Kidney Research: Experimental Protocols (Methods in Molecular Biology, 466) 1588299457, 9781588299451

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Kidney Research

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For other titles published in this series, go to www.springer.com/series/7651

Kidney Research Experimental Protocols

Edited by

Tim D. Hewitson, Ph.D. The Royal Melbourne Hospital, Melbourne, VIC, Australia

Gavin J. Becker, M.D. The Royal Melbourne Hospital, Melbourne, VIC, Australia

Editors Tim D. Hewitson Ph.D. The Royal Melbourne Hospital Parkville, VIC Australia

Gavin J. Becker The Royal Melbourne Hospital Parkville, VIC Australia

ISBN: 978-1-58829-945-1 e-ISBN: 978-1-59745-352-3 DOI: 10.1007/978-1-59745-352-3 Library of Congress Control Number: 2008927373 © 2009 Humana Press, a part of Springer Science + Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface

There is a growing realisation that ultimately the world will not be able to afford the expense of treating the ever-increasing number of patients with end-stage kidney disease. This has focused even greater attention on strategies to prevent and treat early kidney disease, and to slow its progression. Application of these strategies relies on clarification of the mechanisms involved, which in turn are largely dependent on experimental studies in nephrology. Does nephrology research differ from other medical research? In an age where techniques in cell biology, pathology, biochemistry, and genetics are applied so generically, it seems reasonable to assume that laboratory studies in nephrology do not differ greatly from any other field. However, the study of renal disease has a number of inherent problems, most notably the complexity of renal anatomy and physiology, and the associated plethora of different cell types found in the kidney. This has therefore meant that laboratory researchers have had to develop a number of specialised techniques. It is these problems and solutions to address them that form the basis of this book. In editing this volume we have attempted to collate both well-established and novel methods used in the study of experimental kidney disease. Part I of the book covers the preparation and culture of the main cell types used to study the mechanisms of renal disease. The three chapters in this section provide detailed guidance on cell culture methods in general and the propagation of mesangial cells (Chap. 1), tubules (Chap. 2), and fibroblasts (Chap. 3). In Part II we provide a critical review of the common animal models used to mimic the various forms of human renal disease (Chap. 4). For the purposes of our discussion, the review is organised by initiating factor; surgical, toxic, immunemediated, and metabolic injuries. The bibliography provides key examples of the use and application of these models. We hope that our emphasis on mouse techniques is useful to those seeking to take advantage of transgenic and knock-out mice. The third and final section of the book (Part III) describes a number of specific applications and techniques used in vivo and in vitro. Consistent with our emphasis on murine models, the first chapter in Part III (Chap. 5) provides detailed guidance on measuring renal function in mice. One of the major problems encountered in the study of renal disease is the fact that it often involves selective injury to the glomerular, vascular, tubular, and interstitial compartments. Protocols for in situ v

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Preface

hybridization and laser capture microdissection are therefore included. Chapters 8–13 cover a number of imaging techniques used to study renal pathophysiology including methods to observe leukocyte recruitment in real time, generic histochemical protocols, and applications to specifically localize tissue hypoxia and apoptosis. Chapters 14 and 15 provide in vitro designs that can be used to model the matrix contraction and glomerular stretch that occurs in progressive renal disease. Other chapters in this section detail methods for the quantitative analysis of gene expression, measurement of renal collagen, proteomics, and finally, a gene transfer strategy for targeting glomeruli. In many cases, the authors provide case studies as well as detailed protocols to illustrate their application. In editing this book, we have attempted to provide a collection of protocols useful to those with some laboratory experience in nephrology and those new to the field. We are grateful to all the authors for their generosity in sharing their expertise and experience for this book. Their technique notes in particular provide invaluable guidance for those seeking to establish these methodologies in their own laboratory. Tim D. Hewitson Gavin J. Becker

Contents

Preface ............................................................................................................... v Contributors ..................................................................................................... ix List of Color Plates........................................................................................... xiii Part I

Isolation and Propagation of Cell Populations

1

Isolation and Propagation of Glomerular Mesangial Cells.................... Paolo Menè and Antonella Stoppacciaro

1

2

Isolation and Primary Culture of Human Proximal Tubule Cells ........ David A. Vesey, Weier Qi, Xinming Chen, Carol A. Pollock, and David W. Johnson

19

3

Propagation and Culture of Renal Fibroblasts ....................................... Lauren Grimwood and Rosemary Masterson

25

Part II 4

Animal Models

Small Animal Models of Kidney Disease: A Review............................... Tim D. Hewitson, Takahiko Ono, and Gavin J. Becker

Part III

39

Techniques and Applications

5

Measurement of Glomerular Filtration Rate in Conscious Mice .......... Zhonghua Qi and Matthew D. Breyer

59

6

Laser Capture Microdissection of Kidney Tissue ................................... Robert P. Woroniecki and Erwin P. Bottinger

73

7

Quantitative Gene Expression Analysis in Kidney Tissues .................... Chris Tikellis, Philip Koh, Wendy Burns, and Phillip Kantharidis

83

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Contents

8

In Vivo Imaging of Leukocyte Recruitment to Glomeruli in Mice Using Intravital Microscopy ...................................................... 109 A. Richard Kitching, Michael P. Kuligowski, and Michael J. Hickey

9

Using In Situ Hybridization to Localize Renal Gene Expression in Tissue Sections ..................................................................................... 119 Ian A. Darby, Alexis Desmoulière, and Tim D. Hewitson

10

Immuno and Lectin Histochemistry for Renal Light Microscopy ...... 133 Tim D. Hewitson and Lauren Grimwood

11

Immuno and Lectin Histochemistry for Renal Electron Microscopy ................................................................................ 149 Mitsuru Nakajima

12

Pimonidazole Adduct Immunohistochemistry in the Rat Kidney: Detection of Tissue Hypoxia ................................... 161 Christian Rosenberger, Seymour Rosen, Alexander Paliege, and Samuel N. Heyman

13

Identification of Apoptosis in Kidney Tissue Sections .......................... 175 Glenda Gobe

14

Cell-Populated Floating Collagen Lattices: An In Vitro Model of Parenchymal Contraction .................................. 193 Kristen J. Kelynack

15

Mechanical Stretch-Induced Signal Transduction in Cultured Mesangial Cells .................................................................... 205 Joan Krepinsky

16

Determination of Collagen Content, Concentration, and Sub-types in Kidney Tissue.............................................................. 223 Chrishan S. Samuel

17

SELDI-TOF Mass Spectrometry-Based Protein Profiling of Kidney Tissue ....................................................................................... 237 Eleni Giannakis, Chrishan S. Samuel, Wee-Ming Boon, Mary Macris, Tim D. Hewitson, and John D. Wade

18

In Vivo Transfer of Small Interfering RNA or Small Hairpin RNA Targeting Glomeruli ....................................................................... 251 Yoshitsugu Takabatake, Yoshitaka Isaka, and Enyu Imai

Index .................................................................................................................. 265

Contributors

Gavin J. Becker, M.D. Department of Nephrology, The Royal Melbourne Hospital, Melbourne, Victoria, Australia Wee-Ming Boon, Ph.D. Department of Physiology, Monash University, Clayton, Melbourne, Victoria, Australia Erwin P. Bottinger, M.D. Department of Medicine, Mount Sinai School of Medicine, New York, NY, USA Matthew D. Breyer, M.D. Division of Nephrology, Vanderbilt University, Nashville, TN, USA Wendy Burns, B.Sc. Danielle Alberti Memorial Centre for Diabetes Complications, Baker Heart Research Institute, Melbourne, Victoria, Australia Xinming Chen, Ph.D. Department of Medicine, University of Sydney, Kolling Institute, Royal North Shore Hospital, Sydney, New South Wales, Australia Ian A. Darby, Ph.D Department of Nephrology, School of Medical Sciences, RMIT University, Bundoora, VIC, Australia Alexis Desmoulière, Ph.D. Department of Physiology, Faculty of Pharmacy, University of Limoges, Limoges, France Eleni Giannakis, Ph.D. Howard Florey Institute, The University of Melbourne, Victoria, Australia and Bio-Rad Laboratories, Hercules, CA, USA Glenda Gobe, Ph.D. Molecular and Cellular Pathology, School of Medicine, University of Queensland, Brisbane, Queensland, Australia

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Contributors

Lauren Grimwood, B.Sc. (HONS) Department of Nephrology, The Royal Melbourne Hospital, Melbourne, Victoria, Australia Tim D. Hewitson, B.Sc., Ph.D. Department of Nephrology, The Royal Melbourne Hospital and Department of Medicine, University of Melbourne, Melbourne, Victoria, Australia Samuel N. Heyman, M.D. Hadassah University Hospital Mt. Scopus and The Hebrew Medical School, Medicine, Jerusalem, Israel Michael J. Hickey, Ph.D. Centre for Inflammatory Diseases, Monash University Department of Medicine, Clayton, Melbourne, Australia Enyu Imai, M.D., Ph.D. Department of Nephrology, Osaka University Graduate School of Medicine, Osaka, Japan Yoshitaka Isaka, M.D., Ph.D. Department of Advanced Technology for Transplantation, Osaka University Graduate School of Medicine, Osaka, Japan David W. Johnson, M.D. Centre for Kidney Disease Research, Department of Medicine, University of Queensland, Princess Alexandra Hospital, Woolloongabba, Queensland, Australia Phillip Kantharidis, Ph.D. Danielle Alberti Memorial Centre for Diabetes Complications, Baker Heart Research Institute, Melbourne, Victoria, Australia Kristen J. Kelynack, Ph.D. Department of Nephrology, The Royal Melbourne Hospital, Melbourne, Victoria, Australia A. Richard Kitching, M.B. Ch.B., FRACP, Ph.D. Centre for Inflammatory Diseases, Monash University Department of Medicine, Clayton, Melbourne, Victoria, Australia Philip Koh, Ph.D. Danielle Alberti Memorial Centre for Diabetes Complications, Baker Heart Research Institute, Melbourne, Victoria, Australia Joan Krepinsky, M.D. McMaster University, St. Joseph’s Hospital, Hamilton, Ontario, Canada Michael P. Kuligowski, B.Sc. Centre for Inflammatory Diseases, Monash University Department of Medicine, Clayton, Melbourne, Australia

