci U S A 104(48):18892–18897 7. Tang W et al (2020) Channel innovations for inertial microfluidics. Lab Chip 20 (19):3485–3502

8. Moloudi R et al (2019) Scaled-up inertial microfluidics: retention system for microcarrier-based suspension cultures. Biotechnol J 14(5):1800674 9. Rafeie M et al (2016) Multiplexing slanted spiral microchannels for ultra-fast blood plasma separation. Lab Chip 16(15):2791–2802 10. Warkiani ME et al (2015) Membrane-less microfiltration using inertial microfluidics. Sci Rep 5:11018 11. Kwon T et al (2017) Microfluidic cell retention device for perfusion of mammalian suspension culture. Sci Rep 7(1):1–11 12. Moloudi R et al (2018) Inertial-based filtration method for removal of microcarriers from mesenchymal stem cell suspensions. Sci Rep 8 (1):12481 13. Bazaz SR et al (2020) 3D printing of inertial microfluidic devices. Sci Rep 10(1):1–14 14. Shrestha J et al (2019) A rapidly prototyped lung-on-a-chip model using 3D-printed molds. Organs-on-a-Chip 1:100001 15. Vasilescu SA et al (2020) 3D printing enables the rapid prototyping of modular microfluidic devices for particle conjugation. Appl Mater Today 20:100726

INDEX A Acetylated low density lipoprotein (AcLDL) uptake assay .............................................................. 216 Achilles tendon.............................................................. 138 Aerobic microorganisms ................................................. 17 Allogenic therapies .......................................................... 87 Alvetex® membranes ........................................... 243, 244, 251–253, 256 Anchorage-dependent cells .......................................... 155 Animal derived component free (ADCF) ........................ 2 Antibiotics .................................. 170–173, 176, 179–181 Arginine-glycine-aspartate (RGD) sequence................. 85 Autologous/allogeneic therapeutic approaches............ 84

B Bacterial nanocellulose-based grafts, for cell colonization studies ................................................. 205–207 materials antibodies .......................................................... 208 assays and staining solutions............................. 208 bioreactor setup.........................................207–208 cell culture ......................................................... 208 chemicals and solutions .................................... 208 consumables ...................................................... 209 hardware ....................................................209–210 methods AcLDL uptake assay.......................................... 216 acridine orange staining.................................... 218 bioreactor setup.........................................210–211 CD31 immunofluorescence staining .......218–219 cell seeding ................................................211–212 considerations.................................................... 210 lactate monitoring.....................................212–213 Phalloidin—FActin staining .....................216–218 small diameter vascular grafts, connection ...... 211 termination of experiment................................ 216 WST-1 proliferation assay .........................213–215 Bambanker™................................................................. 107 BioBLU® 0.3c bioreactor............................................... 86 BioBLU® 5c stirred bioreactor ...................................... 91 Biological scaffolds........................................................ 180 Biomanufacturing of human stem cells .............. 193, 194 Bioreactor ............................................................. 127, 243 cell cultivation ............................................................. 2

commercially available single-use bioreactors ......... 84 gas transfer................................................................. 13 macrosparger ............................................................... 3 pitched blade impellers ............................................... 3 preparation .................................................................. 7 setpoints................................................................... 7–8 single-use bioreactors................................................ 84 stirred tank (see Stirred tank reactors (STR)) Bioreactor-based adherent cells harvesting ........ 257–259 3D printing technology-based method ................. 258 inertial microfluidics ............................................... 258 materials cell harvesting.................................................... 260 3D printing................................................259–260 PDMS device..................................................... 260 membrane-based technologies ............................... 257 methods device design .............................................260–261 direct fabrication ............................................... 261 mold fabrication ........................................261–262 operation....................................................262–263 PDMS ...................................................................... 265 PMMA sheet ........................................................... 263 SLA printer .............................................................. 264 Biosafety cabinet (BSC) ................................................ 168

C Caco-2 cell line.............................................................. 245 Carbon cataboite repression (CCR) ............................ 224 Cardiomyocytes............................................................... 78 Cascaded continuous cultivation......................... 223–225 materials host cells ............................................................ 225 required devices or cultivation .................225–227 required equipment for process analysis .................................................227–228 required media .................................................. 225 methods biomass determination..............................233–234 continuous adaptation phase ....................230–232 cultivation setup ........................................228–229 determination of metabolite accumulation ..... 234 flow rates calculation......................................... 235 induction phase ................................................. 232

