CARDIOVASCULAR DEVELOPMENT methods and protocols. [2 ed.] 9781071614808, 1071614800

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Methods in Molecular Biology 2319

Xu Peng Warren E. Zimmer Editors

Cardiovascular Development 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 PubMed.

Cardiovascular Development Methods and Protocols Second Edition

Edited by

Xu Peng, and Warren E. Zimmer Med. Physiology, Health Science Center, Texas A&M University, Bryan, TX, USA

Editors Xu Peng Med. Physiology, Health Science Center Texas A&M University Bryan, TX, USA

Warren E. Zimmer Med. Physiology, Health Science Center Texas A&M University Bryan, TX, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1479-2 ISBN 978-1-0716-1480-8 (eBook) https://doi.org/10.1007/978-1-0716-1480-8 © Springer Science+Business Media, LLC, part of Springer Nature 2012, 2021 This work is subject to copyright. All rights are reserved 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. 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 cardiovascular system is the first system formed during embryogenesis, and interfering with the process of cardiovascular development results in congenital cardiovascular disease, which affects nearly 1% of live human births. Cardiovascular development is a complicated process, involving the coordination and alignment of a plethora of cell types. Benefiting from technological advances in genomics and imaging, recent studies have identified many genetic, epigenetic, and signal transduction regulators that are important for cardiovascular development. A pumping heart is constructed from a few progenitor cells, derived from both the primary and secondary heart fields. First, bilaterally located cardiomyocytes migrate toward the embryonic midline and fuse to form a single heart tube. After recruiting cardiomyocytes from the secondary heart field, the heart tube begins to elongate, which involves initiating a rightward looping and then eventually forming separated chambers. Vascular endothelial cells, differentiated from hematopoietic progenitors, also play an essential role in cardiovascular development by dominating the blood vessel formation process. Blood vessels are formed through both vasculogenesis and angiogenesis. First, the primitive blood vessel structure is formed through vasculogenesis, which includes in situ hemangioblast differentiation and proliferation. Once the primitive blood vessels are established, new blood vessels can grow from the pre-exiting vessels to form the final complicated network—a process known as angiogenesis. Following the creation of vasculature, a group of vein vascular endothelial cells express a series of lymphatic markers and migrate out of the vein to form lymphatic vessels. One of the main functions of lymphatic vasculature or the lymphatic system is to transport absorbed tissue fluid back into blood circulation to maintain tissue homeostasis. Cardiovascular development II was designed to bring together a variety of methods routinely used in cardiovascular development studies. One main obstacle in studying embryonic development in mice is that the embryo becomes opaque by the mid to late embryogenesis stages. With newly established tissue clearing techniques, performing threedimensional vascular constructions became a reality. Over the past two decades, lymphatic development has been a flourishing field, witnessing many new developments that have projected scientific inquiry and experimentation at unprecedented speeds. In this book, we added several protocols for lymphatic development, including lymphatic endothelial and muscle cell isolation and culture. We also include lymphatic staining techniques for different organs such as the embryonic skin and mesentery. Bryan, TX, USA

Xu Peng Warren E. Zimmer

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

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1 Tissue Clearing and 3-D Visualization of Vasculature with the PEGASOS Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Dian Jing, Yi Men, and Hu Zhao 2 Preparation and Identification of Cardiac Myofibrils from Whole Heart Samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Heidi A. Creed and Carl W. Tong 3 RNA-Sequencing and Transcriptome Analysis after Fluid Shear Stress Stimulation in Lymphatic Endothelial Cells . . . . . . . . . . . . . . . . . . . . . 25 Hongjiang Si and Xu Peng 4 Preparation of Neonatal Rat Papillary Muscles for Contractile Studies . . . . . . . . . 31 Steven Jokerst, Damir Nizmutdinov, Charley Edgar, April M. Kaspick, Carl W. Tong, and David E. Dostal 5 In Vitro Study of Permeability in Lymphatic Endothelial Cells Responding to Histamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Hongjiang Si 6 Optimizing the Differentiation of Cardiomyocytes from Human Induced Pluripotent-Derived Stem Cells . . . . . . . . . . . . . . . . . . . . . . 51 Melanie Gartz and Jennifer L. Strande 7 Analysis of Angiogenesis in Mouse Embryonic Dorsal Skin by Whole-Mount Fluorescent Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 Jian Wang, Yuwei Dong, Yu Xi, and Xu Peng 8 Isolation and Culture of Mouse Lymphatic Endothelial Cells from Lung Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Philip E. Lapinski and Philip D. King 9 Laser-Induced Choroidal Neovascularization in Rats . . . . . . . . . . . . . . . . . . . . . . . . 77 Min Zhao, Wankun Xie, Travis W. Hein, Lih Kuo, and Robert H. Rosa Jr 10 Detecting Three-Dimensional Vascular Networks in the Mouse Embryonic Hindbrain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Chenshen Huang, Liangjing Wu, Jian Wang, Binu Tharakan, and Xu Peng 11 Visualizing Blood Vessel Development in Cultured Mouse Embryos Using Lightsheet Microscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Samantha J. Fredrickson, Tanner G. Hoog, and Ryan S. Udan 12 Laser Capture Microdissection of Vascular Endothelial Cells from Frozen Heart Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Tianhao Zhou and Jian Wang

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Visualization of Retinal Blood Vessels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wankun Xie, Min Zhao, Travis W. Hein, Lih Kuo, and Robert H. Rosa Jr Transient Transgenics: An Efficient Method to Identify Gene Regulatory Elements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Warren E. Zimmer and Robert J. Schwartz Isolation of Lymphatic Muscle Cells (LMCs) from Rat Mesentery . . . . . . . . . . . . Xueyang Zhang, Sanjukta Chakraborty, Mariappan Muthuchamy, and David C. Zawieja Isolation of Adult Mouse Cardiomyocytes Using Langendorff Perfusion Apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yang Liu, David E. Dostal, and Carl W. Tong Analysis of Lymphatic Vessel Formation by Whole-Mount Immunofluorescence Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jian Wang, Yuwei Dong, Mariappan Muthuchamy, David C. Zawieja, and Xu Peng

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

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Contributors SANJUKTA CHAKRABORTY • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA HEIDI A. CREED • Department of Medical Physiology, College of Medicine, Texas A&M University of Health Science Center, Bryan, TX, USA YUWEI DONG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA DAVID E. DOSTAL • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA; Central Texas Veterans Health Care System, Department of Internal Medicine, Dell Medical School, The University of Texas at Austin, Austin, TX, USA CHARLEY EDGAR • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA SAMANTHA J. FREDRICKSON • Department of Biology, Missouri State University, Springfield, MO, USA MELANIE GARTZ • Department of Cell Biology, Neurobiology and Anatomy‘, Medical College of Wisconsin, Milwaukee, WI, USA; Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, WI, USA; Neuroscience Research Center, Medical College of Wisconsin, Milwaukee, WI, USA TRAVIS W. HEIN • Department of Medical Physiology, College of Medicine, Texas A&M University Health Science Center, Bryan, TX, USA; Department of Ophthalmology, Baylor Scott & White Eye Institute, Temple, TX, USA TANNER G. HOOG • Department of Biology, Missouri State University, Springfield, MO, USA CHENSHEN HUANG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA DIAN JING • State Key Laboratory of Oral Diseases and National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China; Department of Comprehensive Dentistry, Texas A&M University College of Dentistry, Dallas, Texas, USA STEVEN JOKERST • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA APRIL M. KASPICK • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA PHILIP D. KING • Department of Microbiology and Immunology, University of Michigan Medical School, Ann Arbor, MI, USA LIH KUO • Department of Medical Physiology, College of Medicine, Texas A&M University Health Science Center, Bryan, TX, USA; Department of Ophthalmology, Baylor Scott & White Eye Institute, Temple, TX, USA PHILIP E. LAPINSKI • Department of Microbiology and Immunology, University of Michigan Medical School, Ann Arbor, MI, USA YANG LIU • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA YI MEN • State Key Laboratory of Oral Diseases and National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China;

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Department of Comprehensive Dentistry, Texas A&M University College of Dentistry, Dallas, Texas, USA MARIAPPAN MUTHUCHAMY • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA DAMIR NIZMUTDINOV • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA XU PENG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA; Department of Medical Physiology, College of Medicine, Texas A&M University of Health Science Center, Bryan, TX, USA ROBERT H. ROSA • Department of Medical Physiology, College of Medicine, Texas A&M University Health Science Center, Bryan, TX, USA; Ophthalmic Vascular Research Program, Department of Ophthalmology, Baylor Scott & White Eye Institute, Temple, TX, USA ROBERT J. SCHWARTZ • Department of Biology & Biochemistry, University of Houston, Houston, TX, USA HONGJIANG SI • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA JENNIFER L. STRANDE • Department of Cell Biology, Neurobiology and Anatomy‘, Medical College of Wisconsin, Milwaukee, WI, USA; Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, WI, USA; Neuroscience Research Center, Medical College of Wisconsin, Milwaukee, WI, USA BINU THARAKAN • Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA CARL W. TONG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA RYAN S. UDAN • Department of Biology, Missouri State University, Springfield, MO, USA JIAN WANG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA LIANGJING WU • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA YU XI • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA WANKUN XIE • Department of Medical Physiology, College of Medicine, Texas A&M University Health Science Center, Bryan, TX, USA; Ophthalmic Vascular Research Program, Department of Ophthalmology, Baylor Scott & White Eye Institute, Temple, TX, USA DAVID C. ZAWIEJA • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA XUEYANG ZHANG • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA HU ZHAO • Department of Comprehensive Dentistry, Texas A&M University College of Dentistry, Dallas, Texas, USA MIN ZHAO • Department of Medical Physiology, College of Medicine, Texas A&M University Health Science Center, Bryan, TX, USA; Ophthalmic Vascular Research Program, Department of Ophthalmology, Baylor Scott & White Eye Institute, Temple, TX, USA TIANHAO ZHOU • Department of Medical Physiology, College of Medicine, Texas A&M University, Bryan, TX, USA WARREN E. ZIMMER • Department of Medical Physiology, Texas A&M College of Medicine, Bryan, TX, USA

Chapter 1 Tissue Clearing and 3-D Visualization of Vasculature with the PEGASOS Method Dian Jing, Yi Men, and Hu Zhao Abstract Tissue clearing techniques turn tissue transparent through a series of chemical and physical treatments. They have provided a useful tool for three-dimensional (3-D) imaging to study tissue spatial organization and interactions. Many tissue clearing methods have been developed in recent years. Each method has its own application range depending on the purposes of the study. Three criteria for selecting an appropriate clearing method include clearing transparency, fluorescence preservation, and broad tissue applicability. PEG-associated solvent system (PEGASOS) emerged recently as a solvent-based tissue clearing method capable of rendering diverse tissues highly transparent while preserving fluorescence. Combined with vascular labeling techniques, PEGASOS method enables 3-D visualization of vasculature in whole tissues at subcellular resolution. Here, we describe the standard PEGASOS passive immersion protocol and several compatible vascular labeling techniques. Methods of 3-D imaging, data processing, and annotations are also briefly introduced. Key words Tissue clearing, 3-D imaging, Vasculature, PEGASOS

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Introduction Tissue opacity is mainly caused by light scattering and light absorption. The heterogeneous optical properties of different tissue components lead to refractive index (RI) mismatches, which in turn cause light scattering [1]. Moreover, endogenous pigments including heme, melanin, lipofuscin, and cytochrome strongly absorb light [2]. The calcified minerals in hard tissues also block the light transmission. In order to achieve transparency, both chemical and physical approaches are, therefore, applied in various tissue clearing methods to eliminate light scattering components and to achieve uniform internal RI, known as RI matching. Achieving of RI matching enables images being acquired even at depth of several millimeters into tissues.