Contributors

Mary Macris, B.Sc. (HONS) Howard Florey Institute, The University of Melbourne, Victoria, Australia Rosemary Masterson, M.B., Ch.B., Ph.D. Department of Nephrology, The Royal Melbourne Hospital, Parkville, Victoria, Melbourne, Australia Paolo Menè, M.D. Division of Nephrology, Department of Clinical Sciences University of Rome “La Sapienza”, Rome, Italy Mitsuru Nakajima, M.D. Director of Pediatrics, Hoshigaoka Koseinenkin Hospital, Osaka, Japan Takahiko Ono, M.D. Department of Molecular Medicine, School of Pharmaceutical Sciences, University of Shizuoka, Shizuoka, Japan Alexander Paliege, Ph.D. Department of Anatomy, Humboldt University, Berlin, Germany Carol A. Pollock, Ph.D. Department of Medicine, University of Sydney, Kolling Institute, Royal North Shore Hospital, Sydney, New South Wales, Australia Weier Qi, B.Sc., M.Sc. Department of Medicine, University of Sydney, Kolling Institute, Royal North Shore Hospital, Sydney, New South Wales, Australia Zhonghua Qi, M.D., Ph.D. Division of Nephrology, Vanderbilt University, Nashville, TN, USA Seymour Rosen, M.D. Beth Israel Deaconess Medical Centre and Harvard Medical School, Pathology, Boston, MA, USA Christian Rosenberger, M.D. Charité Universitaetsmedizin, Nephrology and Medical Intensive Care, Berlin, Germany Chrishan S. Samuel, Ph.D. Howard Florey Institute, The University of Melbourne, Melbourne, Victoria, Australia Antonella Stoppacciaro, M.D. Department of Experimental Medicine and Pathology, University of Rome “La Sapienza”, Rome, Italy Yoshitsugu Takabatake, M.D., Ph.D. Department of Nephrology, Osaka University Graduate School of Medicine, Osaka, Japan

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Contributors

Chris Tikellis, B.Sc. (HONS) Danielle Alberti Memorial Centre for Diabetes Complications, Baker Heart Research Institute, Melbourne, Victoria, Australia David A. Vesey, B.Sc., Ph.D. Centre for Kidney Disease Research, Department of Medicine, University of Queensland, Princess Alexandra Hospital, Woolloongabba, Queensland, Australia John D. Wade, Ph.D., FRACI, FRSC Howard Florey Institute, The University of Melbourne, Melbourne, Victoria, Australia Robert P. Woroniecki, M.D. Department of Pediatrics, The Children’s Hospital at Montefiore, Albert Einstein College of Medicine, New York, NY, USA

List of Color Plates

Plate 1

Fig. 3.2 Staining characteristics of sub-cultured cells (passage 3); SMA, smooth muscle actin (reproduced from ref. (11) with permission from Elsevier) ...........................................

I-1

Fig. 4.1 Interstitial accumulation of monocytesmacrophages after unilateral ureteric obstruction (UUO). Micrograph shows immunoperoxidase staining with the monoclonal antibody clone ED-1, 3 days post-UUO .......................

I-2

Plate 3 Fig. 4.2 Increased immunoperoxidase staining for fibronectin at day 8 in an acute model of Thy-1 nephritis induced by rabbit ATS. Mesangial proliferation, matrix expansion, and the formation of a small crescent are clearly seen .................................................................................

I-2

Plate 4 Fig. 4.3 Severe crescent formation in a rat model of antiglomerular basement membrane (GBM) nephritis. Day 21, periodic-acid Schiff (PAS) stain (micrograph courtesy of Dr. Toshiaki Makino, Nagoya City University, Nagoya, Japan) ..................................................................................

I-3

Plate 5 Fig. 7.1 Schematic representation summarising the quantitative RT-PCR cycle .........................................................

I-3

Plate 2

Plate 6 Fig. 10.3a Immunohistochemical staining of rat glomerular and peritubular capillaries with mouse anti-rat JG12, a mouse monoclonal antibody to the endothelial cell enzyme aminopeptidase P. b Double labelling of proliferating myofibroblasts in an experimental model of renal infection. Cell proliferation is localised using

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Plate 7

Plate 8

Plate 9

Plate 10

List of Color Plates

DAB and a biotinylated antibody against bromodeoxyuridine (refer to ref. [9] for study design). Myofibroblasts are localised with a monoclonal antibody to smooth muscle actin, detected with alkaline phosphatase and Fast Red(tm). Co-localisation indicates myofibroblast proliferation, confirmed in this photograph by the presence of a mitotic figure (arrow). c and d Differential lectin labelling of tubules with biotinylated (c) PNA and (d) PHA-L using DAB as a chromogen ...........................................

I-4

Fig 14.1 Solidified fibroblast-populated collagen lattices (a) immediately after rimming with a scalpel blade (time 0) and (b) after 48 h. The comparison shows a 40% reduction in lattice diameter ...................................................

I-5

Fig. 15.2 Phosphorylated Raf-1 at Ser338 translocates to the membrane with stretch. MC are stretched for 5 min and phosphorylated Raf-1 at Ser338 visualized by immunofluorescence as outlined in Sect. 3.7. Arrows identify phosphorylated Raf-1 at cell membrane locations ........................................................................

I-6

Fig. 17.1 SELDI-TOF MS Analysis. Arrays with specific chromatographic properties (a) are equilibrated in binding buffer. Samples are applied to the array surface and incubated for 1 h (b). Arrays are washed to remove non-specifically bound proteins (c), followed by EAM application (d). The fraction of the proteome retained on the array is directly analysed by TOF-MS, resulting in a profile of proteins characterised by the m/z and signal intensities (e, f). Data is represented in spectra or virtual gel view (f). Figure adapted from ProteinChip(tm) technology training course (Bio-Rad Laboratories) .......................

I-7

Fig. 18.2 siRNA-mediated silencing of EGFP expression in the glomerular cells. siRNA targeting EGFP was transferred to the kidney of EGFP-transgenic rats via the renal artery using the electroporation method. Fluorescence micrographs of glomeruli in the siRNAtransfected (right) and contralateral (left) kidney were taken 7 days after transfection (upper panels). Sections were stained with Texas red-labeled OX-7 antibody, a marker of mesangial cells (middle panels), and the merged photos are shown in the lower panels (original magnification, ×400). In the transfected kidney, EGFP

List of Color Plates

expression was diminished substantially in almost all of the glomeruli (95%), whereas it was unchanged in the tubules. This inhibition seemed nearly complete in the mesangial cells, whereas in other glomerular cells, endothelial and epithelial cells, inhibition of EGFP expression was not observed. The reduction of mesangial EGFP expression was observed for up to 2 weeks, and had recovered completely by 3 weeks after transfection (data not shown) (reproduced from ref. (3)) ...................................

xv

I-8

Part I

Isolation and Propagation of Cell Populations

Chapter 1

Isolation and Propagation of Glomerular Mesangial Cells Paolo Menè and Antonella Stoppacciaro

Abstract Cultures of glomerular mesangial cells (MC) of rodent or human origin have been extensively employed in renal research laboratories since the early 1980s. Cultured MC retain extensive analogies with the fairly undifferentiated in vivo phenotype of an intercapillary mesenchymal cell population, i.e., a myofibroblast. MC proliferating in response to mitogens and growth factors can be growth-arrested by withdrawal of serum or 3D culture in collagen gels. They synthesize an extracellular matrix that includes interstitial collagens and has analogies with the glomerular basement membrane; a prominent cytoskeleton acts as a functional contractile apparatus. Cultured MC have been extensively employed as a tool for studying pathophysiological events such as mesangial expansion, scarring, and glomerulosclerosis. Current technology for MC isolation and culture is reviewed, with emphasis on methodological issues relevant to characterization, propagation, and long-term maintenance of homogeneous clones. Keywords Mesangial cell, Mesangium, Kidney glomerulus, Tissue culture, Myofibroblast, Mesenchymal cell

1

Introduction

Mesangial cells (MC) have been recognized as one the three major cell lines of the kidney glomerulus by electron microscopy in the second half of the 20th century through the independent work of H. Latta and G. Marinozzi (1, 2). MC have a spatial organization far less specialized than the endothelial and visceral epithelial cells lining the glomerular capillary wall. In fact, they seem to “fill the gap” between adjacent capillary loops surrounded by the glomerular basement membrane (GBM) and a network of visceral epithelial cells, i.e., the podocytes. The word “mesangium” indeed describes the intercapillary space within the glomerulus, and was coined by Zimmermann in 1929 (3). Their random distribution between endothelium and podocytes is further complicated by a fairly intermediate phenotype, sharing features of fibroblasts and smooth muscle cells as well.

From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_1, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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P. Menè, A. Stoppacciaro

The term “myofibroblast” has been often used to emphasize the uncommitted and nondifferentiated nature of this cell population (4, 5). Unlike podocytes, highly differentiated cells that express specific proteins such as nephrin, podocin, podocalyxin, NPHS1, ZO-1, etc. (6, 7), MC hardly display a mesangial-specific molecular marker. Indeed, the lack of an obvious phenotypic marker still plagues the field of MC culture, after more than 30 years of laboratory work trying to reproduce disorders that involve these cells in vivo with in vitro models (8). Elegant imaging studies and electron photomicrographs have shown that there are some 200–300 MC in an average rat glomerulus, visible as sparse nuclei embedded in narrow cytoplasms and trace amorphous extracellular matrix within a single mesangial space (1, 9). Three or fewer nuclei are generally observed in an intercapillary section of a normal murine or human glomerulus. Larger numbers of cells (hyperplasia) or more abundant cell bodies and/or extracellular matrix are the hallmark of a proliferative glomerular disorder. It seems that MC maintain direct contact with the GBM and occasionally with the overlying endothelium, thus, suggesting direct filtration of macromolecules and possibly deposition of plasma proteins in glomerular diseases (1, 4). MC have a prominent contractile apparatus, based on a network of microfilaments containing actin, myosin, and regulatory proteins. This has prompted speculation on their possible mechanical role on glomerular hemodynamics, perhaps synchronous to a similar response to angiotensin II and other vasoconstrictors in podocytes. The hypothesis has been put forward that MC contraction may impact on the glomerular ultrafiltration coefficient (Kf) in vivo by modulating hydraulic conductivity (Lp) and/or the capillary parietal area for filtration (A). Shunting of blood from capillaries narrowed by mesangial/podocyte contraction may underlie this theoretical decrease of A (1, 4, 5). Kriz and coworkers emphasized the tensile strength that a tonic contraction of MC may exert on the GBM, antagonizing the transcapillary pressure forces while holding together the entire network of capillary loops (9). MC may not only serve as the framework of the capillary tuft, but also participate in immunological or metabolic damage involving the glomerulus. Three types of structural change occur in response to mesangial damage. “Mesangiolysis” is induced by injection of anti-Thy 1.1 antiserum in a popular rat model of mesangial proliferative nephritis, with MC undergoing apoptosis and actually disappearing from the intercapillary space (10). Mesangial proliferation is often a prominent response to endothelial damage and vasculitis. Mesangial expansion occurs whenever increased matrix or other amorphous material accumulates without mitosis or migration of MC, as commonly seen in diabetes, amyloidosis, or light-chain deposition. Glomerulosclerosis is believed to result from prolonged mesangial expansion, although focal scarring of glomeruli may also be induced by podocyte injury, as in several forms of proteinuric glomerular diseases. It is unclear whether the two cell types cooperate in the deposition of an amorphous matrix or rather have independent phenotypic changes—as in the case of podocytes undergoing foot process effacement during proteinuria—eventually resulting in glomerular scarring.