Kursad Turksen (ed.), Bioreactors in Stem Cell Biology: Methods and Protocols, Methods in Molecular Biology, vol. 2436, https://doi.org/10.1007/978-1-0716-2018-2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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268 Index

preculture and batch phase.......................229–230 product determinatio ........................................ 235 substrate uptake rates........................................ 235 Cedex Bio Analyzer....................................................... 105 CelGro® scaffold .................................................. 136, 140 Cell activity, in 3D scaffolds bioartificial tissues ................................................... 127 bioreactor system ........................................... 129–130 mechanical compression load and fluid flow parameters.................................................... 131 sample preparation .................................................. 130 sample transfer and chamber assembly .................. 131 Cell adhesion ................................................................... 85 Cell attachment and matrix synthesis .......................... 131 Cell attachment efficiency............................................... 88 Cellbag container ................................149–152, 154, 155 Cell colonization studies, bacterial nanocellulose-based grafts for .............................................. 205–207 materials antibodies .......................................................... 208 assays and staining solutions............................. 208 bioreactor setup.........................................207–208 cell culture ......................................................... 208 chemicals and solutions .................................... 208 consumables ...................................................... 209 hardware ....................................................209–210 methods AcLDL uptake assay.......................................... 216 acridine orange staining.................................... 218 bioreactor setup.........................................210–211 CD31 immunofluorescence staining .......218–219 cell seeding ................................................211–212 considerations.................................................... 210 lactate monitoring.....................................212–213 Phalloidin—FActin staining .....................216–218 small diameter vascular grafts, connection ...... 211 termination of experiment................................ 216 WST-1 proliferation assay .........................213–215 Cell culture medium composition ................................. 88 Cell expansion ...........................................................73–75 Cell growth...................................................................... 85 Cell harvesting............................................................... 260 CELLine AD 1000 bioreactor flask adapting cells, to long-term bioreactor media................................................... 186–187 cell culture ............................................................... 185 characterization materials ....................................... 185 EVs characterization .........................................188–189 harvesting .......................................................... 187 isolation and purification ......................... 188, 191 isolation materials.............................................. 185 RNA ................................................................... 190 illustration................................................................ 184

imaging bioreactor growth surface ............... 189–190 inoculation of cells ......................................... 185–186 preparation .............................................................. 185 shed cells ......................................................... 187, 188 ultrafiltration or multiple ultracentrifuge spins.............................................................. 183 Centrifugation ...................................................... 197, 199 Chemostat cultivation.......................................... 223, 224 Chinese hamster ovary (CHO) cells ................................ 1 Chondrocytes, rocked disposable bioreactor advantages................................................................ 145 cellbag container with inlet and outlet port.......... 150 cell culture ............................................................... 148 cell density and viability .......................................... 153 cell staining .............................................................. 148 CP5 cell inoculum preparation ..................... 147–148 DO and pH level..................................................... 154 glucose consumption rate......................149, 153–154 intracellular oxidoreductases ......................... 148–149 lactate dehydrogenase .................................... 149, 154 maintaining CP5 cells in bioreactor....................... 152 metabolic activity of cells adhered to microcarriers ................................................ 153 microcarrier conditioning ....................................... 147 preparation of CP5 chondrocyte inoculum ............................................. 151–152 ReadyToProcess WAVETM 25 bioreactor system.................................................. 149–150 rehydration, conditioning, and sterilization of microcarriers ................................................ 151 samples for analysis ........................................ 148, 152 Classic fed-batch cultivation ............................................. 2 CLAVE™ sampling port ............................................... 155 Commercially available single-use bioreactors .............. 84 Compression bioreactors .............................................. 128 Conventional continuous process times ...................... 224 CP5 chondrocytes............................................... 146, 147, 151, 152, 154, 155 Cryovials ............................................................................ 2 Cultivation container ...................................................... 84 Cytodex 3 microcarriers .....................146, 147, 149, 151

D Decellularization .................................158, 164, 167, 168 Decellularized tracheal ECM-based porcine bio-scaffold generation....................................................... 160–163 repopulation ...........................................161–162, 164 Detergent circuitries ............................................ 173–174 Dissolved oxygen (DO) concentration, STR aeration ...................................................................... 18 bioreactor.............................................................19, 20 chronometer .............................................................. 20 dynamic method .................................................23, 24 electrode of................................................................ 24