Xu Peng and Warren E. Zimmer (eds.), Cardiovascular Development: Methods and Protocols, Methods in Molecular Biology, vol. 2319, https://doi.org/10.1007/978-1-0716-1480-8_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 PEGASOS passive immersion procedure for both soft and hard tissues. (a) Treatment steps and timetables of PEGASOS passive immersion procedure for clearing soft tissues. (b) For hard tissues, perform decalcification between fixation and decolorization

Over the last decade, numerous tissue clearing techniques have been developed, and they all followed above principles to turn tissue transparent. These clearing methods can be classified into three major categories: (1) organic solvent-based tissue clearing techniques, including the DISCO series [3–5], fluoclear BABB [6], and polyethylene glycol (PEG)-associated solvent system (PEGASOS) [7]; (2) Aqueous reagent-based tissue clearing techniques, such as SeeDB2 [8], the CUBIC series [9, 10], and Ce3D [11]; (3) hydrogel-based TC techniques, including CLARITY [12], PACT [13], and various modified protocols [14, 15]. Here, we introduce the PEGASOS tissue clearing method developed by our lab, which is a solvent-based clearing method capable of turning both soft and hard tissues highly transparent with excellent fluorescence preservation [7]. The standard PEGASOS workflow includes the following steps: perfusion, decalcification, decolorization, delipidation, dehydration, and clearing. Performing whole-body PEGASOS via recirculation system renders the whole mouse transparent, whereas the PEGASOS passive immersion procedure is suitable for individual organs (Fig. 1). We will describe the passive immersion protocol in the following section. Vascular system forms complicated spatial pattern within organs. 3-D imaging based on tissue clearing provides a powerful research tool. Many methods have been used to label the vasculature. Transgenic mouse models are the most popular and recommended method for vasculature labeling. With proper transgenic mouse strain, vasculature components can be efficiently labeled with GFP or other fluorescent proteins [7, 16–18]. Dye injection and dye perfusion are conventional labeling approaches. Commonly used dyes include GS-IB4, dextran sulfate, and DiI [19– 21]. However, not all the dyes are compatible with every clearing method. PEGASOS method is compatible with GS-IB4 and dextran sulfate but not DiI. Solvent components including tertbutanol and benzyl benzoate (BB) dissolve DiI. In addition, the vasculature labeling efficiency of dye is usually lower than using transgenic model and can vary significantly among different animals. Traditional immunostaining with antibodies is also a valuable

Tissue Clearing and 3-D Visualization of Vasculature with the PEGASOS Method

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Fig. 2 Combination of vascular labeling with the clearing process. Overview of the PEGASOS tissue clearing method and compatible vascular labeling methods. Transgenic mice with endogenous vascular fluorescence are most commonly used with this tissue clearing method (①). Retro-orbital injection and tail vein injection provide effective means of labeling the vasculature (②, ③) before sacrifice. In transcardial-circulatory perfusion, dye can be also perfused via the vascular system (④). For dissected thick tissue slices, wholemount immunostaining is another labeling approach that can be applied before the delipidation process (⑤)

tool for labeling vasculature. Penetration of antibodies into deep regions is the major challenge. Antibody incubation conditions should be optimized to improve staining efficiency [7, 16, 19]. In our experience, whole-mount immunostaining with regular antibodies should be applied for soft tissue slices of no more than 500 μm thickness. Our efforts of whole-mount immunohistochemical staining for hard tissue organs or thick slices were never successful. Combined with these vascular labeling techniques, we demonstrated PEGASOS method is able to visualize vasculature within different organs and tissues in 3-D [7, 22, 23] (Fig. 2). Here, we describe the protocol of the PEGASOS passive immersion method in detail. Four compatible vessel labeling methods, including transgenic mouse models, retro-orbital dye injection, postmortem dye perfusion, and whole-mount immunostaining, are introduced. Technical details for confocal imaging and data processing are briefly introduced.

2 2.1

Materials Animals

1. Transgenic mice of 4–8-week-old with endogenous vascular labeling. Commonly used mouse strains for labeling vasculatures: Cre strains for labeling endothelium include Tie2-Cre (JAX #008863) and Cdh5-CreERT2 [23]. aSMA-CreERT [7] mouse can be used for labeling smooth muscle tissue to distinguish arteries from veins and capillaries. NG2-DsRed (JAX #008241), NG2-CreER™ (JAX #008538), or Leptin-Cre (JAX #008320) mouse strains can be used for labeling pericytes. Commonly used reporter strains include tdTomato (Ai14) (JAX #007908), tdTomato (Ai9) (JAX #007909), EYFP (Ai3) (JAX #007903), and ZsGreen (Ai6) (JAX #007906). To generate Tie2-Cre, Ai14 mice, male Tie2-Cre

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mice were bred with female Ai14 mice. PEGASOS method is compatible with fluorescent proteins used in above reporter mouse strains. 2. Dye labeling or immunohistochemical staining strategy is applicable to mice of any genotype. 3. All animal experiments should be approved by the relevant institutional animal care and performed in accordance with guidelines from the NIH or the corresponding national entity governing such procedures. 2.2 Anesthesia System

1. Isoflurane. 2. Oxygen cylinders. 3. Ketamine (80 mg/kg body weight). 4. Xylazine (10 mg/kg body weight).

2.3 Retro-Orbital Injection

1. Dyes for vascular labeling: (a) Fluorescein isothiocyanate–dextran (2000 kDa, SigmaAldrich FD2000S). Dilute FITC-Dextran in PBS with a concentration of 75 mg/mL. (b) DyLight 594 labeled Griffonia Simplicifolia Lectin I (GSL I) isolectin B4 (GS-IB4) (Vector Lab DL1207). Dilute 0.5 mg GSL I-B4 in PBS or saline water to reach a final concentration of 1 mg/mL. 2. 28-gauge insulin syringe.

2.4 Perfusion and Fixation

1. Heparin PBS: 10 U/mL heparin sodium in PBS. 2. 4% paraformaldehyde (PFA) (pH 7.4). 3. 22-gauge needle. 4. 25 mL syringe. 5. Silicone tube. 6. Dissecting straight scissors. 7. Surgical scissors. 8. Straight tweezers. 9. 50 mL centrifuge tubes. 10. Foam plate and pins. 11. Blood container.

2.5 PEGASOS Passive Immersion Procedure

1. Decalcification solution: 0.5 M ethylenediaminetetraacetic acid (EDTA). Add sodium hydroxide to dissolve EDTA and adjust the pH to 7.0. 2. Decolorization solution: 25% N,N,N0 ,N0 -Tetrakis(2-Hydroxypropyl) ethylenediamine (Quadrol) (see Note 1).

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3. Delipidation solution: Dilute pure tert-butanol (tB) (SigmaAldrich 360,538) with dH2O to prepare 30% tB, 50% tB, and 70% tB (v/v). Add pure Quadrol (3% w/v final concentration) to adjust the pH to above 9.5 (see Note 2). 4. Dehydration solution: tB-PEG, a mixture of 70% v/v tB, 30% v/v PEG methacrylate Mn 500 (PEGMMA500) supplemented with 3% w/v Quadrol. 5. Clearing medium: BB-PEG, a mixture of 75% v/v BB and 25% v/v PEGMMA500 supplemented with Quadrol (3% w/v final concentration). The RI of BB-PEG is 1.543. 6. 50 mL centrifuge tubes. 7. A temperature-controlled shaker. 2.6 Whole-Mount Immunostaining

1. Blocking buffer composed of 10% dimethyl sulfoxide, 0.5% IGEPAL CA-630, and 1 casein buffer in PBS. 2. Primary antibody: Anti-α-smooth muscle antibody (dilution 1:500) is commonly used for artery labeling. Anti-CD31 antibody (dilution 1:100) is commonly used for vessel endothelium labeling. Isolectin B4 (GS-IB4) can also be used for endothelium staining. Optimal dilution should be determined individually, and 1:100 is a good starting dilution to test. 3. Alexa Fluor conjugated secondary antibody. 4. 1.5 mL Eppendorf tubes. 5. Aluminum foil paper. 6. A temperature-controlled shaker.

2.7 3-D Imaging Acquisition for Cleared Samples

1. Depression slide.

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1. ImageJ software (National Institutes of Health).

Data Processing

2. Cover glass. 3. Imaging system: confocal microscopy/two-photon microscopy/light-sheet fluorescent microscopy.

2. Imaris software (Bitplane). 3. High performance computer workstation. The optimum configuration includes: 128 GB RAM, dual CPU, high-end graphic card (Nvidia GTX 1080 Ti or AMD Radeon 580), and large-storage hard drives (>5 TB).

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Methods (For endogenously labeled mice, skip methods in Subheading 3.1 and proceed to methods in Subheading 3.2)

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3.1 Retro-Orbital Injection of Vasculature Dye (See Note 3)

1. Anesthetize a mouse using 3% isoflurane and 4% oxygen. Alternatively, a cocktail of ketamine (80 mg/kg body weight) and xylazine (10 mg/kg body weight) could be intraperitoneally administered (see Notes 4 and 5). 2. Prepare one of the vascular labeling dyes in a 28-gauge insulin syringe: 200 μL (a) 75 mg/mL FITC-Dextran or (b) 1 mg/ mL GS-IB4. 3. Place the mouse in left recline position with the head facing to the right. Apply gentle pressure with fingers to the skin around the right eye to slightly protrude the eye from the eye socket (see Note 6). 4. Carefully introduce the needle via the medial canthus at an angle of 30 with bevel down (see Note 7). The bone beneath can be felt as a guide. Once the needle tip is at the base, slowly and smoothly depress the plunger to deliver the dye (see Notes 8–10). 5. Slowly withdraw the needle after injection. Wait for 5–30 min until dye circulates through the whole body before sacrificing the animal. Optimal circulation time for different organs can vary and should be determined individually (see Note 11).

3.2 Perfusion (Including Postmortem Dye Perfusion) and Tissue Preparation

1. Prepare the surgical setup including 50 mL cold heparin PBS (see Note 12), 25 mL cold 4% PFA, and surgical instruments. Connect a 22-gauge needle to a 25 mL syringe using a silicone tube. Eliminate the bubbles in the injection system. 2. Limbs of the anesthetized mouse can be directly fixed on a foam plate. All the following steps must be performed in a biosafety hood. 3. Open the chest and abdominal cavity with surgical scissors, then insert the 22-gauge needle into the mouse’s left cardiac ventricle with bevel up (see Note 13). Inject about 2 mL heparin PBS to fill the heart and cut an incision on the right atrium immediately using a surgical scissor. The right atrium incision provides the outlet for the perfusion fluid. 4. Inject 50 mL heparin PBS to flush out as much blood as possible (see Note 14). 5. Postmortem dye perfusion (optional): Vasculature labeling dyes can also be perfused through the circulation at this stage. After flushing out the blood, inject dye solution to label the vessels: Dilute 200 μL (a) 75 mg/mL FITC-Dextran or (b) 1 mg/mL GS-IB4 in 3 mL PBS for perfusion (see Note 15). Wait for 5 min for complete staining and proceed to the next step. 6. Perfuse 25 mL 4% PFA transcardially for fixation (see Note 14). 7. Dissect carefully the target samples.