1 Isolation and Propagation of Glomerular Mesangial Cells

1.1

5

Isolation and Propagation of Mesangial Cells

MC are the easiest cells to grow from glomeruli of mammals, avian species, and humans. Since 1970 at least three groups (11–13) employed standard culture techniques for mammalian cells, taking advantage of the minimal growth requirements and brisk proliferative potential of MC. When grown in one of the common media for eukaryotic cells (MEM, DMEM, Waymouth, RPMI 1640), these cells retain their in vivo phenotype as stellate, arborized elements that tend to form “hills and valleys” in monolayer culture (11–16). Homogeneous cultures of MC resemble smooth muscle cells or fibroblasts; when confluent, they tend to form a syncytium, with specialized junctions that have been examined by microinjection and lucifer yellow transfer (17). This too duplicates the in vivo setting, where an electrical continuity of MC with cells of the juxtaglomerular apparatus has been theorized (18). An unlimited number of passages in culture is common for rodent MC, while usually not more than 15 – 20 passages in culture can been obtained from human MC (14, 15, 18, 19). The reason is unclear, but probably relates to a greater adaptation to bidimensional culture on plastic surfaces, and thus phenotype transition, for rat MC. Human MC are either committed to a fixed number of divisions, or lack some substrate that ordinary culture media do not provide over the long term. Apparently, rat MC have lower requirements to support sustained proliferation, although the persistent need for substrate attachment indicates that they are not transformed in culture as certain neoplastic clones. Researchers have often emphasized the “inflammatory” conditions of tissue culture as opposed to the in vivo situation. MC (and other glomerular cells as well) are a largely quiescent population in vivo, with a slow turnover in healthy animals and individuals, unlike tubular epithelial cells of the kidney, which have a constant rate of proliferation (14, 15, 18–20). When grown in culture on a plastic surface in the presence of fetal bovine serum (FBS; a product of the platelet release reaction) MC become activated to proliferate and express phenotypic features that are different from those occurring in a healthy kidney in vivo. MC in culture undergo constant cycles of proliferation leading to exponential growth, until some sort of contact inhibition mechanism or antiproliferative mediator in the culture media halts repeated cell divisions. Moreover, loss of the normal 3D network formed by other types of glomerular cells populations (endothelial, epithelial cells) and by a structured GBM is likely to deprive the MC of cell-to-cell communication and regulatory influences, contributing to an artificial environment (6, 18, 21). Tissue culture techniques have been standardized through a 30-year routine in hundreds of laboratories throughout the world. Frozen stocks of cells circulate in many renal research units for scientific and even commercial purposes. Our own culture technique for rat cells (Table 1.1) is a modification of the method described by Striker et al. (14–25).

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P. Menè, A. Stoppacciaro

Table 1.1 Protocol for isolation of mesangial cells Four ether-anesthetized rats are exsanguinated through a midline incision after thorough disinfection of the abdomen Excise eight kidneys, which are decapsulated and placed in EBSS on ice Cut away cortical tissue from rat kidneys (or human kidneys not suitable for transplantation), mince with a razor blade to a paste-like consistency Perform sequential sieving of dispersed rat cortex through 105-µm-diameter brass or nylon filters (120-µm for human glomeruli), then collect glomeruli onto 75-µm-diameter nylon filters Wash, decapsulate glomeruli by forcing suspension 3× through a 21-gauge needle Wash, digest glomerular “cores” with collagenase for approximately 20 min Plate glomerular “cores” resuspended in 12 mL RPMI 1640 medium + 17% FBS + antibiotics + 0.1 U/mL insulin - 2 wells with 2 mL each in 3 plates Outgrowths of spindle-shaped cells at 6–36 h Characterize outgrowth at the time of first passage

2 2.1

Materials Isolation of Mesangial Cells

1. Four to eight kidneys obtained from ether-anesthetized rats (Sprague-Dawley, Wistar-Kyoto, etc.) at weaning, specimens weighing about 100–150 grams (see Note 1). 2. For human cells, 4- to 6-cm3 blocks of cortex cut with a scalpel from the cortex of kidneys not suitable for transplantation, or from the healthy segment of nephrectomy specimens (see Note 2). 3. Earle’s balanced salt solution (EBSS), 2× 100-mL bottles supplemented with 10 µg/mL ceftriaxone (Roche, Basel, Switzerland). 4. Complete set of presterilized (steam, gamma-rays, ethylene oxide, or overnight dipping in 70% ethyl alcohol) surgical instruments to excise kidneys from ether-anesthetized rats, including fine-tip tweezers, curved-tip scissors, and two stainless steel spatulas. 5. Iodopovidone and gauze to wipe abdomen prior to excision. 6. Disposable razor blades or sterile scalpels. 7. Petri dishes, 50-mm diameter, sterile. 8. 105-µm (human kidney, 120-µm)-mesh brass or nylon sieve mounted on a 250mm-diameter wooden/plastic circular frame. 9. 75-µm-mesh nylon sieve mounted on a cut-away plastic bottle neck, 50-mm diameter. 10. Plastic sterile 50-mL Falcon tubes. 11. 10-mL sterile plastic syringe with 21-gauge needle. 12. Sterile 1,000-mL manual Gilson / Eppendorf pipette tips or disposable 1,000mL plastic pipettes. 13. Collagenase: 750 U/mL Worthington type I (Worthington Biochemical, Lakewood, NJ, USA). Weigh powder and dissolve shortly before use in 2 mL Earle’s BSS.

1 Isolation and Propagation of Glomerular Mesangial Cells

7

14. Inverted stage light microscope, × 4 and × 10 objectives. 15. Rosewell Park Memorial Institute (RPMI) 1640 culture medium supplemented with 2 mM glutamine, 17% FBS, antibiotics (10 µg/mL ceftriaxone plus 100 µg/mL gentamicin), and 0.1 U/mL human recombinant insulin. 16. Plastic six-well sterile culture dishes.

2.2

Subculture of Mesangial Cells

1. 2. 3. 4.

Inverted stage light microscope, × 4 and × 10 objectives. Ca2+- and Mg2+-free phosphate-buffered saline (PBS) solution, pH 7.2. Trypsin solution: 0.05% trypsin in 0.02% EDTA in Ca2+- and Mg2+-free PBS. RPMI 1640 culture medium supplemented with 2 mM glutamine, 17% FBS, antibiotics (10 µg/mL ceftriaxone plus 100 µg/mL gentamicin), and 0.1 U/mL insulin. 5. Plastic six-well sterile culture dishes. 6. Plastic, disposable, individually wrapped sterile pipettes (1, 10, and 25 mL). 7. Sterilized fine long tip disposable glass micropipettes connected by a Teflon hose to a vacuum reservoir (to aspirate and remove spent culture media).

2.3 1. 2. 3. 4. 5.

Long-term Storage

9. 10.

Inverted stage light microscope, × 4 and × 10 objectives. MC cultures in early passages (passages 2–5) plated onto 75-cm2 tissue culture flasks. Ca2+- and Mg2+-free PBS solution, pH 7.2. Trypsin solution: 0.05% trypsin in 0.02% EDTA in Ca2+- and Mg2+-free PBS. RPMI 1640 culture medium supplemented with 2 mM glutamine, 17% FBS, antibiotics (10 µg/mL ceftriaxone plus 100 µg/mL gentamicin), and 0.1 U/mL insulin. Plastic 2-mL sterile polypropylene vials with rubber seal. Plastic disposable, individually wrapped sterile pipettes (1, 10, and 25 mL). Sterilized fine long tip disposable glass micropipettes connected to a vacuum reservoir (to aspirate and remove PBS washing). Dimethylsulfoxide (DMSO) stock solution. Liquid N2 storage tank.

3

Methods

6. 7. 8.

3.1

Protocol for Isolation

1. The kidneys of four to eight ether-anesthetized rats are excised with aseptic procedures after exsanguination by cutting the heart through a midline thoracoabdominal incision. Prior thorough disinfection of the skin is obtained by wiping

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2.

3.

4.

5.

P. Menè, A. Stoppacciaro

repeatedly with iodopovidone-soaked gauze. For human cells, 4- to 6-cm3 blocks of cortex are cut with a scalpel from surgical specimens. Kidneys (or fragments) are transferred to a sterile plastic 50-mL tube containing chilled EBSS supplemented with 10 µg/mL ceftriaxone (Roche), and decapsulated by traction with two fine-tip tweezers. After slicing the kidney in half vertically, cortical tissue is manually cut away, poured onto a Petri dish, and chopped into 1- to 2-mm cubes with fine-tip scissors. The kidney fragments are further minced with a razor blade into a paste-like preparation, which is then gently pressed with a spatula through a 105-µm mesh nylon or metal sieve (120 µm for human glomeruli). This glomeruli-enriched paste is collected from underneath the sieve with a second, smaller spatula repeatedly wetted with EBSS, and transferred to a 50-mL tube containing chilled EBSS. This diluted suspension is poured onto a second 75-µm filter and washed extensively with the same solution. Glomeruli and larger tubular fragments are retained on this filter, and transferred to a third 50-mL tube by gentle aspiration through a 1,000- or 5,000-µL automatic pipette tip (see Note 3). From this point on, maximum sterility must be ensured, although it is a safe habit to perform the entire procedure under a laminar hood (see Note 4). The glomerular suspension is aspirated with a 10-mL disposable syringe and force-pressed two to three times through a 21-gauge needle. This procedure will decapsulate about 75% of the glomeruli, which are then viewed on an invertedstage microscope for purity of the preparation. A yield of approximately 80% intact or fragmented glomeruli is acceptable. The glomerular suspension is then spun for 10 min at 900 × g, resuspended in a tube containing a previously prepared sterile 750 U/mL solution of Worthington type I collagenase in RPMI 1640 medium, and gently stirred at 37°C in a Dubnoff bath for 20 ± 5 min, checking digestion of the glomeruli on the microscope every 5 min. The glomerular “cores,” about half the size of the initial decapsulated tuft, are then spun again for 5 min, resuspended in 12 mL complete RPMI 1640 medium, and plated onto the middle two wells of three six-well dishes. The dishes are placed in a 37°C incubator with a controlled, humidified atmosphere of 95% air/5% CO2. Again, sterility of the media, incubator, disposable plastic ware, and laminar flow hood must be ensured throughout the procedure and all subsequent steps (see Note 4).

3.2

Protocol for Subculture and Cloning

1. After 6–12 h, the glomerular “cores” become adherent to the plastic surface, and centrifugal spreading of individual cells is immediately detectable by light microscopy (see Note 5). The medium is changed at 48 h, gently washing the dish to remove nonadherent cells, debris, and floating aggregates. Obviously, all procedures should be carried out under a laminar air flow hood ensuring maximum sterility.