BIOREACTORS growth media ............................................................ 20 microorganism ............................................. 19, 21, 23 oxygen transfer coefficient.................................. 21–22 spectrophotometer .................................................... 20 steady-state technique............................................... 19 sterilization process ................................................... 21 unsteady-state technique .................................... 18–19 Double-chamber perfusion bioreactor ............... 168, 169

E E. coli BL21 (DE3) ....................................................... 224 Eppendorf BioBLU® series ............................................ 86 Exosomes................................................................ 83, 193 Extracellular matrix (ECM)...........................85, 167, 241 Extracellular vesicles (EVs) bioreactor preparation ............................................ 196 CELLine bioreactor (see CELLine AD 1000 bioreactor flask) hiPSC-NPCs............................................................ 194 hiPSCs (see Human induced pluripotent stem cells (hiPSCs)) hMSCs (see Human mesenchymal stromal/stem cells (hMSCs)) hUC-MSCs.............................................................. 194 HUVECs ................................................................. 194 isolation from collected media ............................... 201 isolation from human stem cells grown in bioreactor cultures......................................................... 196 PBS-VW bioreactors ............................................... 194 wave bioreactor ....................................................... 194

F Fabrication protocol, thermoplastic microfluidic devices biological and medical applications.......................... 27 chip design and preparation electrochemical wet etching ............................... 32 photolithography .......................................... 29–31 using AutoCAD software ................................... 31 drilling and planarization.......................................... 34 of electrodes blank COP substrate .............................. 32–33, 36 Cr/Au etching .................................................... 33 design creation .................................................... 32 hot embossing ..................................................... 34–36 Market and Market reports ...................................... 28 materials..................................................................... 29 microfluidic device with electrodes .......................... 36 microfluidic device without electrodes .............. 35–36 modification and durability ...................................... 28 optical properties....................................................... 28 thermo-compression bonding .................................. 35 FiberCell bioreactor ...................................................... 184 Fibroblasts ..................................................................... 246

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Flow cytometry analysis .................................................. 90 Fluid flow....................................................................... 242

G Gas transfer................................................................13, 17 Geltrex coating .............................................................. 201 Glucose consumption ..................................................... 85 Glucose consumption rate...........................149, 153–154 Good manufacturing practice (GMP) ........................... 84

H HepG2 tissue model ............................................ 245, 248 hiPSC complete culture medium (hiPSC-CCM)........ 195 hiPSC-derived neural progenitor cells (hiPSC-NPCs) .................................... 194, 202 hMSC complete culture medium (hMSC-CCM) .................................... 195–196 Horseradish peroxidase (HRP) .................................... 191 Human induced pluripotent stem cells (hiPSCs) in BioBLU® ............................................................... 93 chemically defined, serum-free expansion ......... 87–89 Corning’s spinner flasks ...................................... 94–97 culture medium, preparation of ............................... 70 differentiation in planar and bioreactor cultures......................................................... 195 differentiation of NPC organoid in spinner flasks .................................................... 197–198 enzymatic dissociation of........................................ 106 in Eppendorf’s Instrumented BioBLU® 0.3c.........................................................97–100 equipment and consumables .................................... 68 extracellular matrices, preparation of ................. 69–70 media and consumables ............................................ 68 microcarrier cultures media and small molecules for ........................... 69 preparation of ................................................ 70–71 reagents and consumables for ............................ 69 Ns1u criterion........................................................... 107 planar culture, culture and expansion in....... 196–197 process analytics ........................................................ 93 sampling and quality control ......................... 100–104 screening for high cardiac differentiation potency..................................................... 72–73 single-use spinner flasks ............................................ 92 small molecules, preparation of ................................ 70 in spinner flasks ......................................................... 93 in stirred single-use bioreactors.......................... 84–87 T75-flask-based inoculum production............... 93–94 in T75-flasks .............................................................. 92 Human mesenchymal stromal/stem cells (hMSCs)......................................................... 83 expansion in PBS-VW bioreactors ................ 199–200