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8. Immerse the dissected samples in 4% PFA at 4  C overnight. On the following day, wash samples with PBS three times for 20 min each on a shaker. 3.3 PEGASOS Passive Immersion Procedure (Fig. 1)

1. Decalcification (only for hard tissues): immerse samples in 0.5 M EDTA (pH 7.0) at room temperature (RT) on a shaker at 60 rpm for around 4 days. Change the decalcification solution daily. Wash samples with pure H2O for 30 min to elute excessive EDTA. 2. Decolorization: place samples in 25% Quadrol for 1–2 days at 37  C under constant shaking at 60 rpm. Change medium daily until the medium does not turn yellow anymore (see Note 16). 3. Delipidation: treat samples with gradient delipidation solutions of 30%, 50%, and 70% tB at 37  C under constant shake at 60 rpm. The treatment time depends on the tissues volume. Suggested durations for different tissues are summarized (Table 1). This step takes approximately 2 days. 4. Dehydration: immerse samples into tB-PEG solution at 37  C on a shaker at 60 rpm for 2 days. Change the medium at least once. After dehydration, the samples should be kept away from any water containing reagent. 5. Clearing: immerse samples in the final BB-PEG clearing medium for 1 day on a shaker at 37  C until tissues turn transparent (see Note 17) (Fig. 3a). Cleared tissue can be preserved in the BB-PEG solution at room temperature (RT).

3.4 Whole-Mount Immunostaining Within PEGASOS Passive Immersion Procedure

The following protocol is based on the whole-mount immunohistochemical performed on brain slice of 500 μm thickness. 1. Decalcification and decolorization are the same as in Subheading 3.3. 2. Wash samples with PBS solution for 30 min before blocking. 3. Blocking: prepare the blocking buffer in a 1.5 mL Eppendorf tube containing blocking solution. Tissue sections are placed into 1 mL blocking buffer and incubated overnight at RT on a shaker (60 rpm) (see Note 18).

Table 1 Time schedule of delipidation for different tissues

Delipidation

Tissue organs at 4–6 weeks age

Tissue organs at 8 weeks age

Tissue sections 90% are positive by PCR and stain for LacZ (reporter gene) activity. This number ensures that adequate, reproducible data are collected. We ask the core to inject enough zygotes to implant 15–25 into each pseudopregnant females for each time point we wish to analyze. The core does the injections by the general guidelines described in Manipulating the Mouse Embryo: a laboratory manual [15] although each have derived site-specific protocols and then notifies our laboratory of the animals for our harvest at designed developmental time points for analysis as described below (Subheading 3.3). 1. To initiate BAC isolation for microinjection, streak from the 80  C bacterial stock containing the modified—reporter gene substituted—BAC clone onto an agar plate containing antibiotic selection (kanamycin and chloramphenicol, 20 mg/mL and 25 mg/mL, respectively) the evening before starting liquid cultures and incubating at 37  C. 2. Pick a single colony and seed into a flask of 800 mL LB media containing antibiotics and shake at ~300 RPM overnight in a temperature-controlled incubator/shaker at 37  C (see Note 4). 3. Centrifuge the cells to a pellet at 2000  g for 15 min. We use a Beckman centrifuge equipped with a JA-10 rotor spinning at ~4000 RPM. Discard the supernatant and drain fully by turning up on a paper towel. 4. Each pellet is resuspended in 100 mL of P1 buffer, centrifuge as above to pellet the bacteria, and once the supernatant is removed, resuspend the pellet in 60 mL of P1 buffer. Add 100,000 units of Ready-lyse (catalog R1082M, Epicenter Technologies, Madison, WI) and allow the mixture to incubate at room temperature for 15–20 min (see Note 5). 5. Carefully add 30 mL of P2 buffer to tube and allow to incubate at room temperature for an additional 5 min. Do not mix strongly or vortex, which will damage the BAC DNA molecule. This step can be enhanced by rolling the tubes on the bench top. 6. Add 30 mL of chilled P3 buffer and incubate on ice for 15 min. Do not mix vigorously or vortex. The P2 and P3 buffers can be added by washing down the sides of the tube when adding; this gently adds the buffers and leads to better distribution of the buffer.

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7. Centrifuge the bacterial slurry at 15,000  g (9000 RPM in the JA-10 rotor) for 30 min at 4  C. Carefully remove the supernatant into a cleaned chilled tube and recentrifuge as above to completely clear any sediments, which can clog the column (next step) and significantly reduce the DNA yield. Filter the lysate through a filter and directly onto a Maxiprep column. 8. The column is washed with kit-provided buffers and then the DNA eluted by addition of 15 mL of supplied buffer. 9. DNA is precipitated by addition of 11 mL room temperature isopropanol, mixing carefully (no vortex) and incubated for 5 min on the bench top. The DNA is collected by centrifugation at 15,000  g for 30 min at 4  C. 10. Carefully pour off the supernatant and air-dry the pellet for 5 min at room temperature. Resuspend the pellet in 1 mL of P1 buffer and add RNase A (Sigma) to 100 units/mL and incubate at 37  C for 30 min. Add 100 μL of 10% SDS and proteinase K to 0.2 mg/mL and incubate at 50  C for 2 h. This step can be extended to an overnight incubation if necessary. 11. Add an equal volume of buffered phenol (pH 8.0) by gentle inversion (no vortex) and centrifugation. 12. Transfer the top aqueous layer to a new tube for a second phenol extraction and finally transfer the supernatant to a fresh tube to add an equal volume of chloroform. 13. The chloroform extraction is repeated and the aqueous layer (top layer) transferred to a fresh tube and the DNA precipitated by adding 7.5 M to final concentration of 2.5 M and 0.5 volume of isopropanol and incubation at room temperature for 5 min. 14. Centrifuge at 15,000  g for 30 min and carefully remove the supernatant and air-dry the pelleted DNA for 30 min at room temperature. 15. Resuspend the DNA pellet into ~0.5 mL P1 buffer. This needs to be a gentle resuspension, which may require an incubation overnight at room temperature to allow the BAC DNA to gently be absorbed into the buffer solution. 16. DNA concentration can then be quantitated via absorption measures at 260 nM. The DNA can be stored at 4  C. 17. The DNA can be used at this point or it may be useful to produce a linearized BAC before injection. To linearize the DNA, place 10 μg DNA into a tube and add 100 μ/mL of lambda terminase and incubate at 37  C for 30 min (see Note 6).

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18. If the DNA is linearized, the DNA should be purified with buffered phenol/chloroform extraction and precipitation with ammonium acetate as described above. Resuspend the DNA into P1 buffer and store at 4  C until needed for injection. 19. Prepare the DNA for injection by diluting to a concentration of 1 ng/μL with microinjection buffer and send to the core for injection on ice. 3.3 Collecting and Analysis of Reporter Gene Expression

The crux of the transient transgenic experiment is the collection of timed embryos for analysis. Once we receive notice from the core of successful injections and transfer of zygotes to the pseudopregnant females, we take the mice into separate cages (if different DNAs are injected) and assign them for developmental timing for analysis. We count the day after implantation as day 1 and then set developmental times for collection based upon this determination. Depending upon the reporter gene used, there are some differences in the actual data collection (e.g., using EGFP—fluorescence analysis or LacZ staining for b-galactosidase activity), and here, we will describe our work with LacZ reporter analysis. We illustrate results using both EGFP and LacZ reporter gene placed in the Nkx 2.5 locus in Fig. 3. 1. Pseudopregnant recipients of the fertilized embryos injected with the reporter gene/BAC clones are placed into cages, and the embryos allowed to “incubate” until collected at the desired developmental time points. For our experiments, we collected embryos at times between 7.5 and 13.5 embryonic days. 2. At the designated day, the carrier mouse surrogates are euthanized using approved techniques (by CO2 inhalation per protocols approved by institutional committees) and prepared for embryo collection by spraying 70% ethanol on the ventral abdominal surface to limit hair sticking to surgical instruments. 3. The abdominal cavity is opened ventrally using sharp surgical scissors and the uterus located by moving through the fascia tissue using a pair of dissecting forceps. Mice contain a pair of uterine horns, which will appear as “beads on a string” when pregnant. 4. Once the uterine horns are located, they are dissected and placed into a petri dish containing DEPC-treated PBS. The mesometrium of the uteri is opened with scissors, which reveals the individual embryos. 5. Carefully remove the embryos and covering membranes (yolk sac) into a separate petri disk containing DEPC-treated PBS. 6. The visceral yolk sac surrounding each embryo is pierced with forceps and then separated from the embryo with gentle pressure.

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Fig. 3 Examples of “transient transgenics” and BAC recombineering in the identification of Nkx 2.5 tissuespecific transcriptional enhancers. (a) shows the results of using LacZ reporter constructs to locate sequences that regulate Nkx 2.5 transcription in specific-tissue fields. A number of LacZ reporter gene DNAs were constructed to contain various lengths of sequences 50 to the transcriptional start of the 2.5 gene are diagrammed. These were injected into 0.5 day embryos, which were allowed to develop for 12.5 days, after which the embryos were collected and analyzed. As described in this text, yolk sacs from embryos were examined for the presence of the LacZ gene sequence by PCR, and almost all that showed PCR-positive developed LacZ staining. Using this analysis, an enhancer important for transcriptional activity in the A-V canal of the heart is located between 10,765 bp and 5011 bp in front of the Nkx 2.5 gene (compare panel d and i) and an enhancer directing transcription in the right ventricle is located between 10,765 and 8554 bp in front of the gene. This right ventricle enhancer needs the context of the immediate 50 Nkx 2.5 promoter because deleting this segment results in reduced right ventricle expression (compare panel d with panel m). Thus, the transient transgenic technique successfully identified sequences located far from the gene that are critical for the appropriate transcription of the gene and that some enhancers require the correct context relative to the gene for their activity. (Figure from Reecy et al., Ref. 1, The Company of Biologist Limited with permission)

7. The yolk sac is placed into a 1.5 mL microcentrifuge tube containing DNA isolation buffer and the released embryo placed into a separate numbered 1.5 mL tube containing PBS and kept on ice. 8. This is continued until each yolk sac and corresponding embryo are dissected and placed into correspondingly numbered tubes after which the yolk sacs are placed into a 55  C water bath for an overnight incubation and the embryos processed as below (go to step 22). 9. Each yolk sac is incubated in 30–40 μL of DNA isolation buffer at 55  C overnight (~16 h). DNA is then isolated from the disrupted tissue by the addition of 40 μL buffered phenol (pH 8.0) and the tube vortexed.