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2. After 1–2 additional days, a subconfluent array of cobblestone-like patches of cells and spindle-shaped individual cells can be observed. At 7–10 days, stellate, spindle-shaped cells generally overtake the cobblestone of epithelial cells, and each set of two wells should be subcultured into a single six-well dish (1:3 split). 3. First, spent RPMI 1640 medium is aspirated without scratching the monolayer (e.g., from the side of the gently tilted dish). The cells are washed once for 60 sec with Ca2+- and Mg2+-free PBS solution, pH 7.2, followed by application of 1 mL/well trypsin solution (0.05% trypsin in 0.02% EDTA in Ca2+and Mg2+-free PBS) for 60 sec at 37°C (returning the dishes to incubator). Gently tapping the dish with the lid on helps detachment. On the microscope, cells can be seen to round up, detach from one another, and occasionally float. Most of the trypsin solution is then aspirated, the dish is transferred to the incubator for a further 3 min, and the enzymatic digestion is stopped by adding 2 mL/well complete RPMI 1640 medium (containing serum protease inhibitors). 4. The content of each well, usually an even suspension of rounded cells or small clusters of cells, is transferred to a sterile 50-mL plastic tube, diluted 1:3 or 1:4 with fresh media depending on the desired cell density, and plated onto appropriate flasks or culture dishes. Clumps of cells can usually be broken down by repeated pipetting prior to final plating (see Note 6 and below, “mesangial hillocks”). If needed for experiments, an aliquot of the suspension can be diluted and counted on a Bürker or Thoma chamber to adjust the number of cells to be plated. 5. Dilution cloning can be useful to select out homogeneous populations derived from a single cell or a small cluster of cells. It is usually done by increasing the split ratio to 1:10 or 1:20 and plating 200 µL of the resulting cell suspension onto each well of 96-well sterile dishes. The resulting clones, viewed by inverted-stage light microscopy, can be marked and individually trypsinized, to yield cultures resulting from divisions of a single cell. Cloning rings have also been used, but this procedure suffers from the complication of silicone-sealing each ring to the bottom of large Petri dishes, and the need to sterilize the entire assembly. Nevertheless, a certain degree of clonal selection cannot be avoided with time in culture, since cells with the fastest growth tend to outnumber those that proliferate more slowly or become quiescent. Moreover, those cells that display tighter adhesion to plastic surfaces are less easily propagated, since dislodgment by trypsin is less effective during subculture (see below). Usually, time in culture favors less adhesive cell populations, which probably display dysregulated matrix synthesis. 6. After the first two subcultures, the cell lines should be checked for purity and characterized according to the protocol in Table 1.2, usually confirming that a fairly homogeneous and pure (95–98%) population of MC has been obtained. The cells should be subcultured every 3 to 5 days, depending on the speed of replication, often very high in earlier passages of human cells (passages 2–8) and later passages of rat cells (from passage 10 on). This avoids the build-up of a very packed monolayer, in which individual cells can hardly be identified. When subcultured, superconfluent monolayers tend to be released by trypsin as

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“clumps” of cells rather than suspensions of individual elements. Upon plating, these clumps give rise to “hillocks” in subsequent cultures, that is, radial foci of clonal proliferation around a central core of fibrous, amorphous material (18–22, 25, 26). Obviously, this prevents the formation of a thin, even monolayer whenever this is required for microscopy, staining, microinjection, patch-clamp, or fluorometric recordings. On the other hand, hillocks have been useful as a model of the nodular deposition of extracellular matrix in various glomerular diseases (21, 26).

3.3

Characterization of the Cells

A unique and reliable marker of MC is yet to be identified, unlike glomerular epithelial and endothelial antigens. Therefore, establishing the mesangial origin of a cell line in culture is mostly based on exclusion criteria, as well as fulfillment of certain prerequisites (27, 28). Table 1.2 summarizes most accepted standards for a MC culture, including the lack of endothelial (factor VIII, CD34, AC-LDL binding, Weibel-Palade bodies, and angiotensin-converting enzyme activity) and epithelial markers (cobblestone morphology, “domes” of polarized cells, cilia, and various podocyte antigens, such as CD10, nephrin, podocin, NEPH1, ZO-1, podoplanin, α-actinin-4, etc.) (13–22, 27, 28, 29). Nephrotoxins such as puromycin aminonucleoside can be useful to rule out the presence of epithelial cells, most sensitive to aminoglycosides. Fibroblasts are apparently unable to grow when the media are supplemented with the d-isomer of valine only, unlike MC. Mitomycin C is reportedly toxic to MC, which can also be targeted by antisera against the Thy 1.1 epitope, constitutively expressed only by rat cells (5, 21). Markers suitable for immunofluorescence or immunoperoxidase staining include desmin, vimentin, and smooth muscle myosin and actin (SMA). It should be noted that the latter is fairly regulated by growth factors and cytokines, and may not be constitutive even in cells proliferating in response to fetal bovine serum. SMA has been often regarded as an indicator of a “myofibroblastic” phenotype, and accompanies

Table 1.2 Characterization of cultured mesangial cells Morphology: stellate, spindle-shaped, longitudinally oriented in bundles (“hills and valleys”) when confluent; no cobblestones, no “domes,” no cilia (epithelial features); no WeibelPalade bodies (endothelial features) Stains positive by immunofluorescence or immunoperoxidase: fibronectin, vimentin, Thy 1.1, cytokeratins, desmin, myosin; occasionally, -smooth muscle actin Stains negative by immunofluorescence or immunoperoxidase: Factor VIII, AC-LDL; CD45, CD10, Ia antigens, CD10, CD34 Enzymes: no angiotensin-converting enzyme Toxins: sensitive to mitomycin C, resistant to puromycin Growth: positive in d-valine-containing media Contractility by videomicroscopy: positive ANG II, vasoconstrictors

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mesangial proliferation and/or glomerular inflammation in vivo (5, 21, 29, 30). (Cyto)keratins are usually viewed as a phenotypic marker of epithelial cells, although a number of obvious mesangial preparations stain positive (see Note 7). Another issue that has often been debated is the presence of a “mesangial phagocyte,” that is, a blood-borne bone marrow-derived cell with features of a monocyte/ macrophage or of antigen-presenting cells. A small population (< 5% of total MC in a glomerulus) that stains positive for the leukocyte common antigen, CD45, and/ or the MHC Class II Ia antigen has been initially described by Kreisberg and Karnovsky (13), and later examined through bone marrow irradiation studies by Schreiner et al. (31). The likelihood that such cells survive and proliferate in culture is minimal, particularly since cell cultures are systematically negative for leukocyte markers. Interestingly, dendritic cells have been isolated from the renal mesangium of LEW.1A rats (32). On average, only two of these antigen-presenting cells, strongly stimulating allogeneic mixed leukocyte reactions, can be seen in the whole glomerulus (32).

3.4

Protocol for Freezing/Thawing Cultured MC

1. Typically, trypsinized cell suspensions (see above) are resuspended in full serum-containing, sterile filtered media supplemented with 10% (v/v) DMSO as a cryopreservative. Cell count is best adjusted to 1×106 cells/mL, and 1-mL aliquots are frozen in sterile polypropylene vials with rubber seals. 2. Freezing in liquid N2 preserves cells for an unlimited time (see Note 8). Slow freezing is usually accomplished by holding the vials in a –80°C freezer for 24 h before transferring them to the liquid N2 storage tank (see Note 9). When thawing, the vials are simply transferred to a 37°C Dubnoff bath; immediate thawing is not harmful. Cells are then slowly (10 min) diluted to 10 mL with fresh complete medium, in order to minimize the osmotic shock due to decreasing extracellular DMSO, and plated at the desired density (typically, 1 mL frozen stock to 12 mL RPMI 1640).

3.5

Prolonged Culture

For experimental reasons, it is often necessary to maintain MC in culture for several days or weeks, examining functional features including cell viability and extracellular protein synthesis. MC synthesize and release in culture an extracellular matrix whose composition resembles the in vivo intercellular substance and also the GBM. Its spatial organization is obviously randomly distributed (18, 21). MC matrix regulates cell growth and function by binding growth factors and matrix receptors on the cells themselves (18, 21, 33). This applies to cultured MC

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as well, because their rate of proliferation slows down whenever matrix synthesis is enhanced. This is the case of culture in high glucose media (see Note 10), often used to reproduce the diabetic microenvironment (34, 35). Indeed, long-term culture of MC has an impact on matrix synthesis and release even when ordinary media are employed. Schnaper et al. have shown that later passages (> passage 11) of human fetal MC in culture increase steady-state expression of mRNA for the α1 chains of collagen type III and IV, and laminin β1 and γ1 (36). Conversely, matrix-degrading enzymes such as interstitial collagenase (MMP-1), gelatinase A (MMP-2), tissue-type plasminogen activator (tPA), tissue inhibitor of metalloproteinases (TIMP-1), and plasminogen activator inhibitor (PAI-1) tend to decrease or disappear with time in culture. As a result, collagen type IV accumulation describes a matrix-accumulating phenotype in later passages of cultured MC, likely resulting from progressively higher levels of transforming growth factor (TGF)-β1 (36). Further evidence of a regulatory role of matrix on mesangial proliferation in culture comes from the concept of 3D cultures (37, 38). This approach, first introduced by Yaoita and Marx, uses a collagen type I gel to grow cells embedded within the extracellular matrix. In this microenvironment the cells are quiescent, as demonstrated by various functional assays. This technique more closely resembles the in vivo situation of MC, surrounded by GBM, other types of cells, and their own extracellular matrix (37–39). One drawback of this experimental setup is the low density of the cells, which complicates biochemical assays and messenger RNA (mRNA) extraction. Difficult diffusion of nutrients and test substances may also occur, due to gelification of the substrate. Savill and coworkers emphasized the absolute requirement of MC for growth factors to support not only proliferation but viability as well (40–42). Indeed, when studied after omitting serum from the culture medium, rat MC exhibit higher rates of apoptosis, consistent with growth factors acting as “survival factors.” Thus, working on serum-starved cells may provide potentially misleading results, due to MC entering a programmed cell death mode (40–42). Over the years, a number of laboratories have developed MC lines stabilized by transfection with simian virus (SV) 40 or isolated from SV-40 transgenic mice (43, 44). There is no evidence that this approach is necessary to stabilize murine cell lines or that it can prevent dedifferentiation in vitro. Since the same degree of stability can be obtained by limiting the number of passages, thawing and expanding frozen batches of the same cell preparation can yield the same results, without the complications and artifacts of viral transformation. The phenotype of cultured MC is also affected by physical forces, much like is believed to occur in vivo during glomerular hypertension or hyperfiltration, such as in remnant kidneys or in diabetes. At least three laboratories have grown MC under conditions of controlled pressure resembling the in vivo intraglomerular transcapillary pressure of 40–60 mmHg. Interestingly, increasing pressure resulted in enhanced matrix synthesis (45–47). In an effort to minimize variability in growth rates, phenotype, and biochemical responses among different cell lines, various laboratories have introduced

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“defined” media, consisting of formulations of growth factors in controlled amounts, limiting the amount of added protein. Defined media are more expensive than supplements of raw, decomplemented FBS, and have not gained wide popularity. Indeed, an acceptable degree of stability and reproducibility can be obtained by pooling different batches of FBS and using identical amounts (typically 10–17–20%) after careful storage at –70°C. UV exposure of complete media and serum as well as repeated freezing-thawing cycles should be avoided. Preformed combinations of insulin, transferrin, and selenium salt have been popular, although no obvious advantages over conventional media have been convincingly shown.