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270 Index

Human mesenchymal stromal/stem cells (hMSCs) (cont.) expansion in planar and bioreactor cultures................................................ 195–196 expansion in planar culture..................................... 199 Human pluripotent stem cells (hPSCs) random positioning machine APLNR+ mesoderm cells .................................... 56 bioreactor equipment ......................................... 57 cell culture medium and reagents ...................... 57 cell lines ............................................................... 57 electronic control system .................................... 65 feeder-free expansion .......................................... 59 flow cytometry .................................................... 64 hematopoietic differentiation .......................55, 56 hematopoietic progenitor cells ..................... 61–62 hemogenic endothelium progenitor ............ 60–61 IF9S medium formulation.................................. 58 immunofluorescence staining ....................... 62–64 immunostaining and flow cytometry ................. 59 Matrigel® solution .............................................. 64 mesoderm induction ........................................... 60 T12.5 cell culture flasks ...................................... 59 stirred tank bioreactors autoclave cycle ..................................................... 50 bioreactor....................................................... 41–42 bioreactor setup............................................. 44–46 buffers and reagents ............................................ 40 cardiomyoyctes .................................................... 40 cell-based therapies ............................................. 39 cell count during expansion ......................... 47–48 cell harvest ..................................................... 48–49 cell inoculum ....................................................... 46 C-flex tubing ....................................................... 46 DeltaV control platform ..................................... 51 DO probe ............................................................ 51 extracellular matrix coating ................................ 43 F3 hPSC passaging solution .........................49, 52 harvest line extension.......................................... 42 L7™ culture system ................................ 40, 49, 50 microcarrier coating ............................................ 44 passaging solution ............................................... 40 Pharmed tubing portion..................................... 47 T-75 flask ............................................................. 43 tubing ............................................................ 40–41 2D seed train ................................................. 43–44 using peristaltic pump......................................... 52 Human umbilical cord-derived MSCs (hUC-MSCs) ...................................... 194, 200 Human umbilical cord MSC-secreted EVs ................. 194 Human umbilical vein endothelial cells (HUVECs)................................................... 194

I Inertial microfluidics ..................................................... 258 Intestinal mucosa .......................................................... 246 Intestinal tissue model .................................................. 248 In vitro bioreactor perfusion model ................... 205–207 materials antibodies .......................................................... 208 assays and staining solutions............................. 208 bioreactor setup.........................................207–208 cell culture ......................................................... 208 chemicals and solutions .................................... 208 consumables ...................................................... 209 hardware ....................................................209–210 methods AcLDL uptake assay.......................................... 216 acridine orange staining.................................... 218 bioreactor setup.........................................210–211 CD31 immunofluorescence staining .......218–219 cell seeding ................................................211–212 considerations.................................................... 210 lactate monitoring.....................................212–213 Phalloidin—FActin staining .....................216–218 small diameter vascular grafts, connection ...... 211 termination of experiment................................ 216 WST-1 proliferation assay .........................213–215 In vitro 3D mechanical stimulation of TDSCs ............................................ 135–141 Isopropyl-α-D-1-thiogalactopyranoside (IPTG) ........ 224

L Lactate dehydrogenase (LDH) ........................... 149, 154 Lactic acid ...................................................................... 242 Lactose Dehydrogenase Elevating Virus (LDEV) ........................................................ 195 Long-term stable productivity ..................................... 224

M Mammalian cell culture, bioreactors antifoaming agents ...................................................... 5 automatization and detailed control ........................ 11 bioreactors ................................................................... 3 gas transfer........................................................... 13 preparation .............................................................7 setpoints............................................................. 7–8 cell culture media ................................................ 3–5, 9 cell expansion .............................................................. 7 controllers.................................................................... 5 equipment.................................................................... 3 feeding strategy ......................................................... 10 feeds and supplements ............................................ 3–5 harvest and clarification ............................................ 10

BIOREACTORS inoculation................................................................... 9 metabolite analyzers.................................................. 11 paper autoclaving bags.............................................. 12 parameters ................................................................... 6 probe calibration and sterilization.............................. 7 probes .......................................................................... 5 sampling port .............................................................. 4 seeding cell density.................................................... 13 supplements ................................................................. 6 tubing welders ........................................................... 11 Mature cardiomyocytes (CM) ........................................ 67 Mechanical stimulation ................................................. 128 Mechanobiology ........................................................... 135 Media composition ......................................................... 85 Medium adaptability ....................................................... 88 Membrane-based technologies..................................... 257 Microbial continuous cultivation ................................. 224 Microcarrier-based cell culture ..................................... 257 Microcarrier beads ...................................... 146, 151, 155 Microcarrier spinner cultures cardiac differentiation in ..................................... 75–77 media and small molecules for ................................. 69 preparation of ...................................................... 70–71 proper suspension of ................................................. 77 reagents and consumables for .................................. 69 in suspension and ensure adequate mass transfer ........................................................... 85 MTT assay ..................................................................... 245 Mycoplasma............................................................ 88, 191