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10. Phases are separated by centrifugation in a microfuge at max speed for 2 min. 11. The nucleic acids (with genomic DNA) will partition into the top, aqueous layer, which is removed into a fresh tube. 12. The phenol extraction is repeated, and the top layer is again removed to a fresh tube. 13. The aqueous layer is extracted again with 40 μL of buffered phenol and an equal volume (~40 μL) of chloroform:isoamyl alcohol (24:1 v/v). 14. Following this organic extraction, the aqueous layer is placed into a fresh tube and the DNA collected by precipitation with the addition of 4 μL 7.5 M ammonium acetate and 40 μL of 100% ethanol. 15. Mix thoroughly and incubate at room temperature for 10–30 min. The DNA is collected by spinning at maximal speed in a microcentrifuge for 5 min and removing the ethanol containing supernatant. 16. The pelleted DNA is washed by adding 700 μL of 70% ethanol to the tube, mixing thoroughly and centrifugation as described above. This is an important step to remove traces of SDS remaining with the DNA that can inhibit subsequent PCR analysis. 17. The DNA pellet is air-dried and then suspended in TE buffer and stored at 4  C until needed. 18. The presence of the reporter/BAC is determined by PCR on the yolk sac DNA. PCR is done with a combination of three primers—one located 2–500 base pairs upstream of the original gene, one located within the 50 coding segments of the original gene, and one located in the 50 sequences of the LacZ gene. PCR reaction conditions we have employed are: 1 μg yolk sac DNA into a PCR tube with 10 μL of the complete PCR mix from Denville Scientific and add 1 pM, each of the three oligonucleotide primers (see Note 3). 19. Reactions are preformed using an MFJ PCR machine with an initial DNA denaturation at 94  C for 10 min, followed by 30 cycles of 94  C for 1 min, 60  C for 1 min, 72  C for 1 min, and a final extension at 72  C for 5 min. 20. PCR reactions products are analyzed on a 1% agarose gel run in Tris-acetate (TAE) buffer. 21. Embryos from step 4 above were stripped of external membranes and placed into numbered 1.5 mL tubes containing PBS. They are numbered in order to correlate staining data with the PCR confirmation of LacZ gene presence.

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22. For fixing and staining of the whole mount embryos, transfer them into a 24 well plates that have been numbered for identification and contain ~0.5 mL PBS. We use the 24-well plates for the ease of processing multiple embryos, gently shaking on lab rotator, and aspiration of solutions. 23. Fix the embryos by aspirating the PBS and replacing with fix solution and place the plate on a flat lab rotator on low speed for a gentle mixing of the embryos in solution. Young embryos require shorter fixation times, 7.5 day to ~12.5 day need 15–30 min with 12.5 and older requiring up to an hour fixation time (see Note 7). 24. Prior to staining, the fixed embryos are washed with 3–4 changes of detergent rinse buffer, 15–30 min per rinse. 25. Remove the last rinse and replace with enough staining solution to each well that just covers the embryos. 26. Incubate with gentle shaking in the dark for ~2 h (1–3 h) for embryos up to 12.5 days or ~4 h (3–5 h) for embryos over 12.5 days. For longer incubations (3 h or longer), it is important to add Tris–HCl (pH 7.3) to the staining solution. 27. Rinse the embryos with PBS, then they can be stored in 70% ethanol at 4  C until imaged in whole mount and or sectioning with subsequent imaging. 28. For whole embryo imaging, place the embryos into 1.5 mL (small embryos) or 15 mL screw top tubes (larger embryos) and rinse with PBS. 29. Remove the PBS and replace with PBS containing 4% paraformaldehyde overnight to post fix the embryos. 30. Rinse the embryos several times [3–5], 15 min each with PBS, then transfer to a small petri dish and image in the stereo microscope. 31. Gentle movement of the embryos to image from different views can be accomplished using watchman #5 or smaller forceps. 32. To examine reporter gene expression on the cellular level, the stained embryos need to be embedded in wax and sectioned for microscopy. The first step in this process is to dehydrate the tissue through a series of graded ethanol. They have been stored in 70% ethanol (step 27 above) and are dehydrated by sequential incubation in 80, 90, and 100% ethanol for 45 min each (see Note 8). 33. The samples are then ready to embed in wax by incubation in fresh xylene for 30 min and then in a 50:50 mixture of xylene: paraffin wax and incubate at 60  C for 1 h.

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34. Transfer samples to neat (100%) wax at 60  C for two changes and then transfer to histology molds to allow the wax to harden (see Note 9). The samples can then be stored at 4  C until ready for cutting sections for analysis. 35. The wax-embedded sample is trimmed to allow for mounting on the microtome and cutting appropriate sections. We generally cut embryos for an experiment in a sagittal and others in cross sections continuously through the entire embryo. Thus, the sample needs to be trimmed with a sharp razor blade and the affixed to the microtome that will allow the sections desired. 36. Set the microtome at 10 mM sections for analysis. 37. The sectioning is automated and as the sections are cut, they are floated in flotation bath of water and then collected upon microscope slides (see Note 10). Specimens may be counter stained for nuclear content to identify specific cells (see Note 11). 38. The sectioned specimens are examined by microscopy and documented using a digital camera attached to the microscope.

4

Notes 1. As with most PCR protocols to generate fragments for cloning, we have found it necessary to conduct experiments to optimize the yield of the desired product. This is a particular special case as the experiment is designed to add ~150 bp (75 on both the 30 and 50 ends) to the reporter gene segment that conforms to the target gene in order to place the reporter in frame with the genomic gene sequence on the BAC. We routinely do two steps to enhance our obtaining the PCR product; first, we preform 3–5 cycles of PCR with a low-annealing temperature (50–52  C) to facilitate having a template containing the overhanging nucleotides, and we do an initial PCR with this template using a gradient protocol with 12, using 1  C, degree steps or increments from 55  C to 67  C annealing temperatures. With the MFJ PCR machines, we utilize the gradient function to perform this experiment. 2. The overnight incubation of the bacteria was found to enhance the recombination events in the bacteria, resulting in more colonies carrying the reporter gene inserted into the BAC DNA. We have found that leaving the plates to incubate for up to 72 h can allow the slow growing colonies time to appear on the selective media plates. We also have found that the reduction in the level of kanamycin in the plates to 15 μg/mL helps with colony yields.

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3. Using a nested primer approach, the PCR product for the genome copy of the gene will be visualized at a specified size, and any band corresponding to LacZ should run to a different size on agarose gels. We try to make them >250 base pair in difference to make analysis clear, and any band in the LacZ region confirms the presence of this gene. 4. Start all BAC clone cultures from a single colony on a freshly streaked plate containing the appropriate selective media. We have also started a 10 mL culture from a single colony to incubate overnight to seed the larger cultures the next day. 5. It is critical to lyse BAC-containing bacteria gently to keep the large DNA of the BAC clone intact. Additionally, all steps in the isolation protocol should be performed in a manner that is thorough but gentle to maintain BAC integrity. 6. The DNA can be made linear by brief incubation with a restriction enzyme that digests DNA infrequently—such as Not I. These digests should be done for very brief times and monitored to ensure that the DNA is only linearized and not completely digested. 7. Larger embryos (12 day or older) should be split into two sagittal segments to allow the penetration of stain into internal tissues. To image smaller embryos (up to ~11.5 day), we use bright-field illumination with dark-field illumination used on larger embryos. 8. All posttreatments of embryos stained for LacZ (b-galactosidase) should be done to minimize exposure to organics (such as xylenes, ethanol), which can leach the color from the stained embryo. 9. To position the embryos in the wax molds for ease of sectioning (cross or sagittal sections), we use heated forceps before the wax hardens. If the wax does begin to harden before the positioning, place the mold in a bath of melted wax (60  C) to melt the wax around the specimen, use heated forceps to position and then allow to cool. 10. We routinely use slides that are coated with poly-L-lysine (Sigma P1274) for section adherence to the slides. The use of coated slides is necessary if the experiment needs to have further staining, for example, staining nuclei with nuclear fast red. 11. To counterstain the LacZ sections with nuclear fast red in order to better localize cells within tissues, we have rinsed the slides containing sections in xylene (two changes for 2 min each) then rehydrate by 1 min incubation in 100%, 90%, 80%, 70%, 50%, 30%, and then DEPC-treated water. The hydrated sections are then incubated in nuclear fast red solution (0.1%

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nuclear fast red Kernechtrot (Fluka, 60700) with 5% aluminum sulfate hydrate (Mallinckrodt, 3208) for 2–3 min, followed by 2–3 min in DEPC-treated water. The slides are then dehydrated by incubation for 1 min in 30–100% ethanol as above, followed by a brief rinse in xylenes. Sections are covered with a drop of Permount and coverslip added to top the specimens. Mounted slides are stored in slide boxes at room temperature until analyzed. References 1. Reecy JM, Li X, Yamada M, DeMayo FJ, Newman CS, Harvey RP et al (1999) Identification of upstream regulatory regions in the heartexpressed homeobox gene Nkx2-5. Development 126(4):839–849 2. Verzi MP, Stanfel MN, Moses KA, Kim BM, Zhang Y, Schwartz RJ et al (2009) Role of the homeodomain transcription factor Bapx1 in mouse distal stomach development. Gastroenterology 136(5):1701–1710 3. Chi X, Chatterjee PK, Wilson W 3rd, Zhang SX, Demayo FJ, Schwartz RJ (2005) Complex cardiac Nkx2–5 gene expression activated by noggin-sensitive enhancers followed by chamber-specific modules. Proc Natl Acad Sci U S A 102(38):13490 4. Schwartz RJ, Olson EN (1999) Building the heart piece by piece: modularity of cis-elements regulating Nkx2-5 transcription. Development 126(19):4187–4192 5. Stanfel MN, Moses KA, Schwartz RJ, Zimmer WE (2005) Regulation of organ development by the NKX-homeodomain factors: an NKX code. Cell Mol Biol (Suppl 51): OL785–OL799 6. Carson JA, Fillmore RA, Schwartz RJ, Zimmer WE (2000) The smooth muscle gamma-actin gene promoter is a molecular target for the mouse bagpipe homologue, mNkx3-1, and serum response factor. J Biol Chem 275 (50):39061–39072 7. Lee SH, Kim S, Hur JK (2018) CRISPR and target-specific DNA endonucleases for efficient DNA Knock-in in eukaryotic genomes. Mol Cells 41(11):943–952

8. Ahmadzadeh V, Farajnia S, Baghban R, Rahbarnia L, Zarredar H (2019) CRISPR-Cas system: toward a more efficient technology for genome editing and beyond. J Cell Biochem 120(10):16379–16392 9. Dubchak I, Brudno M, Loots GG, Pachter L, Mayor C, Rubin EM et al (2000) Active conservation of noncoding sequences revealed by three-way species comparisons. Genome Res 10(9):1304–1306 10. Mayor C, Brudno M, Schwartz JR, Poliakov A, Rubin EM, Frazer KA et al (2000) VISTA : visualizing global DNA sequence alignments of arbitrary length. Bioinformatics 16 (11):1046–1047 11. Fire A, Harrison SW, Dixon D (1990) A modular set of lacZ fusion vectors for studying gene expression in Caenorhabditis elegans. Gene 93 (2):189–198 12. Warming S, Costantino N, Court DL, Jenkins NA, Copeland NG (2005) Simple and highly efficient BAC recombineering using galK selection. Nucleic Acids Res 33(4):e36 13. Gordon JW (1993) Micromanipulation of gametes and embryos. Methods Enzymol 225:207–238 14. Gordon JW (1993) Production of transgenic mice. Methods Enzymol 225:747–771 15. Nagy A, Gertsenstein M, Vintersten K, Behringer R (2003) Manipulating the Mouse Embryo: A Laboratory Manual. Cold Spring Harbor Press, Cold Spring Harbor, New York, p 751