3.6

Long-term Storage of Cultured MC Lines

Storing MC lines frozen after isolation, characterization, and clonal expansion is highly recommended, in order to minimize interassay variability. Experiments can be done on multiple batches of cells kept in culture for a short time, instead of using repeatedly subcultured populations in which dedifferentiation and chromosomal abnormalities commonly occur with time. In our opinion, frozen stocks of cells should be preferred to prolonged cultures and late passages. Pure cultures of rat or human MC can be obtained by several sources such as cell banks or even purchased. Table 1.3 lists some of the major organizations providing established cell lines of smooth muscle or MC origin. No matter what the source of cultured MC, the freezing/thawing procedures are fairly standardized.

3.7

Conclusions

About 30 years and a few thousands of papers from laboratories worldwide since the first methodological reports, the art and science of growing MC in renal research is still a matter of debate. In the age of molecular biology, genomics and proteomics, the scarcely differentiated phenotype of MC attracts undoubtedly less attention than the more fashionable podocyte. MC “are out, podocytes are in” nowadays,

Table 1.3 Sources of commercially available smooth muscle cell/MC cultures American Type Culture Collection (ATCC)—LGC Promochem: http://www.lgcpromochem-atcc. com/ Invitrogen: http://www.invitrogen.com/ European Collection of Cell Cultures (ECACC): http://www.ecacc.org.uk/ These organizations are three of the major providers of cell lines used in tissue culture work worldwide. Several other cell banks and commercial companies offer smooth muscle cells of various origin and even mesangial cells. Inquiries should be placed directly for availability and characteristics of specific cell lines

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as suggested by a recent provocative review (8). The vast number of slit diaphragm and intrinsic podocyte proteins and their relationship with the nephrotic syndrome account for the booming interest in this peculiar cell type (6–8, 48). Yet, much has still to be learned about the role of MC in health and disease, with particular emphasis on the interaction with podocytes and endothelial cells. While it is likely that the next few years will see a shift in the focus of research from cultured cells to experimental models and human disease, MC should still remain a powerful tool for genetic manipulation, drug development, and testing (49). Time will tell whether there will be room for MC culture as a gene therapy and bioengineering tool, such as we are now seeing for artificial skin cultures from dermal fibroblasts or cartilage and bone from chondrocytes or osteoblasts.

4

Notes

1. Using young animals is critical to the success of the procedure, because cells from adult kidneys grow slowly and tend to last less in culture. 2. Kidneys from young individuals, children, or even fetal tissue are more likely to yield successful cultures than samples from mature subjects. For human tissue, a thorough microscopic evaluation should be carried out on a fragment excised from apparently healthy cortex (that is, distant from the lesion that led to nephrectomy). This can be done in parallel while performing glomerular isolation; evidence of neoplastic infiltration, necrosis or inflammatory reaction/leukocyte infiltration should halt the cell culture procedure. 3. Cutting the final 5 mm of the plastic tip with a sterile scalpel or razor blade helps picking up larger fragments. 4. FBS is the most likely source of bacterial or mycoplasma contamination. Whenever unexplained contamination of complete culture media occurs, it is advisable to run control incubations at 37°C of 4–5 mL of media in sterile Petri dishes with and without FBS. Should outgrowth of microorganisms occur over 36–72 h, the batch of FBS is returned to the producer or vacuum-filtered over Millipore filters. Foaming of serum and adhesion of growth factors to Millipore membranes is a frequent complication of this procedure, which is not advisable. 5. It is not advisable to disturb the initial attachment by removing often the culture dishes from the incubator. Dishes should be left standing for at least 24 h. 6. MC usually firmly adhere to the plastic growth substrates, although certain lines or clones tend to detach easily as a single sheet when challenged with vasoconstrictors or manipulated for biochemical experiments. Fibronectin or collagen coating of dishes is not necessary under most circumstances, at variance with epithelial or endothelial cells. In our experience, detachment occurs mostly when cells are superconfluent, and likely there is synergy of action between the mechanical forces through specialized tight junctions establishing electrical continuity of the monolayer (16). Synchronous depolarization occurs when a contractile stimulus or shear stress is applied, thus pulling the monolayer off as a single sheet of cells. A simple solution is to schedule studies on cells that are still slightly subconfluent (that is, while individual cells can still be recognized). 7. The distribution of certain markers may be fairly uneven across the population studied, occasionally within the same cell line and passage in culture. This may be due to clonal selection (see above) and the asynchronous position of cells within their growth cycle. 8. In our experience, fully functional cells can be recovered after 10 years of storage, provided that no freezing/thawing has ever occurred. Viability is up to 90%, as assessed by trypan blue or acridine orange dye exclusion. Storage of the tanks (cell repository and reservoir) and all handling are best done in a –4°C or –20°C walking refrigerator, in order to minimize cold loss. Protective clothing (hands, eyewear) is advisable.

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9. Polystyrene/cotton wrapping (approx. 5 cm thickness per individual vial) slows initial cooling in the -80°C freezer, where vials should be kept for 24 h prior to transferring to the liquid N2 tank. This minimizes damage due to icing within the cytosol and organelles. 10. Glucose concentration is usually set at 30 mM versus control cultures at 5.5 mM; a third parallel culture should be employed to control for the effects of hyperosmolar growth conditions, using mannitol or sorbitol as an inert, nonmetabolized sugar.

References 1. Latta, H. (1973) Ultrastructure of the glomerulus and juxtaglomerular apparatus in, Handbook of Physiology (Orloff J., Berliner R.W., Geiger S.R., eds.). American Physiological Society, Washington DC, pp. 1–29. 2. Marinozzi, V. (1961) Struttura e istofisiologia del glomerulo, in Atti del II Corso di aggiornamento professionale “Nefrologia Moderna”, Rome, Italy, pp. 33–51. 3. Zimmermann, K.W. (1929) Über der Bau des Glomerulus der Menschlichen Niere. Z. Mikrosk. Anat. Forsch. 18, 520–552. 4. Latta, H. (1992) An approach to the structure and function of the glomerular mesangium. J. Am. Soc. Nephrol. 2 (suppl 10), S65–S73. 5. Johnson, R.J., Floege, J., Yoshimura, A., Iida, H., Couser, W.G., Alpers, C.E. (1992) The activated mesangial cell: a glomerular “myofibroblast”? J. Am. Soc. Nephrol. 1992; 2 (suppl 10), S190–S197. 6. Pavenstadt, H., Kriz, W., Kretzler, Μ. (2003) Cell biology of the glomerular podocyte. Physiol. Rev. 83, 253–307. 7. Johnstone, D.B., Holzman, L.B. (2006) Clinical impact of research on the podocyte slit diaphragm. Nature Clin. Pract. Nephrol. 2, 272–282. 8. Jefferson, J.A., Shankland, S.J. (2006) Glomerular disease: the podocyte is ready for prime time and may be already center stage. NephSAP 5, 331–338. 9. Kriz, W., Elger, Μ., Mundel, P., Lemley, K.V. (1995) Structure-stabilizing forces in the glomerular tuft. J. Am.Soc. Nephrol. 5, 1731–1739. 10. Baker, A.J., Mooney, A., Hughes, J., Lombardi, D., Johnson, R.J., Savill, J. (1994) Mesangial cell apoptosis: the major mechanism for resolution of glomerular hypercellularity in experimental mesangial proliferative nephritis. J. Clin. Invest. 94, 2105–2116. 11. Quadracci, L., Striker, G.E. (1970) Growth and maintenance of glomerular cells in vitro. Proc. Soc. Exp. Biol. Med. 135, 947–950. 12. Burlington, H., Cronkite, E.P. (1973) Characteristics of cell cultures derived from renal glomeruli. Proc. Soc. Exp. Biol. Med. 142, 143–149. 13. Kreisberg, J.I., Hoover, R.L., Karnovsky, Μ.J. (1978) Isolation and characterization of rat glomerular epithelial cells in vitro. Kidney Int. 14, 21–30. 14. Striker, G.E., Striker, L.J. (1985) Biology of disease. Glomerular cell culture. Lab. Invest. 53, 122–131. 15. Davies, Μ. (1994) The mesangial cell: a tissue culture view. Kidney Int. 45, 320–327. 16. Ennulat, D., Brown, S.A. (1995) Canine and equine mesangial cells in vitro. In Vitro Cell. Dev. Biol. Anim. 31, 574–578. 17. Ijima, K., Moore, L.C., Goligorsky, Μ.S. (1991) Syncytial organization of cultured rat mesangial cells. Am. J. Physiol. 260, F848–F855. 18. Menè, P., Simonson, Μ.S., Dunn, Μ.J. (1989) Physiology of the mesangial cell. Physiol. Rev. 69, 1347–1424. 19. Menè, P. (2001) Mesangial cell cultures. J. Nephrol. 14, 198–203. 20. Pabst, R., Sterzel, R.B. (1983) Cell renewal of glomerular cell types in normal rats. An autoradiographic analysis. Kidney Int. 24, 626–631. 21. Rupprecht, H.D., Sterzel, R.B. (1997) Glomerular mesangial cells, in Immunologic renal diseases (Neilson E.G., Couser W.G., eds.). Lippincott-Raven, Philadelphia, pp. 595–626.