P Partial decellularization................................................. 168 Patellar tendon .............................................................. 138 PBS vertical wheel (PBS-VW) bioreactors..................194, 196, 199, 200 PEG6000..................................................... 196, 201, 202 Perfusion............................................................... 242, 243 Poisson distribution ........................................................ 85 Polydimethylsiloxane (PDMS) ............................ 260–265 Polyethylene terephthalate (PET)................................ 190 Polytetrafluoroethylene (PTFE)-coated magnetic stir bar................................................................. 243 Population doubling level (PDL) .................................. 90 PureLink™ RNA Mini Kit ............................................ 143

R ReadyToProcess WAVE 25 disposable bioreactor ............................................ 146–150 Recellularization............................................................ 167 Rocked disposable bioreactor, chondrocytes advantages................................................................ 145 cellbag container with inlet and outlet port.......... 150 cell culture ............................................................... 148

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cell density and viability .......................................... 153 cell staining .............................................................. 148 CP5 cell inoculum preparation ..................... 147–148 DO and pH level..................................................... 154 glucose consumption rate......................149, 153–154 intracellular oxidoreductases ......................... 148–149 lactate dehydrogenase .................................... 149, 154 maintaining CP5 cells in bioreactor....................... 152 metabolic activity of cells adhered to microcarriers............................................ 153 microcarrier conditioning ....................................... 147 preparation of CP5 chondrocyte inoculum ............................................. 151–152 ReadyToProcess WAVETM 25 bioreactor system.................................................. 149–150 rehydration, conditioning, and sterilization of microcarriers ................................................ 151 samples for analysis ........................................ 148, 152 Rotating bioreactor..................................... 158, 159, 161 RT-qPCR analysis ......................................................... 142

S Saccharomyces cerevisiae cells........................................... 37 Scaffold-based method ................................................. 136 Shed cells ..................................................... 187, 188, 191 Single-use bioreactors ..................................................... 84 Sodium dodecyl sulfate (SDS)............................. 168, 172 Spheroid-based cultures.................................................. 87 Stem cell expansions ....................................................... 90 Stirred bioreactor system .............................................. 243 Stirred tank reactors (STR) DO concentration aeration ................................................................ 18 bioreactor.......................................................19, 20 chronometer ........................................................ 20 dynamic method ...........................................23, 24 electrode of.......................................................... 24 growth media ...................................................... 20 microorganism ....................................... 19, 21, 23 oxygen transfer coefficient ............................ 21–22 spectrophotometer.............................................. 20 steady-state technique......................................... 19 sterilization process ............................................. 21 unsteady-state technique .............................. 18–19 human pluripotent stem cells autoclave cycle ..................................................... 50 bioreactor....................................................... 41–42 bioreactor setup............................................. 44–46 buffers and reagents ............................................ 40 cardiomyoyctes .................................................... 40 cell-based therapies ............................................. 39 cell count during expansion ......................... 47–48 cell harvest ..................................................... 48–49 cell inoculum ....................................................... 46

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272 Index

C-flex tubing ....................................................... 46 DeltaV control platform ..................................... 51 DO probe ............................................................ 51 extracellular matrix coating ................................ 43 F3 hPSC passaging solution .........................49, 52 harvest line extension.......................................... 42 L7™ culture system ................................ 40, 49, 50 microcarrier coating ............................................ 44 passaging solution ............................................... 40 Pharmed tubing portion..................................... 47 T-75 flask ............................................................. 43 tubing ............................................................ 40–41 2D seed train ................................................. 43–44 using peristaltic pump......................................... 52 Suspension bioreactors .......................194–197, 199, 202