Chapter 15 Isolation of Lymphatic Muscle Cells (LMCs) from Rat Mesentery Xueyang Zhang, Sanjukta Chakraborty, Mariappan Muthuchamy, and David C. Zawieja Abstract Lymphatic muscle cells (LMCs), with unique characteristics resembling a combination of both cardiac and smooth muscle cells, play an essential role in the spontaneous contraction of the lymphatic vessels to pump fluid forward. However, our understanding of the more detailed molecular phenotypes of LMCs is limited. Here, we described a method to isolate the LMCs from rat mesentery and then culture the cells in vitro, which will serve a lot more molecular biology study of LMCs and significantly improve our knowledge about the unique characteristics of LMCs. Key words Rat mesentery, Lymphatic muscle cells (LMCs), Isolation

1

Introduction An essential function of the lymphatic system is to transport lymph containing immune cells, macromolecules, and lipids, etc. throughout the body in which the spontaneous contraction of the lymphatic muscle cells (LMCs) play an important role [1]. LMCs have traditionally been regarded as a type of smooth muscle cells. Nevertheless, lymphatics not only perform tonic contractions to regulate resistance, but also create phasic contractions to generate flow [2–4]. Correspondently, the molecular basis of the contractile proteins in LMCs also exhibits a unique combination of cardiac and smooth muscle elements [5]. However, the study of more detailed characteristics of LMCs including the origin of LMCs, the mechanism of LMCs differentiation, recruitment and migration during development, and the molecular mechanism regulating lymphatic muscle contraction remains very limited. One of the research barriers is the difficulty in dissecting and culturing LMCs in vitro. The possibility of culturing LMCs in vitro enables us to perform a lot more molecular and cell experiments to study the phenotype of

Xu Peng and Warren E. Zimmer (eds.), Cardiovascular Development: Methods and Protocols, Methods in Molecular Biology, vol. 2319, https://doi.org/10.1007/978-1-0716-1480-8_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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LMCs and will also further facilitate the physiological and pathological study of lymphatics. Here, we described an effective method to dissect the primary LMCs and to culture the cells to P7 for experimental use [6, 7]. Briefly, mesenteric lymphatics are dissected out from a young rat. There are two major types of cells within the lymphatic vessel wall: lymphatic endothelial cells (LECs) within the inside surface and LMCs within the outside surface of the vessel wall. After dissecting the vessels out, the vessel will be cultured for about a week, allowing the LMCs within the outside surface of the wall to migrate out from the vessel. The vessels will be discarded, and the cells that have migrated off of the vessel will be cultured using DMEM cell culture media in a 37
C incubator with 90% air and 10% CO2. There are some possibilities that a limited portion of LECs will also migrate out. Nevertheless, the survival of LECs in vitro requires special growth factors to be included in specific culture media for endothelial cells [6]. LECs will not be able to survive in the LMCs culture medium (DMEM). Immunostaining of smooth muscle markers SM22 and a-SMA will be performed to characterize the muscle cell phenotype of the cultured cells to check the purity of the cells we acquired. Other LMCs markers can also be used [5].

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Material

2.1 Mesenteric Lymphatic Vessel Dissection

1. 70% ethanol. 2. Surgical instruments including scissors, forceps, fine-tipped forceps, and clamps. 3. Surgical suture. 4. Horizontal laminar flow hood. 5. Dissection microscope. 6. 6-well cell culture plate. 7. Digital heating water bath. 8. Dissection petri dish. 9. Dissection microscope.

2.2 Culture and Split Primary LMC

1. 1% gelatin: add 1 g gelatin into 100 ml PBS and sterilize by autoclaving. 2. DMEM with 10% fetal bovine serum (FBS): add 55 ml FBS into 500 ml DMEM. 3. PBS with 1 antibiotics (penicillin & streptomycin). 4. 0.25% trypsin-EDTA. 5. Cell culture plates (6-well plate). 6. 37
C CO2 incubator. 7. Inverted microscope.

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1. Anti-rat SM22 and anti-rat-SMA primary antibodies and corresponding secondary antibodies for immunefluorescence staining. 2. Synth-a-freeze. 3. Cryovials and an isopropanol chamber. 4. Liquid nitrogen.

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Methods

3.1 Dissect Rat Mesentery

1. Sterilize the surgical instruments by autoclave. 2. Coat the cell culture plates with 1% gelatin to help the newly isolated cells to attach. 3. Warm the culture media up to 37
C by water bath heating. 4. Following CO2 euthanasia, open the chest and abdomen of the animal with surgical scissors. 5. Find the mesentery and tie the two ends of mesentery using surgical suture. Take the mesentery out and rinse it into PBS with antibiotics. Re-stretch the mesentery in a PBS-filled dissection petri dish. 6. Dissect several mesenteric lymphatic vessels in hood. Carefully remove the attached fat tissue around the lymphatics using sterilized fine-tipped forceps under a dissection microscope.

3.2 Culture and Split Primary LMCs

1. Create a scratch on the bottom of the 6-well plate (one well for each vessel) using sterilized forceps. Fill each well with 2 ml prewarmed culture media in hood. 2. Set each vessel down to the bottom of each well at the scratched area. Culture the vessel in a 37
C incubator with 90% air and 10% CO2. 3. Check the cell status every day and keep the vessel in the culture plate until when most of the LMCs migrate out from the vessel (normally after 1–1 1/2 week). Then use a sterilized forceps to get rid of the vessel from the well. Change the culture plate with fresh media and continue culturing the cells in the incubator. 4. Check the cell status to make sure there is no contamination or severe cell death every day. Replace with fresh culture media every 2 days. 5. Split the cells (P0 ! P1) when they reach the peak of growth. It is usually when cells get very close to each other, cover almost 70% area of the well, and generate a “hill and valley” phenotype after 2–3 weeks after LMCs dissection. To split the cells, remove the culture media and wash the cells with prewarmed

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PBS (37
C). Then trypsinize the cells with 200 μl 0.25% trypsin-EDTA for 1 min in the 37
C incubator. Then add 2 ml prewarmed culture media and split and transfer the cells into new wells. 6. Continue culturing the cells. Check cell status every day and replace with fresh media every 2 days. Split the cells again when they reach full coverage. 3.3 LMCs Characterization and Storage

1. To characterize the acquired LMCs, perform immunefluorescence staining targeting LMCs markers SM22 and a-SMA. Since there are only two major types of cells within the vessels wall, the point is to confirm the muscle cell phenotype distinguishable from lymphatic endothelium cells. 2. To freeze and store the cells, trypsinize the cells with 0.25% trypsin-EDTA and add enough fresh culture media. Then centrifuge the cell suspension, get rid of the supernatant and resuspend the cells with 1 ml synth-a-freeze, and transfer the cell suspension into the cryovials. Place the cryovials containing the cells in an isopropanol chamber and store them at 80
C overnight. For long-term storage, place the cryovials containing the cells into liquid nitrogen later.

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Notes 1. During the mesentery dissection, the purpose of tying the two ends of mesentery is to avoid the releasing of contents of the gut. 2. When dissecting the vessels, there is a balance of getting rid of most of the attached fat and keeping the vessel intact. For this experiment purpose, it is more important to get the most rid of the attached fat. 3. Make sure to use DMEM without HEPES since HEPES might affect the growth of LMCs [6, 7]. 4. Do not treat the cells with trypsin for more than 5 min, otherwise the cells will get damaged, and the morphology might change dramatically. 5. When discard the vessel from the cell culture plate, you may find the vessel gets a little bit degraded, and it appears not to be integrated. Besides, a significant amount of cells migrating out from the vessel indicates a good preparation. If the population of migrate-out cells is too small, it is likely that they won’t be able to grow further enough to create a culturable population. You may try to dissect a longer vessel without damaging the vessel too much.

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6. When LMCs grow a little too dense, they tend to “pile up” on each other and generate a “hill and valley” phenotype rather than perform cell death. It is a bad sign if you notice a significant scale of dead cell while not showing obvious “hill and valley” phenotype. 7. Dissected primary LMCs grow very slowly. After P4, they might duplicate approximately every 3–5 days. References 1. Zawieja DC (2009) Contractile physiology of lymphatics. Lymphat Res Biol 7:87–96 2. Granger HJ (1979) Role of the interstitial matrix and lymphatic pump in regulation of transcapillary fluid balance. Microvasc Res 18:209–216 3. McHale NG, Meharg MK (1992) Co-ordination of pumping in isolated bovine lymphatic vessels. J Physiol 450:503–512 4. Bohlen HG, Wang W, Gashev A, Gasheva O, Zawieja D (2009) Phasic contractions of rat mesenteric lymphatics increase basal and phasic nitric oxide generation in vivo. Am J Physiol Heart Circ Physiol 297:H1319–H1328

5. Muthuchamy M, Gashev A, Boswell N, Dawson N, Zawieja D (2003) Molecular and functional analyses of the contractile apparatus in lymphatic muscle. FASEB J 17:920–922 6. Zawieja SD, Wang W, Chakraborty S, Zawieja DC, Muthuchamy M (2016) Macrophage alterations within the mesenteric lymphatic tissue are associated with impairment of lymphatic pump in metabolic syndrome. Microcirculation 23:558–570 7. Muthuchamy M, Gashev A, Boswell N, Dawson N, Zawieja D (2003) Molecular and functional analyses of the contractile apparatus in lymphatic muscle. FASEB J 17:920–2

Chapter 16 Isolation of Adult Mouse Cardiomyocytes Using Langendorff Perfusion Apparatus Yang Liu, David E. Dostal, and Carl W. Tong Abstract Heart disease is one of the leading causes of death in the United States. Isolation and culture adult cardiomyocytes are important for studying cardiomyocyte contractility, heart hypertrophy, and cardiac failure. In contrast to neonatal cardiomyocyte isolation, adult mice cardiomyocytes isolation is challenging due to firm connections among cardiomyocytes through intercalated discs. The availability of newly generated genetically modified mouse lines requires to establish protocols to isolation and culture adult mouse cardiomyocyte for in vitro studies. In this manuscript, we described a straightforward method of isolating adult mouse cardiomyocytes using Langendorff perfusion apparatus. Briefly, the hearts were harvested from adult mice and the heart was mounted to Lagendorff apparatus. After perfusion with calcium depletion and collagenase digestion, the left ventricles were minced and filtered. Lastly, the separated cardiomyocytes were treated with CaCl2. The isolated cardiac myocytes can be utilized in a broad range of experiments including screening for drugs. Key words Adult cardiomyocytes isolation, Trypsin, Langendorff

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Introduction The advances in genetically modified mice make it possible to investigate gene functions in cardiac development and diseases. However, the genetically modified mice models are not suitable for large-scale screening and studies on complex signal transduction pathways. The cultured adult cardiac myocytes can be utilized in a broad range of experiments, including cell electrical conduction, cell contractile activity, cell mechanics, and intracellular Ca2+ homeostasis, etc. With the recent advances in gene transfer technology, direct analysis of gene functions on isolated cardiomyocytes is possible. Therefore, establishing a straightforward protocol for isolation adult mouse cardiomyocyte will not only benefit in understanding cardiac physiology, but also employable to test the effect of gene therapy for clinical treatment of cardiovascular disorders.