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22. Striker, G.E., Killen, P.D., Farin, F.Μ. (1980) Human glomerular cells in vitro: isolation and characterization. Transplant Proc. 12 (Suppl 1), 88–99. 23. Menè, P., Dunn, Μ.J. (1986) Contractile effects of TxA2. and endoperoxide analogues on cultured rat glomerular mesangial cells Am. J. Physiol. 251 (Renal Fluid Electrolyte Physiol. 20): F1029–F1035. 24. Menè, P., Dubyak, G.R., Abboud, H.E., Scarpa, A., Dunn, Μ.J. (1988) Phospholipase C activation by prostaglandins and thromboxane A2 in cultured mesangial cells. Am. J. Physiol. 255 (Renal, Fluid and Electrolyte Physiol. 24): F1059–F1069. 25. Men è, P., Pugliese, F., Faraggiana, T., Cinotti, G.A. (1990) Identification and characteristics of a Na-. /Ca2+ exchanger in cultured human mesangial cells Kidney Int. 38, 1199–1205. 26. Sterzel, R.B., Lovett, D.H., Foellmer, H.G., Perfetto, Μ., Biemesderfer, D., Kashgarian, Μ. (1986) Mesangial cell hillocks. Nodular foci of exaggerated growth of cells and matrix in prolonged culture. Am. J. Pathol. 125, 130–140. 27. Wilson, H.Μ., Stewart, K.N. (2005) Glomerular epithelial and mesangial cell culture and characterization. Methods Mol. Med. 107, 269–282. 28. Ardaillou, R. (1996) Biology of glomerular cells in culture. Cell Biol. Toxicol. 12, 257–261. 29. Menè, P., Fofi, C., Domenici, A., Stoppacciaro, A. (2005) Immunophenotyping glomerular cells in renal biopsies, a novel approach to the diagnosis of kidney diseases. J. Am. Soc. Nephrol. 16, 816A. 30. Elger, Μ., Drenckahn, D., Nobiling, R., Mundel, P., Kriz, W. (1993) Cultured rat mesangial cells contain smooth muscle alpha-actin not found in vivo. Am. J. Pathol. 142, 497–509. 31. Schreiner, G.F., Unanue, E.R. (1984) Origin of the rat mesangial phagocyte and its expression of the leukocyte common antigen. Lab. Invest. 51, 515–523. 32. Gieseler, R., Hoffmann, P.R., Kuhn, R., Fayyazi, A., Stojanovic, T., Schlemminger, R., Peters, J.H. (1997) Enrichment and characterization of dendritic cells from rat renal mesangium. Scand. J. Immunol. 46, 587–596. 33. Ballermann, B.J. (1989) Regulation of bovine glomerular endothelial cell growth in vitro. Am. J. Physiol. 256 (Cell Physiol. 25), C182–189. 34. Pricci, F., Pugliese, G., Menè, P., Romeo, G., Romano, G., Galli, G., Casini, A., Rotella, C.Μ., Di Mario, U., Pugliese, F. (1996) Regulatory role of eicosanoids in extracellular matrix overproduction induced by long-term exposure to high glucose in cultured rat mesangial cells. Diabetologia 39, 1055–1062. 35. Menè, P., Pugliese, G., Pricci, F., Di Mario, U., Cinotti, G.A., Pugliese, F. (1997) High glucose inhibits capacitative Ca2+. influx in cultured rat mesangial cells by a protein kinase C-dependent mechanism. Diabetologia 40, 521–527. 36. Schnaper, H.W., Kopp, J.B., Poncelet, A.C., Hubchak, S.C., Stetler-Stevenson, W.G., Klotman, P.E., Kleinman, H.K. (1996) Increased expression of extracellular matrix proteins and decreased expression of matrix proteases after serial passage of glomerular mesangial cells. J. Cell. Sci. 109:2521–2528. 37. Yaoita, E. (1989) Behavior of rat mesangial cells cultured within extracellular matrix. Lab. Invest. 61, 410–418. 38. Marx, Μ., Daniel, T.O., Kashgarian, Μ., Madri, J.A. (1993) Spatial organization of the extracellular matrix modulates the expression of PDGF-receptor subunits in mesangial cells. Kidney Int. 43, 1027–1041. 39. Floege, J., Radeke, H.R., Johnson, R.J. (1994) Glomerular cells in vitro versus the glomerulus in vivo. Kidney Int. 45, 360–368. 40. Marx, Μ., Dorsch, O. (1997) pp60c-src is required for the induction of a quiescent mesangial cell phenotype. Kidney Int. 51, 110–118. 41. Mooney, A., Jobson, T., Bacon, R., Kitamura, Μ., Savill, J. (1997) Cytokines promote glomerular mesangial cell survival in vitro by stimulus-dependent inhibition of apoptosis. J. Immunol. 159, 3949–3960. 42. Mooney, A., Jackson, K., Bacon, R., Streuli, C., Edwards, G., Bassuk, J., Savill, J. (1999) Type IV collagen and laminin regulate glomerular mesangial cell susceptibility to apoptosis via beta(1) integrin-mediated survival signals. Am. J. Pathol. 155, 599–606.

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43. Vicart, P., Schwartz, B., Vandewalle, A., Bens, Μ., Delouis, C., Panthier, J.J., Pournin, S., Babinet, C., Paulin, D. (1994) Immortalization of multiple cell types from transgenic mice using a transgene containing the vimentin promoter and a conditional oncogene. Exp. Cell. Res. 214, 35–45. 44. Sraer, J.D., Delarue, F., Hagege, J., Feunteun, J., Pinet, F., Nguyen, G., Rondeau, E. (1996) Stable cell lines of T-SV40 immortalized human glomerular mesangial cells. Kidney Int. 49, 267–270. 45. Mattana, J., Singhal, P.C. (1995) Applied pressure modulates mesangial cell proliferation and matrix synthesis. Am. J. Hypertens. 8:1112–1120. 46. Singhal, P.C., Sagar, S., Garg, P. (1996) Simulated glomerular pressure modulates mesangial cell 72 kDa metalloproteinase activity. Connect. Tissue Res. 33, 257–263. 47. Mertens, P.R., Espenkott, V., Venjakob, B., Heintz, B., Handt, S., Sieberth, H.G. (1998) Pressure oscillation regulates human mesangial cell growth and collagen synthesis. Hypertension 32, 945–952. 48. Mundel, P., Reiser, J., Kriz, W. (1997) Induction of differentiation in cultured rat and human podocytes. J. Am. Soc. Nephrol. 8, 697–705. 49. Rodriguez-Barbero, A., L’Azou, B., Cambar, J., Lopez-Novoa, J.Μ. (2000) Potential use of isolated glomeruli and cultured mesangial cells as in vitro models to assess nephrotoxicity. Cell. Biol. Toxicol. 16, 145–153.

Chapter 2

Isolation and Primary Culture of Human Proximal Tubule Cells David A.Vesey, Weier Qi, Xinming Chen, Carol A. Pollock, and David W. Johnson

Abstract Primary cultures of renal proximal tubule cells (PTC) have been widely used to investigate tubule cell function. They provide a model system where confounding influences of renal haemodynamics, cell heterogeneity, and neural activity are eliminated. Additionally they are likely to more closely resemble PTC in vivo than established kidney cell lines, which are often virally immortalised and are of uncertain origin. This chapter describes a method used in our laboratories to isolate and culture pure populations of human PTC. The cortex is dissected away from the medulla and minced finely. Following collagenase digestion, the cells are passed through a sieve and separated on a Percoll density gradient. An almost pure population of tubule fragments form a band at the base of the gradient. Cultured in a hormonally defined serum-free growth media, they form a tightly packed monolayer that retains the differentiated characteristics of PTC for up to three passages. Keywords Renal proximal tubule cells, Hormonally defined serum free medium, Percoll gradient, Polarised monolayers

1

Introduction

The kidney is a complex organ composed of at least 12 functionally distinct epithelial cell types (1). The proximal tubule cells (PTC), which, along with tubular vasculature, form greater than 80% of the renal cortex, are one of the most prominent epithelial cell types. Not only do they play important roles in fluid, amino acid and sodium reabsorption, they also contribute significantly to pathological changes within the cortical tubulointerstitium (2, 3). In order to study the function of PTC in the absence of confounding influences of other renal cell types, haemodynamics, and neural activity, methods have been developed to isolate and culture these cells. Established cell lines such as the opossum kidney-derived cell line, OK; the porcine tubular epithelial cell line, LLC-PK1; and the human tubular cell line, HK2, exhibit degrees of de-differentiation and loss of PTC-specific biochemical and transport properties (4–7). From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_2, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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In this chapter we describe a method used in our laboratories for the isolation and culture of human PTC, based on the method of Vinay et al. (8). The cortical tissue is dissected away from the kidney section, minced finely, and digested with collagenase. Following filtration to remove undigested and fibrous tissue, cells are separated on a Percoll gradient. Proximal tubule fragments form a band near the base of the gradient. Using this method, a highly enriched population of proximal tubular fragments and cells can be obtained. Yields are typically in the order of 2 million cells per gram of original cortical tissue. Cultured in a hormonally defined serum-free media, they form confluent polarised monolayers in 5–8 days with a characteristic cobblestone appearance. Dome formation is apparent when the monolayers reach higher confluency. For experimentation, we use these cells at passage 2 when growth and differentiation characteristics are well preserved (9).

2

Materials

1. Water: water for preparation of sterile media and buffers is obtained from a Milli-Q TM water system (Millipore, Billerica, MA, USA). The feed water for this is from a reverse osmosis system. 2. Krebs–Henseleit Solution (KHS): 108 mM NaCl, 4.9 mM KCl, 2.6 mM CaCl2, 3.1 mM NaH2PO4, and 28 mM NaHCO4, adjusted to pH 7.4 and filtered sterilised with a 0.2-µm filter (see Notes 1 and 2). 3. Percoll solution (GE Health Care, Sydney, Australia). Diluted to 45% with double strength KHS before use. 4. Collagenase solution: 30 mg collagenase (Type II, 300–400 U/mg; Worthington Biochemical Company) is dissolved in 30 mL of KHS and filter sterilised with a 0.2-µm filter (see Note 3). 5. Sieve: stainless-steel tissue dissociation sieve with a mesh size of 297 µm (50mesh screen). Heat sterilised before use. 6. Centrifuge tubes: 50-mL sterile polypropylene centrifuge tubes are used throughout the procedure for washing cells. Pre-sterilised 50-mL polycarbonate centrifuge tubes are used for Percoll centrifugation. 7. Cell culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM)/nutrient mixture Ham’s F-12 (DMEM/F-12) supplemented with sodium bicarbonate (Thermo Scientific, Melbourne, Australia). 8. Penicillin/streptomycin: a stock solution of penicillin/streptomycin (5,000 U/mL/5,000 µg/mL) is obtained from Sigma (Sydney, Australia) and is added to culture media at 5 mL per litre. 9. Defined medium supplements: insulin–transferrin–selenium supplement (Cambrex, Melbourne, Australia) is used at 1 mL per litre. The final concentration of insulin, transferrin, and selenium are 5 µg/mL, 5 µg/mL, and 5 ng/mL, respectively. 10 ng/mL epidermal growth factor, 50 nM hydrocortisone, 5 pM triiodothyronine, and 50 µM prostaglandin E1, all from Sigma, are made up as

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stock solutions and stored at –80°C before addition to the medium at the final concentrations indicated.