T Tendon-derived stem cells (TDSCs) definition ................................................................. 135 isolation of mice TDSCs................................ 137–139 mechanical simulation extraction of RNA for validation .................... 138, 141–142 scaffold-based ...................................137, 140–141 scaffold-free ......................................137, 139–140 primer sequences for RT-qPCR analysis................ 142 scaffold-based method ............................................ 136 tenogenesis marker.................................................. 136 Tenogenesis, of TDSCs ................................................ 136 Thermoplastic microfluidic devices, fabrication protocol biological and medical applications.......................... 27 chip design and preparation electrochemical wet etching ............................... 32 photolithography .......................................... 29–31 using AutoCAD software ................................... 31 drilling and planarization.......................................... 34 of electrodes blank COP substrate .............................. 32–33, 36 Cr/Au etching .................................................... 33 design creation .................................................... 32 hot embossing ..................................................... 34–36 market and market reports ....................................... 28 materials..................................................................... 29 microfluidic device with electrodes .......................... 36 microfluidic device without electrodes .............. 35–36 modification and durability ...................................... 28 optical properties....................................................... 28 thermo-compression bonding .................................. 35 3D scaffolds, cell activity in bioartificial tissues ................................................... 127 bioreactor system ........................................... 129–130 mechanical compression load and fluid flow parameters.................................................... 131 sample preparation .................................................. 130

sample transfer and chamber assembly .................. 131 3D TableTrix® ............................................................... 114 Three-dimensional (3D) tissue equivalents epithelial polarization.............................................. 246 fluid flow.................................................................. 242 liver model ............................................................... 245 materials cell culture ......................................................... 248 processing and analysis of samples ................... 249 methods drug toxicity testing, HepG2 liver model growth for .........................................................249–251 perfused intestinal model creation ...........251–253 perfused model, processing and analysis of ..........................................................253–255 static intestine models............................................. 247 techniques................................................................ 243 transport .................................................................. 242 Tissue engineering ...................................... 127, 167, 180 Tracheal graft de-epithelialization antibiotics, media, and detergents cocktail of antibiotics ........................................ 171 DMEM + antibiotics.................................171–172 materials.....................................................170–171 1% sodium dodecyl sulfate................................ 172 1% triton X-100................................................. 172 bioreactor extra components......................... 168–169 bioreactor main components......................... 168, 169 bioreactor setup.............................................. 175–176 decontamination ............................................ 179–180 detergent circuitries ....................................... 173–174 devices...................................................................... 170 graft procurement and preparation ........................ 172 H2O ................................................................ 177, 178 motor and pumps installation ....................... 176–177 other materials................................................ 169–170 PBS........................................................................... 179 positioning trachea.................................................. 176 SDS .......................................................................... 177 tools ......................................................................... 170 triton X-100 ................................................... 177, 178 Tracheal in vitro reconstruction .......................... 157–160 bioreactor................................................................. 165 materials generation, decellularized tracheal ECM-based porcine bio-scaffold.............................160–161 human chondrocyte propagation and maintenance................................................. 161 porcine trachea collection................................. 160 repopulation, decellularized tracheal ECM-based porcine bio-scaffold.............................161–162 methods generation, decellularized tracheal ECM-based porcine bio-scaffold.............................162–163

BIOREACTORS

IN

STEM CELL BIOLOGY: METHODS

AND

PROTOCOLS Index 273

human chondrocyte propagation and maintenance................................................. 163 porcine trachea collection................................. 162 repopulation, decellularized tracheal ECM-based porcine bio-scaffold..................................... 164 tracheal segments .................................................... 164 Tracheal transplantation ............................................... 167 Triton X-100 ......................................................... 73, 162, 171–173, 177–179, 249, 254

cell harvesting................................................. 122–123 cell quality assessment-related reagents ................. 116 3D FloTrix® digest solution ................................... 116 3D FloTrix® vivaSPIN bioreactor ................. 116–118 3D TableTrix® microcarrier tablets .............. 115–116 inoculation on ................................................ 118–119 three-dimension (3D) dynamic cell culture .......... 114 tissue engineering and cell-based therapies ........... 113 Uniaxial stretching ...................................... 136, 137, 143

U

W

Umbilical cord mesenchymal stem cells (UCMSCs) cell culture, medium, replenishment, and growth monitoring.......................................... 119–121 cell culture reagents ....................................... 114–115

Wave-assisted agitation ................................................. 152 Working cell bank (WCB) .............................................. 89 WST-1 proliferation assay .................................... 213–215