Xu Peng and Warren E. Zimmer (eds.), Cardiovascular Development: Methods and Protocols, Methods in Molecular Biology, vol. 2319, https://doi.org/10.1007/978-1-0716-1480-8_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Although techniques for isolating cardiomyocytes from other animals, such as rats and rabbits, have been established, successful isolation of adult cardiac myocytes in mice remains challenging. One possible reason is the complicated arrangement of the cardiac tissue. The cardiomyocytes exist as the fundamental contractile units of the heart. Within the intact cardiac tissue, cardiomyocytes are connected by intercalated discs through different junctional complexes [1]. These are closely associated with neighboring cells and the extracellular matrix [1]. Cardiomyocytes are also equipped with bundles of myofibrils (termed sarcomeres), deep invaginations of T-tubules, and a densely packed mitochondrial network [2, 3]. All these structures allow the heart to work as a single functional organ to respond to (hormonal and mechanical) signal regulations and environmental changes. Cardiomyocyte isolation requires disrupting these structures. In addition, cardiomyocytes are highly sensitive to mechanical stretching, enzymatic digestion, oxygen supply, ionic fluctuations, and metabolism changes [4]. All these characteristics make isolating functionally and morphologically intact cardiomyocytes very challenging because it is difficult to disrupt these structures without damaging the cardiomyocytes. Moreover, cardiomyocytes are terminally differentiated cells and have limited proliferative ability. They stop cell division after birth and thus do not multiply in culture. This means that cell numbers gradually diminish in culture. Therefore, establishing a protocol for isolating high-quality and quantity cardiomyocytes is critical for studying adult cardiac function. Langendorff perfusion system has been the centerpiece for cardiomyocyte isolation for over 45 years [5]. However, the protocols used by different laboratories vary depending on the species. The goal of this chapter is to describe a straightforward method for isolating adult mouse cardiac myocytes. This protocol is a modification of a procedure used from prior work in the mouse [6].

2

Materials

2.1 Heart Perfusion System

The Langendorff perfusion system (Radnoti). (Fig. 1a) includes heating coil (Fig. 1b), mouse heart chamber (Fig. 1c), MINIPUMP variable flow (Fig. 1d), and heated circulating bath (Fig. 1e).

2.2 Heart Cannulation and Cardiomyocytes Isolation System

1. Surgical Tools (Fig. 2): One big forceps, one small forceps, one big scissor, one small scissor, two fine-tipped forceps, and one fine-tipped surgical scissor. 2. 1 ml syringe (Fig. 3b). 3. Blunt-ended 20 G needle (Fig. 3a) (see Note 1).

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Fig. 1 Langendorff perfusion system big forceps

big scissor

small forceps

small scissor

fine-tipped forceps

fine-tipped surgical scissor

Fig. 2 Surgical tools for isolating adult heart and loading heart into Langendorff apparatus

4. 6–0 black-braided silk nonabsorbable, nonsterile surgical suture spool (SP102) (Fig. 3c). 5. Syringe holder (Fig. 3d) (see Note 2). 6. 60
15 mm petri dish. 7. 50 ml conical Falcon tubes with Sefar Nylon Mesh filter top (Fig. 4). 8. Sefar Nylon Mesh Lab Pak, 250 Microns Square Opening, 1200
1200 .

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Fig. 3 Heart cannulation system

Fig. 4 50 ml Falcon tube with Sefar Nylon Mesh filter top for accumulating filtered cardiomyocytes

9. Bottle top filtration unit. 10. 30 ml Luer lock syringe. 11. Syringe top filter, 0.2 μm.

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12. Glassware: 10 ml beaker. 13. Hot plate stirrer. 2.3

Buffers

1. Basal Solution: 135 mM NaCl, 4 mM KCl, 1 mM MgCl2, 10 mM HEPES, 0.33 mM NaH2PO4 (see Note 3). 2. Perfusion Buffer: 1
basal solution with 10 mM D-(+)-Glucose, 10 mM BDM, and 5 mM taurine. Adjust the pH to 7.4 at 37  C and filter with bottle top filtration unit (see Note 4). 3. Digestion Buffer: Add collagenase type 2 to perfusion buffer (pH has been adjusted to 7.4 at 37  C) to a final concentration of 525 U/ml. Filter digestion buffer with a 30 ml Luer lock syringe and 0.2 μm syringe top filter (see Note 5). 4. Transfer Buffer A: 1
basal solution with 5.5 mM D-(+)-Glucose, 10 mM BDM, and 5 mg/ml BSA (see Note 6). Adjust pH to 7.4 at 37  C. Filter with a 30 ml Luer lock syringe and 0.2 μm syringe top filter. 5. Transfer Buffer B: 137 mM NaCl, 5.4 mM KCl, 0.5 mM MgCl2, 10 mM HEPES, 5.5 mM D-(+)-Glucose, and 1.8 mM CaCl2. Adjust pH to 7.4 at 37  C. Filter with a 30 ml Luer lock syringe and 0.2 μm syringe top filter (see Note 7).

2.3.1 Transfer Solution (See Note 8)

1. Transfer Solution 1: Add 0.2 ml transfer buffer B into 5.8 ml transfer buffer A, and the final [Ca2+] is 0.06 mM. 2. Transfer Solution 2: Add 0.8 ml transfer buffer B into 5.2 ml transfer buffer A, and the final [Ca2+] is 0.24 mM. 3. Transfer Solution 3: Add 2 ml transfer buffer B into 4 ml transfer buffer A, and the final [Ca2+] is 0.6 mM. 4. Transfer Solution 4: Add 4 ml transfer buffer B into 2 ml transfer buffer A, and the final [Ca2+] is 1.2 mM.

3 3.1

Methods System Setup

1. The Langendorff Perfusion System (9): Assemble the instruments as indicated in Fig. 1. Make sure the whole system has no bubbles. Set the flow rate at 3 ml/min at the cannulation end (see Note 9). 2. Turn on the circulating water bath (Fig. 1e). Set the temperature of the water bath such that the temperature of the perfusion buffer at the cannulation end is at 37  C. 3. Cannulation Apparatus (Fig. 3d): Make knots with two twist using 6–0 black-braided silk nonabsorbable suture (Fig. 3c). Connect the 20 G blunt-ended needle (Fig. 3a) onto a 1 ml

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syringe (Fig. 3b) and load about 500 μl of perfusion buffer to the syringe (Fig. 3b). Eliminate any bubbles from the syringe and then put the syringe onto the syringe holder (Fig. 3d). Add about 20 ml perfusion buffer to a 60
15 mm petri dish. Place the syringe on top of the petri dish such that the 20 G bluntended needle is immersed in the perfusion solution (Fig. 3d). 4. Keep perfusion buffer and digestion buffer in 100% O2 bubbling for the whole isolation process. 3.2 Heart Cannulation and Cardiomyocytes Isolation

1. Intraperitoneal inject 200 IU heparin sodium per mouse (about 6 weeks old) to prevent blood clotting before anesthetization. Allow about 10 min for the heparin to distribute through the bloodstream (see Note 10). 2. Anesthetize mouse with 2.5% isoflurane and 100% O2 by inhalation until reaching deep anesthetization with no response to a toe pinch. 3. Wipe mouse chest area with 75% ethanol and cut the skin open to expose the rib cage. Use the big forceps to grab the inferior end of the sternum and cut along the diaphragm’s anterior edge to open the thoracic cavity using the big scissors. Use the small scissors to cut the rib cage open to expose the thoracic cavity and the heart (see Note 11). 4. Remove heart by cutting off lungs and surrounding vessels carefully. Immediately immerse the heart in a 10 ml beaker, which contains 3–5 ml of perfusion buffer (see Note 12). 5. Transfer heart to the cannulation plate containing perfusion buffer (Fig. 3d). Make sure the entire heart and main vessels are submerged by the perfusion buffer to avoid air entering the heart. Using the fine-tipped forceps, cannulate the heart onto the 20 G blunt-ended needle through aorta vessel under a dissecting microscope. Leave the needle tip right above the aortic valves to allow for coronary perfusion. Proper perfusion through the coronary arteries will help digestion of the right ventricle. Firmly tie the aorta vessel onto the needle using the suture (Fig. 5a). Once the heart is secured, slowly inject the 500 μl perfusion buffer into the heart through the syringe attached to wash the remaining blood from the heart and vessels. Avoid introducing air bubbles into the heart and vessels (Fig. 5a). 6. Quickly trim off extra tissue if necessary. Remove the needle with the cannulated heart from the syringe and immediately fasten to the Langendorff perfusion system. Avoid introducing air bubbles (Fig. 5b). 7. Perfuse the heart with perfusion buffer for 3 min (see Note 13).

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Fig. 5 Heart cannulation

8. After 3 min of perfusion, switch to digestion buffer. Carefully monitor the digestion process. The heart should gradually become pale from the root to the tip. Gently squeeze the heart once every minute and collect a few drops of perfusate into a clean 60
15 mm petri dish and observe under an inverted microscope. Stop digestion when a single cardiomyocyte is present in the drop. The heart tissue should feel soft with no ventricular resistance (see Note 14). 9. Quickly remove the heart from the Langendroff perfusion system and place it into a 60
15 mm dish containing 5 ml transfer buffer A. Trim off extra tissue and vessels. Starting from the aortic opening of the left ventricle, gently tear the pericardium with fine-tipped forceps and tease the heart into 4–6 pieces (see Note 15). 10. Further dissociate the heart tissue using a plastic Pasteur pipette, whose tip had been cut off at an ~45 angle. Triturate up and down several times until the tissue pieces can easily enter the pipette (see Note 16). 11. Switch to a normal plastic Pasteur pipette. Pipette several times until the large pieces become smaller tissue strips. 12. Transfer the cell suspension into a 50 ml Falcon tube through a 250 μm filter top (Fig. 4). Rinse the petri dish with 3 ml more transfer buffer A and transfer to the same Falcon tube. 13. Place the Falcon tube upright and allow the cells to sediment by gravity for 15 min. 14. After 15 min, carefully remove the supernatant without irritating the cells.

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15. Add 5 ml of transfer solution 1(0.06 mM [Ca2+]) to suspend the cells and mix by gently shaking the tube. 16. Repeat steps 14 and 15 with transfer solution 2 (0.24 mM [Ca2+]), 3 (0.6 mM [Ca2+]), and 4 (1.2 mM [Ca2+]). 17. After adding transfer solution 4, the isolated cardiomyocytes are ready for further culture or other experiments (see Note 17).