3 3.1

Methods PTC Isolation

All procedures were performed using aseptic techniques. Segments of macroscopically and histologically normal renal cortex (5–10 g) were obtained aseptically from the normal pole of adult human kidneys removed surgically because of small (15,000×g for 10 min at 4°C to pellet RNA (see Note 10). The supernatant is decanted or removed with a clean pipette tip, carefully, so as to not loosen the RNA pellet. When most of the supernatant has been removed, 800 µL of 70% ethanol is gently added per tube and the tubes left at room temperature for 5–10 min to allow the diffusion of salts out of the RNA pellet. The samples are then centrifuged again but this time for 5 min at 7,000×g to avoid compacting the pellet and making it difficult to redissolve. All traces of the supernatant are removed and the RNA allowed to air dry for 3–5 min. Over drying should be avoided as this makes it difficult to redissolve the RNA (see Note 11). 3. When dissolving the RNA, there are essentially two options. Dissolving the RNA in formamide is the best option for long-term storage. The RNA dissolves well, and can be quantitated and used in most downstream applications, except where the proportion of RNA volume to reaction volume is appreciable (>5%). In this case the RNA is precipitated and redissolved in sterile water (see Note 12). Typically we would dissolve tissue sample RNA in a volume of between 2 and 5 µL/mg tissue.

3.4

Tissue Culture Cells

3.4.1

Cell Culture Experiments

Tissue culture cells are treated as per standard protocols dictated by each experiment. The NRK52E rat proximal tubular cells are seeded in 1× DMEM supplemented with 10% serum and antibiotics. The next day the medium is replaced with DMEM containing 2% serum with the treatment, for example 10 ng/mL transforming growth factor (TGF)-β1. At the desired time point, cells are harvested as described below. Typically from a confluent monolayer of cells we would obtain 8–16 µg RNA per well of a 12-well plate, or 15–40 µg per well of a 6-well plate.

3.4.2

Harvesting Cells

1. Tissue culture medium is removed from 6- or 12-well plates by vacuum and the cell monolayers washed once with ice-cold PBS. 2. Cells are lysed by the addition of 0.8 mL TrizolTM directly to each well as the plates are sitting on ice. An alternative protocol is to use the Solution D method (3) (see Note 13).

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3. An optional step at this point is to scrape the cells. In our experience this may marginally increase the yield of RNA. Smaller volumes of TrizolTM can be used but will affect RNA purity and yield (see Note 3). 4. The lysed cells in TrizolTM are aspirated from each well using a 3-mL syringe with a 21-gauge needle and transferred to clean 1.5-mL microfuge tubes. The gDNA is sheared by passing the lysate through the syringe three to five times. This step is important as it affects the viscosity of the sample. At the end of each group of treatments a new syringe is used. All samples are kept on ice till the next step. 5. Samples can be stored at −20°C at this stage for up to 3–4 weeks without compromising RNA quality.

3.4.3

RNA Extraction from Cultured Cells

1. The method is similar to that described above for extraction of RNA from tissue homogenates with some modifications. The entire process is carried out in microfuge tubes. 2. To isolate RNA, 160 µL chloroform/isoamyl alcohol is added to each tube containing cell lysate, the samples are vortexed for 15 s and left on ice 5 min. Samples are then centrifuged at >14,000×g for 7 min at room temperature and the upper aqueous phase carefully removed to new tubes (see Note 8). 3. To precipitate RNA, 480 µL isopropanol is added to each tube, the tubes briefly vortexed, and left at –20°C overnight (see Note 9). Samples are then centrifuged at >14,000×g for 10 min at 4°C (see Note 10) to pellet the RNA precipitate. Very carefully decant the supernatant or remove it with a clean pipette tip. The RNA pellet from a well of a 12-well plate is very small and easy to lose. When most of the supernatant has been removed, 800 µL of 70% ethanol is gently added per tube and the tubes left at room temperature for 5 min to allow the diffusion of salts out of the RNA pellet. 4. The samples are then centrifuged again but this time for 5 min at 7,000×g to avoid compacting the pellet and making it difficult to redissolve. All traces of the supernatant are removed and the RNA allowed to air dry for 3–5 min (see Note 11). 5. The volume to redissolve the RNA depends on downstream applications. Typically we dissolve RNA from tissue culture cells at 10 µL/well of a 12-well plate, or 25 µL/well of a 6-well plate.

3.5

Quantitation of RNA

1. For quantitation of RNA the protocols are fairly standard, with access to a quartz cuvettes and a spectrophotometer required. A dilution of RNA is made in water depending on the cuvette volume (see Note 14) and the absorbance 260 and 280 nm is measured to establish concentration and purity of RNA (see Note 15).

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2. The concentration of RNA in the sample is calculated based on the formula that an absorbance of 1 at 260 nm is equivalent to 40 µg RNA when measured in water (see Note 16).

3.6

Storage of RNA

All RNA samples are stored at –80°C. At this temperature, samples are relatively stable for a number of years for use with QRT-PCR (see Note 17).

3.7

DNase Treatment of RNA Samples

All current methods of RNA purification result in preparations that are contaminated with gDNA to a varying extent (see Note 18). To remove gDNA the RNA is digested with RNase-free DNase. An optional step would involve the addition of an RNase inhibitor to the sample during the DNA digestion. 1. Typically, 6 µg of RNA is incubated for 20–30 min at 37°C with 1 µL of RNasefree DNase-I (see Note 19) in 1× reaction buffer in a volume of 10 µL as per the DNA-free kit from Ambion. 2. Inactivation reagent (2 µL) is added to each reaction and the samples are mixed and left at room temperature for 2–3 min with occasional mixing (see Note 20). 3. Samples are centrifuged at >14,000×g for 2 min at RT to pellet the inactivation beads and the sample is removed to fresh, labelled tubes for storage at – 80°C (see Note 21).

3.8 cDNA Synthesis Step A standard cDNA synthesis reaction is carried out as follows. The reagents can be purchased from a single manufacturer as a kit or as individual components. We have opted for the later. 1. Mix and briefly centrifuge each component before use. 2. Make up the following Master Mix: DNase treated RNA (see Note 22) Random hexamers (50 ng/µL) Water Final volume

1.8 µL 2.0 µL 6.2 µL 10 µL

3. Mix and centrifuge briefly (pulse) to pellet droplets.

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4. Incubate samples at 70°C for 5 min followed by 1 min on ice (see Note 23). 5. The following reaction mix is prepared, adding each component in the indicated order. The volumes shown are per cDNA reaction (see Note 24): 5× RT Buffer 10 mM dNTP mix 0.1 M DTT Super RNase-InTM(20 U/µL) MMulV (200 U/µL) Water Final mix volume

4 µL 2 µL 2 µL 0.1 µL 1 µL 0.9 µL 10 µL

6. Add 10 µL of reaction mix (step 5 above) to each RNA/primer mix from step 4 above. 7. Gently mix by tapping at the bottom of the tube and then centrifuge briefly (pulse). Incubate at room temp (25°C) for 10 min. 8. Transfer the tubes to 37°C and incubate for 50–60 min. 9. Terminate the reactions by incubating 70°C for 10 min and finally store cDNA at –20°C.

3.9 Quantitative Real-Time PCR 3.9.1

Background

QRT-PCR measures the degradation of a fluorescent-labelled oligonucleotide (referred to as a probe) in real time, concomitant with PCR amplification (5). The probe has a reporter dye at the 5′ end and a quencher dye at the 3′ end (5), and is designed to anneal between the 5′ and 3′ oligonucleotides sites. When the probe first anneals it is intact and thus no fluorescence is emitted. However, during PCR, the probe is cleaved by the 5′ nucleolytic activity of AmpliTaq GoldTM DNA polymerase (ABI), resulting in the separation of the reporter and quencher dyes and subsequently resulting in an energy transfer from the quencher dye to the reporter dye. This process is repeated in every cycle, resulting in the increase in fluorescence, which is measured by the machine. Thus, the accumulation of PCR products is directly monitored by measuring the increase in fluorescence (Fig. 7.1). In the past a passive reference dye (TAMRA) was included in the reaction mixture and acted as an internal control (normalising the reporter dye signal) and correcting for any fluctuations in fluorescence caused by changes in volume or concentration. ABI now produces MGB (Μ, minor binding groove) probes that contain a non-fluorescent quencher (Q) at the 3′ end instead of TAMRA. The major advantage of these MGB probes is that it allows for a more accurate measure of reporter fluorescence (R). A second advantage of MGB probes is that they have a higher melting temperature (Tm), enabling the design and use of shorter probes.

7 Quantitative Gene Expression Analysis in Kidney Tissues Fig. 7.1 Schematic representation summarising the quantitative RT-PCR cycle (see Color Plate 5)

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Polymerization Forward Primer

R

M Q

5⬘ 3⬘ 5⬘

5⬘ 3⬘ 5⬘

Reverse Primer

Strand Displacement R M Q

5⬘ 3⬘

3⬘ 5⬘

Cleavage R M Q 5⬘ 3⬘

3⬘ 5⬘

Polymerization Completed R

M Q

5⬘ 3⬘

3⬘ 5⬘

5⬘ 3⬘

3⬘ 5⬘

In our laboratory we have standardised on the ABI 7500 Fast PCR platform for a number of reasons. These reasons include the “fast” nature of the amplification reaction (40 min) due to thinner plastic ware and PCR block enhancements, and also the reduced reaction volume, which saves on consumable costs. The work described below was carried out on this platform.

3.9.2

Probe and Primer Design

Probes and primers are selected from cDNA sequences such that intronic sequences are avoided. Where possible, primers are designed to span one or more introns so that only cDNA sequences are amplified during the PCR reaction. Probe/primer design is facilitated with the use of Primer Express software that is part of the Applied Biosystems package. The software automatically sets parameters to favour a standard set of reaction conditions for multiplexing. Briefly:

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1. Probe/primer amplicons are designed to be no longer than 100 bp. 2. Probes are designed such that they do not contain runs of more than three consecutive Gs, contain no Gs at the 5′ end, they are selected from the DNA strand with more Cs than Gs, and the Tm must be between 68 and 70°C. 3. Primers are designed with no runs of more than three consecutive Gs, no more than two GCs in the last five nucleotides at the 3′ end, and the Tm must be between 58 and 60°C. 4. Once potential probe and primer sequences are identified, the resulting amplicon is BLASTed against sequence databases to determine the specificity to the gene of interest (see Note 25) before ordering. We purchase all our probes from ABI. 5. Primers and probes should be made up to a stock solution and working solutions aliquoted and stored at – 20°C (see Note 26).