4

Notes 1. Can be made by removing the sharp tip of the regular 20 G needle and then smoothing the cut end using a sandstone. 2. Anything that can hold the 1 ml syringe to free your hands for the heart cannulation. 3. Basal solution can be made the day before the cardiomyocyte isolation experiment in a 1 L bottle and leave in 37  C water bath over night with the cap tightened. If make the day of experiment, leave the basal solution in 37  C water bath for at least 30 min to bring the temperature up to 37  C. 4. It is recommended to use a hot plate stirrer while making the perfusion solution and adjusting the pH. Set the stirrer at a desired speed and the hot plate at 37  C. Move basal solution from the 37  C water bath to the hot plate stirrer. Add D-(+)Glucose, BDM, and taurine to basal solution to make perfusion buffer as indicated in “Subheading 2.3, item 2” After adjusting the pH, bubble the perfusion buffer with 100% oxygen throughout the entire isolation procedure. One 3-month-old WT mouse heart needs about 155 ml of perfusion buffer to make digestion buffer, transfer buffer A, transfer buffer B, and to perfuse the heart. Thus, if starting with three mice hearts, making 500 ml perfusion buffer is suggested. Perfusion buffer of 1 l is good for six mice hearts. 5. Bubble the digestion buffer with 100% O2 throughout the duration of the isolation procedure. One 3 months old WT mouse heart needs about 25 ml of digestion buffer when the perfusion rate is set at 3 ml/min. 6. One mouse heart needs about 8 ml of transfer buffer A to neutralize the digestion buffer and 17 ml to make the transfer solution with increasing calcium concentration. 7. One mouse heart needs 7 ml of transfer buffer B to make the transfer solution with increasing calcium concentration. 8. This recipe is for one mouse heart, which is 6 ml of each transfer solution.

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9. Before getting started, run the perfusion system with 75% ethanol for 5 min with the setting of the MINI-PUMP variable flow instrument at fast mode, then switch to ddH2O for 5 min. After ddH2O wash, switch to perfusion buffer and set the flow mode to slow. Adjust the flow speed to get the flow rate at 3 ml/min. 10. Make 1 IU/μl heparin in PBS. Inject about 200 μl per mouse. 11. Be careful not to accidentally cut the heart at this step. 12. Leave as much of the ascending aorta as possible to facilitate cannulation. Keeping the thymus gland with heat helps to identify the aorta vessel when cannulating. The aorta arch is right under the thymus when separating the two lobes of the thymus gland. 13. One can gently squeeze the heart to help remove the extra blood. 14. If the right ventricle area remains pink while the rest of the heart appears pale, it indicates that the needle tip was below the coronary opening such that the coronary was not properly perfused and the right ventricle was not well-digested. 15. A well-digested heart tissue will have cotton-like morphology, and cells start to dissociate when the pericardium is lightly torn. 16. Avoid generating bubbles. The transfer solution A starts to turn cloudy as more cells are dissociated. Modification of the pipette tip will decrease the shear stress on the cells during the dissociation process. 17. One can estimate the yield of the cardiomyocytes at this point by adding a few microliters of the cell suspension to a petri dish and observing under an inverted light microscope. A live cardiomyocyte should have a rod shape, and an injured or dead myocyte will become round (Fig. 6). This protocol generally yields about 80% of rod-shaped cardiomyocytes.

Fig. 6 Isolated adult cardiomyocytes

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References 1. Woodcock EA, Matkovich SJ (2005) Cardiomyocytes structure, function and associated pathologies. Int J Biochem Cell Biol 37:1746–1751 2. Ibrahim M, Gorelik J, Yacoub MH, Terracciano CM (2011) The structure and function of cardiac t-tubules in health and disease. Proc Biol Sci 278:2714–2723 3. Piquereau J, Caffin F, Novotova M et al (2013) Mitochondrial dynamics in the adult cardiomyocytes: which roles for a highly specialized cell? Front Physiol 4:102

4. Louch WE, Sheehan KA, Wolska BM (2011) Methods in cardiomyocyte isolation, culture, and gene transfer. J Mol Cell Cardiol 51:288–298 5. Powell T, Twist VW (1976) A rapid technique for the isolation and purification of adult cardiac muscle cells having respiratory control and a tolerance to calcium. Biochem Biophys Res Commun 72:327–333 6. Liao R, Jain M (2007) Isolation, culture, and functional analysis of adult mouse cardiomyocytes. Methods Mol Med 139:251–262

Chapter 17 Analysis of Lymphatic Vessel Formation by Whole-Mount Immunofluorescence Staining Jian Wang, Yuwei Dong, Mariappan Muthuchamy, David C. Zawieja, and Xu Peng Abstract Pathological alterations of lymphatic structure and function interfere with lymph transport, resulting in a wide range of clinical disorders that include edema, tissue inflammation, and metabolic syndromes. Mesentery contains abundant lymphatic vessels and plays an important role in transporting absorbed lipid from the intestine. In this manuscript, we describe a whole-mount staining method on isolated mouse mesentery with VEGFR3, Prox1, and Lyve1 antibodies to visualize the morphology of lymphatic vessels. Keywords Lymphangiogenesis, Lymphatic endothelial cells, Whole-mount staining, Immunofluorescence staining

1

Introduction The lymphatic system plays an important role in tissue fluid balance, lipid transportation, and immunological surveillance [1– 3]. Lymphangiogenesis, the formation of new lymphatic vessels from preexisting vessels, is not only essential for embryo development, but also involved in the pathogenesis of many diseases, including cancer metastasis, diabetes, and other infectious diseases [4]. Insufficient lymphangiogenesis and/or dysfunctional lymphatic vasculature results in lymphedema and impaired immunological response. Unfortunately, therapeutic options for lymphedema are still limited. It is of great importance to unravel the underlying mechanisms of lymphangiogenesis to develop novel therapeutic strategies that target abnormal lymphatic development and function. Compared to the blood vessels, which are relatively even in the lumen, lymphatic vessels are morphologically nonuniform. Thus, whole-mount staining appears to be a valuable technique, as it is

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capable of visualizing whole lymphatic vessels, providing comprehensive information. Given that lymphangiogenesis occurs in the mid to late gestational stage, it is technically challenging to perform whole-mount staining on whole embryos since the embryos are no longer transparent, and it is more difficult for antibodies to penetrate. However, specific organs such as the mesentery can serve as a adequate model to investigate lymphangiogenesis. The mesentery, a transparent membrane attached to the gut wall of the abdominal cavity, serves as a major conduit for intestinal lymphatics. It is a preferable model because of its abundance in lymphatic vessel vascularization and minimal background interference when staining [5]. Moreover, intraluminal lymphatic valves present in the collecting lymphatic vessels can be visualized by specific markers (Prox1, FoxC2, Integrin α9, etc.) in the mesentery, which enables the mesentery to be a model for lymphatic valve formation study [6]. It is believed that a subset of the endothelial cells in cardinal veins gives rise to the building blocks of the lymphatic vasculature, lymphatic endothelial cells (LECs). These LECs form the lymph sacs, the source from which the entire lymphatic vasculature is also derived. We are currently able to specifically label LECs, largely due to deeper understandings of the molecules involved in lymphangiogenesis and advances in lymphatic endothelium-specific marker identification during the past few decades. The most frequently used markers are Prox1, Lyve1, VEGFR3, and podoplanin. Prox1, a transcription factor, is the master regulator controlling LEC differentiation [7]. Prox1 expression level is significantly higher in the lymphatic valve cells, compared to the lymphangion, and is a good marker of lymphatic valve forming cells [6]. Lyve1, a hyaluronan receptor in LECs, is specifically expressed in LECs but not in the blood endothelial cells (BECs). Lyve1 can serve as a lymphatic vessel marker although it plays a dispensable role in lymphatic development [8, 9]. In lymphatic capillary, Lyve1 expression level is higher than that of collecting lymphatic vessels. VEGFR3 is present in LECs and is essential for lymphatic development [10]. BECs also express VEGFR3 before the onset of lymphatic development, but its expression is downregulated in mid-gestation during embryogenesis and becomes restricted to the lymphatic endothelium after mid-gestation [11, 12]. Podoplanin plays an essential role in lymphatic development and is expressed by the LECs that have migrated out of the cardinal vein [13]. Podoplanin is expressed at similar levels in the collection and capillary LECs. In this protocol, we aim to describe detailed methods about how to properly isolating embryonic skin and mesentery from embryonic mice and how to perform whole-mount immunofluorescence staining by using LEC-specific markers, including Prox1, Lyve1, VEGFR3, and podoplanin.

Analysis of Lymphatic Vessel Formation by Whole-Mount Immunofluorescence. . .

2 2.1

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Materials 1. 1 Phosphate-Buffered Saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Ha2HPO4, and 2 mM KH2PO4. Adjust the pH to 7.4 with HCl and store the buffer at room temperature.

Reagents

2. 4% paraformaldehyde (PFA) in PBS. 3. PBST: 0.075% Triton X-100 in PBS. 4. Antifade mounting media (Invitrogen). 5. Blocker: 1% bovine serum albumin (BSA). 6. 5% fetal bovine serum in PBS. 7. Primary antibodies (see Table 1). 8. Secondary antibodies (see Table 2).

Table 1 Primary antibodies for whole-mount staining of the mesentery Primary antibody

Marker

Host Species Company

Dilution

PECAM-1/ CD31

Pan endothelial cell marker

Rat

BD Pharmingen (550274 or 553370)

1:100

LYVE-1a

Lymphatic endothelial cell marker

Rabbit

Abcam (ab14917)

1:500

VEGFR3

Lymphatic endothelial cell marker

Goat

R&D System (AF743)

1:100

Prox1

Lymphatic endothelial cell marker

Rabbit

AngioBio (11-022P)

1:100

Podoplanin

Lymphatic endothelial cell marker

Syrian hamster

AngioBio (11-033)

1:100

a

LYVE1 antibodies can label lymphatic endothelial cells and a subset of macrophages

Table 2 Frequently used immunofluorescence-labeled secondary antibodies Secondary antibody

Source

Reaction

Company

Dilution

AlexaFluor-594

Goat

Goat anti-rat

Invitrogen

1:200

AlexaFluor-488

Goat

Goat anti-rabbit

Invitrogen

1:200

AlexaFluor-488

Donkey

Donkey anti-goat

Invitrogen

1:200

AlexaFluor-594

Donkey

Donkey anti-Rat

Invitrogen

1:200

AlexaFluor-488

Goat

Goat anti-hamster

Invitrogen

1:200

AlexaFluor-647

Goat

Goat anti-hamster

Invitrogen

1:200

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Methods

3.1 Harvest the Mesentery from Time-Crossed Pregnant Female Mice

1. Euthanize the pregnant female with carbon dioxide, followed by cervical dislocation. Then cut open the abdominal cavity and isolate the embryos from the uterus (see Note 1). 2. Humanely decapitate pups and wash off the blood. 3. Pin down the limbs of embryos on a Sylgard-coated petri dish containing ice-cold PBS. Cut open the abdomen and exteriorize the small intestine attached by the mesentery. 4. Cut the intestine and mesentery submerged in PBS (see Note 2). 5. Pin the intestine down onto the dish in a loop to fully spread the mesentery (see Note 3).

3.2 Whole-Mount Immunofluorescence Staining

1. Rinse the tissue with PBS 1–2 times to remove the debris and damaged tissues. 2. Use 4% PFA to fix the tissue overnight at 4  C on the orbital shaker (see Note 4). 3. On the orbital shaker, wash the tissue with PBS four times at room temperature for 5 min each time (see Note 5). 4. Permeabilize the mesenteric preparation with 0.075% Triton X-100 in PBS for 15–20 min at room temperature on the orbital shaker (see Note 6). 5. Wash the tissue with PBS four times for 5 min each at room temperature on the orbital shaker. 6. Block the tissue with 5% fetal bovine serum in 1% BSA for 2 h at room temperature on the orbital shaker (see Note 7). 7. Remove the pins and transfer the samples from the Sylgardcoated petri dish to a 24-well cell culture plate (see Note 8). 8. Perform the appropriate dilution of primary antibody in 5% fetal bovine serum in 1% BSA (see Table 1) and gently agitate the tissues overnight at 4  C on the orbital shaker. 9. Wash the tissue with PBS four times at room temperature on the orbital shaker, for 15 min each time. 10. Perform the appropriate dilution of secondary antibody in 5% fetal bovine serum in 1% BSA and incubate the tissues for 2 h at room temperature on the orbital shaker. 11. On the orbital shaker, wash the tissue with PBS six times for 15 min each time at room temperature. 12. Pin the gut back onto the Sylgard-coated petri dish and remove the gut wall (see Note 9). 13. Spread the mesentery on a glass slide and leave the sample at room temperature for several minutes to dry (see Note 10).

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Fig. 1 Immunostaining on E17.5 mesentery with CD31 and Prox1 antibodies. The isolated mesentery was fixed with 4% PFA and then stained with CD31 (a) and Prox1 (b). Panel (c) is the merged image

14. Apply the ProLong antifade reagent on the samples and cover samples with cover glasses. 15. Allow samples to dry overnight at room temperature before observing. 16. Observe the mesentery with fluorescence or confocal microscopy (Fig. 1).

4

Notes 1. To acquire a well-formed mouse mesentery, it is typically better to euthanize the pregnant female when the embryos are at least E15.5 or older. 2. Since the root of the mesentery is hard to identify, gently pull the intestine out of the abdomen as much as possible and cut at the very end of the mesenteric root. Additionally, embryonic mesenteries are extremely fragile, particularly at E15.5–E16.5. If performing the procedure at earlier stages is difficult, use P1– P3 pups as practices first. 3. It is challenging to pin the intestine down since the gut wall is thin and slippery. Thus, it is most effective to begin pinning at either end of the intestine. Furthermore, the larger the intestine loop is, the more vasculature information can be obtained post staining, and the easier it will be to manipulate in the following processes. 4. To accelerate the process, you can alternatively fix the tissue at room temperature for 1 h on the orbital shaker. It is important to wrap the dishes with polyvinyl-chloride membranes to prevent PFA evaporation and to make sure that the fixative is adequate so that the tissue will not dry out.

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5. The main purpose of this process is to get rid of the fixative since it is able to affect the following staining. It is better to add a large amount of PBS each time and speed up the orbital shaker. 6. The purpose of this step is to make the cell membrane more permeable for the antibodies. Timing is important for this step. Either insufficient permeabilization or over permeabilization could cause unsatisfactory staining results. 7. If staining could not be performed immediately after blocking, keeping the tissue in the blocking buffer (PBS containing 5% fetal bovine serum and 1% BSA) overnight at 4  C would be an alternative. 8. Be very careful when handling the gut wall and removing the pins since the mesentery attached to the gut wall is thin and easily broken. It is useful to press the gut wall down onto the dishes when pulling out every pin, which should protect the mesentery from being torn. 9. This is the most challenging step of this protocol since after the processes, the mesenteries become even more fragile. Furthermore, blood and lymphatic vasculature are abundant on the boundaries between the mesentery and the gut wall. To maintain as much of the mesentery as possible, it is better to leave a little bit of gut wall on the boundaries when cutting off the mesentery. 10. The best time to manipulate and make the mesentery spread well is when the PBS is about to dry out. Do not try to move the mesentery after PBS is completely dry.

Acknowledgments This work was supported by American Heart Association Transformational Project Award (19TPA34900011) to Xu Peng. References 1. Semo J, Nicenboim J, Yaniv K (2016) Development of the lymphatic system: new questions and paradigms. Development 143(6):924–935 2. Oliver G (2004) Lymphatic vasculature development. Nat Rev Immunol 4(1):35–45 3. Zheng W, Aspelund A, Alitalo K (2014) Lymphangiogenic factors, mechanisms, and applications. J Clin Invest 124(3):878–887 4. Alitalo K, Tammela T, Petrova TV (2005) Lymphangiogenesis in development and human disease. Nature 438(7070):946–953

5. Mahadevan A, Welsh IC, Sivakumar A, Gludish DW, Shilvock AR, Noden DM et al (2014) The left-right Pitx2 pathway drives organ-specific arterial and lymphatic development in the intestine. Dev Cell 31(6):690–706 6. Sabine A, Agalarov Y, Maby-El Hajjami H, Jaquet M, Hagerling R, Pollmann C et al (2012) Mechanotransduction, PROX1, and FOXC2 cooperate to control connexin37 and calcineurin during lymphatic-valve formation. Dev Cell 22(2):430–445

Analysis of Lymphatic Vessel Formation by Whole-Mount Immunofluorescence. . . 7. Wigle JT, Oliver G (1999) Prox1 function is required for the development of the murine lymphatic system. Cell 98(6):769–778 8. Gale NW, Prevo R, Espinosa J, Ferguson DJ, Dominguez MG, Yancopoulos GD et al (2007) Normal lymphatic development and function in mice deficient for the lymphatic hyaluronan receptor LYVE-1. Mol Cell Biol 27(2):595–604 9. Luong MX, Tam J, Lin Q, Hagendoorn J, Moore KJ, Padera TP et al (2009) Lack of lymphatic vessel phenotype in LYVE-1/CD44 double knockout mice. J Cell Physiol 219 (2):430–437 10. Zhang Y, Ulvmar MH, Stanczuk L, MartinezCorral I, Frye M, Alitalo K et al (2018) Heterogeneity in VEGFR3 levels drives lymphatic vessel hyperplasia through cell-autonomous

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and non-cell-autonomous mechanisms. Nat Commun 9(1):1296 11. Kukk E, Lymboussaki A, Taira S, Kaipainen A, Jeltsch M, Joukov V et al (1996) VEGF-C receptor binding and pattern of expression with VEGFR-3 suggests a role in lymphatic vascular development. Development 122 (12):3829–3837 12. Dumont DJ, Jussila L, Taipale J, Lymboussaki A, Mustonen T, Pajusola K et al (1998) Cardiovascular failure in mouse embryos deficient in VEGF receptor-3. Science 282(5390):946–949 13. Schacht V, Ramirez MI, Hong YK, Hirakawa S, Feng D, Harvey N et al (2003) T1alpha/podoplanin deficiency disrupts normal lymphatic vasculature formation and causes lymphedema. EMBO J 22(14):3546–3556

INDEX A

H

Adult cardiomyocytes isolation ........................... 143–151 Age-related macular degeneration (AMD)...............5, 77, 78, 83 Angiogenesis......................................... 61–67, 78, 87, 88, 91, 93, 103 Animal..................................2–4, 6, 8, 10, 18, 33–35, 63, 78–80, 82, 83, 113–117, 128, 139, 144 Animal model ......................................................... 52, 111

Hindbrain ..................................................................87–92 Histamine ..................................................................45–49

B Bacterial artificial chromosomes (BAC)............. 120–129, 131, 132, 135 B-galactosidase ..................................................... 130, 135 Blood vessel ............................ 10, 61, 62, 66, 80, 87, 88, 91, 93–103, 111–117, 153

C Cardiac ................................ 6, 10, 18, 31, 42, 43, 52, 55, 57, 58, 137, 143, 144 myocytes ............................................ 31, 32, 143, 144 myofibrils ............................................................. 15–23 Cardiomyocyte ........................................ 51–59, 143–151 Cell culture ............................. 26, 27, 46, 48, 63, 70, 74, 89, 138–140, 156 Cell isolation.......................................................................v Confocal scanning laser ophthalmoscopy (cSLO) .....111, 112, 114, 115 Contractile responses ...................................31, 32, 40–42

D Disease-modeling ......................................................52, 83 Dorsal skin .................................................................61–67

E

I Immunofluorescence ................................ 53, 55, 89, 106 Immunofluorescence staining .......................90, 153–158 Induced pluripotent stem cells (iPSCs) ................. 51, 52, 54, 55, 58 Isolated cardiac tissues .................................................... 42 Isolation ............................ 17, 18, 32, 33, 35–38, 41, 42, 122, 125, 127, 128, 131, 135, 143, 144, 148, 149

L Langendorff.......................................................... 143–152 Laser capture microdissection (LCM) ......................... 105 Laser-induced choroidal neovascularization (CNV) .... 77, 78, 81–83 Lightsheet microscopy ............................................93–103 Lymphangiogenesis ............................................. 153, 154 Lymphatic endothelial cells (LECs)...........25–30, 45–49, 69–75, 138, 154, 155 Lymphatic muscle cells (LMCs).......................... 137–141

M Magnetic separation ........................................... 70, 72–74 Mass spectrometry (MS).................................... 16, 21–23 Mouse ............................................ 2–6, 8, 10, 18, 19, 31, 42, 61–67, 69–74, 78, 84, 87–91, 93–103, 113–116, 120, 121, 124, 126–128, 143–151, 154, 156, 157 Mouse tissue .................................................................... 74

N Neonatal rat heart .....................................................34, 36

Embryo culture .......................................................93–103

F Fluid shear stress .......................................................25–29 Fluorescein angiography (FA) .............................. 83, 111, 112, 115

P Papillary muscles .......................................................31–43 PEGASOS.................................................................... 1–11 Permeability........................................................ 23, 45, 46 Protein isolation .............................................................. 18

Xu Peng and Warren E. Zimmer (eds.), Cardiovascular Development: Methods and Protocols, Methods in Molecular Biology, vol. 2319, https://doi.org/10.1007/978-1-0716-1480-8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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CARDIOVASCULAR DEVELOPMENT: METHODS AND PROTOCOLS

162 Index

Protein separation ........................................................... 21 Protocol ...................................2, 3, 7, 15, 17, 18, 21–23, 29, 54, 62, 66, 70, 74, 82, 121, 127, 128, 131, 135, 143, 144, 151, 154, 158

R Rat mesentery....................................................... 137–141 Red E/T recombineering ............................................. 125 RNA isolation....................................................... 106–108 RNA-sequencing (RNA-Seq) ...................................25–29

Transcriptome ...........................................................25–29 Trans-epithelial/endothelial electrical resistance (TEER) assay ....................................................... 45 Transient transgenics ................................. 120, 121, 123, 127, 128, 131 Transwell....................................................................45–48 Trypsin ....................................26, 27, 29, 46–48, 74, 140

V

Spectral-domain optical coherence tomography (SD-OCT) ................................................ 112, 116 SPIM ....................................................................... 94, 100

Vascular endothelial cells ...............................87, 105–109 Vasculature.......................... 1–11, 61, 62, 65, 69, 87, 93, 94, 102, 103, 111, 114, 153, 154, 157, 158 Vasculogenesis .................................. 61, 87, 93, 102, 103 Vibratome section .....................................................90, 91 VISTA ............................................................................ 121

T

W

3-D imaging ........................................................... 2, 8, 11 Tissue clearing ............................................................... 1–3

Whole-mount staining ............................ 63, 88, 154, 155 Wnt inhibition ................................................................. 53

S