3.9.3

PCR Reaction Setup

RT-PCR reactions contain 500 nM of forward and reverse primers, 50 nM each of FAM/MGB cDNA probe and VICTM/MGB 18S ribosomal probe, in 1× Taqman Fast Universal Master mix (ABI). Each sample is run and analyzed in triplicate. 1. A master mix consisting of all the reaction components is made up for each probe. Below is the recipe used in our laboratory and includes the 18S probe as our internal endogenous control. Water makes up the remainder of the volume: 18S probe (ABI kit) Specific probe (1 µM stock) Forward primer (10 µM stock) Reverse primer (10 µM stock) TaqMan Fast Universal Master Mix Water Final volume

0.35 µL 0.625 µL 0.625 µL 0.625 µL 6.25 µL 4.025 µL 12.5 µL

2. 12-µL aliquots of the above master mix are added to each well of the PCR plate depending on the number of cDNA samples to be analysed. 3. Finally 0.5 µL (up to 1 µL) of cDNA prepared in Sect. 3.8 above is added to the corresponding wells of the PCR plate. 4. The plate is sealed with adhesive optical covers that are compatible with the ABI 7500 Fast PCR machine (ABI) and inserted into the PCR block for the run to begin. 5. The real-time PCR machine is set according to the dyes being used and this setup should never be altered unless different dyes are used with different probes. We have standardised on the dyes mentioned above to avoid changing the setup on the machine in normal use (see Note 27). 6. When the run has completed, the plate is removed and data is saved for analysis as described below.

7 Quantitative Gene Expression Analysis in Kidney Tissues

3.9.4

95

Quantitative Real-Time PCR

The amplification plots generated by the PCR machine are analysed using the ABI 7500 Fast Real-Time PCR Sequence Detection Software. The amplification of the test gene is compared against the endogenous control. An example of the data generated is shown for kidney RNA in Fig. 7.2. The amplification curves of a gene that is in high abundance, 18S in this example, appear at an earlier cycle number indicated by the curves on left side of the plot. In contrast the curves on the right side are those of angiotensin-converting enzyme (ACE), whose expression in these samples is lower than 18S and hence the curves appear at a higher cycle number. 1. To analyse the amplification curve of a sample, the threshold line (arrow heads on right, Fig. 7.3) is positioned in the linear phase of the exponential curve of PCR amplification. The threshold line can also be set automatically by the software. 2. This threshold line is set above background fluorescence and the fluorescence emitted by the no-template controls (NTC) (see Note 28). 3. Where the fluorescence of a particular sample rises significantly above background and crosses the threshold, a cycle number (Ct) is then recorded (Fig. 7.3). Therefore, if a given sample contains more cDNA for the gene of interest it will result in an earlier emission of fluorescence rising above the threshold and thus recording a lower Ct.

Fig. 7.2 A screen capture of an amplification plot for a typical quantitative real-time PCR (QRTPCR) experiment using the ABI 7500 FAST machine. The amplification curves are typical of multiplexing reactions. In this case the curves to the left are the endogenous 18S gene, while those on the right are for ACE gene expression

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Fig. 7.3 Amplification curves for a FAM- and a VICTM-labelled probe. A threshold line is set in the linear phase of the curve and a cycle number is derived from where the linear curve crosses the threshold line

3.9.5

Optimisation of Probe and Primer Concentrations

For results to be valid, optimisation of each probe and primer combination needs to be conducted prior to their use in the quantitation of gene expression levels in cDNA samples. 1. Probes and primers are tested at the standard concentrations of 50 nM and 500 nM respectively, as recommended by ABI. 2. Optimisation of probe and primer concentrations is conducted by increasing the concentration of probes and primers by 1.2-, 1.5-, and 2-fold in an optimisation experiment. 3. If no increase in Ct value is observed at the standard concentrations, these concentrations are accepted as appropriate for the system and samples employed in the study. However, changes in Ct values imply that the reagents are limiting for the template being used and need to be adjusted. It is important that the reagents be in excess. The standard concentrations are used in our laboratory and have consistently been shown to be adequate for most genes we have studied to date.

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Housekeeping Genes for Normalisation

The debate still rages concerning which housekeeping gene is the best to use to normalise gene expression data for RNA quality and input, and variation in the efficiency of cDNA synthesis (see Note 29). This issue was highlighted in a recent publication by de Kok et al. (6) who analysed 13 different housekeeping genes and identified the HPRT gene as the best for use in tumour tissues. Laboratories with a gene profiling background have often used the mean expression of a combination of genes to normalise their data. The endogenous housekeeping gene used in our laboratory and by many others in the field is 18S. The reasons are that 18S is expressed in all cell types and is highly conserved amongst many different species. The primers and probe for 18S are available commercially (ABI) in a ready-to-use mixture. Optimisation experiments for using 18S in multiplex PCR are critical since the gene is abundantly expressed in all cells. There is potential danger when using an excess 18S probe/ primer mix, which can lead to the exhaustion of the reaction components and thus effecting amplification of the target gene. It is therefore essential that competitive and non-competitive reactions be conducted to establish conditions that avoid this problem (see Sect. 3.9.8). The choice of 18S as the housekeeping gene is often criticized because of the high level of expression. Although 18S is abundantly expressed, unlike GAPDH or β-tubulin, diseases such as diabetes do not affect the expression of this gene. This needs to be established in each laboratory with respect to the housekeeping gene used and the sorts of samples and diseases under investigation. To establish the validity of 18S in our experiments it was crucial to determine whether the expression level of this gene was altered between sample groups. For example, below is the data for rat kidney cDNA where the expression of ACE was investigated. The ACE probe is labelled with a fluorescent FAM dye. The PCR reactions were multiplex reactions with 18S (labelled with fluorescent VIC dye) as the endogenous housekeeping gene. A comparison was made between cDNA samples from control and diabetic rat kidney samples (Table 7.1). Note the 18S values in both groups. The mean 18S cycle number for the control and diabetic kidney 13.28 versus 13.52 respectively, indicating a similar level of expression between the two groups. To confirm that there is no significant difference in the 18S Ct values between the groups, it is recommended that one-way variance analysis be performed. In contrast the difference in the mean ACE expression between the two groups is about 1.8 cycles, which is a significant difference (p = 0.0042). Thus we can conclude that the expression of ACE is reduced in diabetic kidney compared with control.

3.9.7

Multiplexing

Multiplexing allows the amplification and measurement of both the gene of interest and the endogenous control gene in the same reaction. The probe reporter dye is typically labelled with FAM and the endogenous control probe is labelled with VICTM. Each dye emits fluorescence at different wavelengths when excited by the laser.

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Table 7.1 Comparison of 18S–VIC and ACE–FAM cycle number values between control and diabetic rat kidney samples Group Control Control Control Control Control Diabetic Diabetic Diabetic Diabetic Diabetic Diabetic

ACE–FAM 28.66 30.24 29.6 28.4 28.7 30.3 30.3 31.93 31.71 31.02 30.08

18S–VIC 12.44 13.38 14.38 12.93 13.29 13.72 13.25 13.49 13.62 13.18 13.85

Mean ACE–FAM 29.12

Mean 18S–VIC 13.28

30.89

13.52

Kidney samples from six different animals were analysed in each group

Table 7.2 Comparison of ∆Ct values in multiplexing versus non-multiplexing reactions Non-multiplexing Multiplexing

FAM (Ct)

VIC (Ct)

Ct

28 29

11 12

17 17

To ensure reproducibility and to generate accurate data, competitive (multiplexing) and non-competitive (non-multiplexing) reactions must be carried out with the VICTM-labelled endogenous control probe. The Ct values are extrapolated from the amplification plot (Fig. 7.2) for both probes, and the difference in Ct (∆Ct) is calculated by subtracting the VIC Ct from the FAM Ct. When the ∆Ct values for multiplexing versus non-multiplexing are identical (or within 0.8 of a cycle), multiplexing experiments are valid (Table 7.2) because the reactions for each probe do not interfere with one another.

3.9.8

Relative Efficiency

The relative efficiency of PCR amplification is used to demonstrate equal amplification efficiencies of both the FAM-labelled probe and the 18S probe in multiplex reactions over different initial template concentrations. This is carried out by varying the amount of template cDNA in the PCR reaction, determining the ∆Ct values for these reactions, and finally plotting ∆Ct versus total cDNA concentration. A line of best fit is then drawn and the value of the slope should be < 0.1, which demonstrates that varying the initial template amount does not significantly alter the ∆Ct value (Fig. 7.4). This ensures accurate and reproducible results in multiplexing reactions.

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20

∆Ct

17.5

15

12.5

10

100

50

25

12.5

6.25

3.125

cDNA (ng) Fig. 7.4 Relative efficiency plot for ACE amplification. The ∆Ct value is shown for each of the starting concentrations of cDNA

3.9.9

Quantitation Gene Expression by RT-PCR

QRT-PCR is a fully quantitative method for determining cDNA levels of specific genes that correspond to mRNA level of those genes in the original sample. The method used to obtain relative gene expression data is the comparative Ct method previously described by Livak (7). In this method a “calibrator” sample is used as a baseline for comparison of the level of gene expression for every unknown sample. In the case study outlined below the calibrator group is the control group. The calculations first involve the subtraction of the 18S Ct value from the Ct value of the FAM probe resulting in the differential Ct or ∆Ct value. The average ∆Ct value is then calculated for the calibrator group and this is subtracted from the ∆Ct value of every sample, including the ∆Ct values of the calibrator group. The resulting value is the ∆∆Ct, and this value is used in the equation, 2–∆∆Ct to derive the fold induction (FI) value. All means, standard error of means, and statistics are calculated using this value.

3.10

Case Studies

3.10.1

Case Study with RNA from Rat Kidney

A case study is outlined here to demonstrate a practical use for QRT-PCR. RNA was isolated and cDNA was synthesised as described above. The aim was to compare ACE mRNA levels in diabetic rat kidney to control rat kidney. The ACE probe is labelled with FAM fluorescent dye. The 18S probe is used as the endogenous

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Table 7.3 Comparison of the FI reveals that ACE mRNA levels in diabetic rat kidney are significantly reduced (mean FI, 0.4±0.2; p< 0.01) as compared with ACE mRNA levels in control rat kidney (mean FI, 1.09±0.44) Group

ACE–FAM

18S–VIC

∆Ct

∆∆Ct

FI

Mean FI

STDV

SE

Control Control Control Control Control Diabetic Diabetic Diabetic Diabetic Diabetic Diabetic

28.66 30.24 29.6 28.4 28.7 30.3 30.3 31.93 31.71 31.02 30.08

12.44 13.38 14.38 12.93 13.29 13.72 13.25 13.49 13.62 13.18 13.85

16.22 16.86 15.2 15.46 15.43 16.53 17.1 18.44 18.09 17.84 16.23

0.38 1.02 –0.64 –0.38 –0.41 0.69 1.26 2.6 2.25 2 0.39

0.77 0.49 1.56 1.30 1.33 0.62 0.42 0.16 0.21 0.25 0.76

1.09

0.44

0.20

0.40

0.20

0.08

housekeeping gene and is labelled with VIC fluorescent dye. FI in each case is calculated and shown in Table 7.3. ACE expression in diabetic kidney is significantly reduced (0.4±0.2; p