Kidney Research. Experimental Protocols

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

xiii

xiv

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

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

3

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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

8

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

2 Isolation and Primary Culture of Human Proximal Tubule Cells

21

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 (<6 cm) renal cell carcinomas. Patients were otherwise healthy and not on medication. Informed consent was obtained prior to each operative procedure and the use of human renal tissue for primary culture was reviewed and approved by the Princess Alexandra Hospital Research Ethics Committee. 1. Upon removal of the kidney tissue, the tissue is placed in a sterile 70-mL container with ice-cold culture media. It is transferred to the laboratory on ice. 2. In the laboratory culture cabinet, transfer the tissue to a sterile shallow Petri dish. Remove the kidney capsule and dissect away the cortex. The remaining tissue can be discarded. We usually fix a small piece of the cortex in formalin for histology and rapidly freeze another piece for storage in liquid nitrogen. The cortical segments are then rinsed with fresh KHS and finely minced with a sterile razor. The minced tissue is washed three times in 40 mL of cold KHS by centrifugation at 600 × g for 10 min at 4°C. 3. The resultant pellet is resuspended in 30 mL of pre-warmed KHS containing 30 mg of class 2 collagenase (Worthington). This is incubated with stirring or shaking at 37°C for 30 min (see Notes 3 and 4). 4. The digested tissue is diluted 1:2 with ice-cold KHS and filtered through a 297µm sieve (50-mesh screen) to remove any undigested fibrous tissue. 5. The filtrate is centrifuged at 600 × g for 4 min at 4°C. The pellet is then resuspended in fresh cold KHS and re-centrifuged. This is repeated a total of three times (see Note 5). 6. The final pellet is then resuspended in 50 mL of 45% Percoll solution, which is divided between two tubes (see Note 6). 7. Centrifuge the tubes at 20,000 × g for 30 min at 4°C in a fixed angle rotor. This procedure establishes a density gradient in which the cells separate into four bands, F1 to F4. The lowermost band, F4, is enriched with proximal tubule fragments and contains the highest alkaline phosphatase and gamma-glutamyl transpeptidase activities. This band is carefully removed and washed three times in media (see Note 7). 8. Resuspended the cell pellet in serum-free, hormonally defined culture media and place in 25-cm2 culture flasks at a seeding density of 1.5 mg pellet/cm2 (see Notes 8 and 9).

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9. Cells are incubated in a humidified atmosphere of 95% air/5% CO2 at 37°C. Change medium every 2 days. When confluence is reached in 5 to 8 days, the cells can be subcultured with trypsin–EDTA. 10. Cells can be cryopreserved in DMEM/F12 medium containing 20% dimethyl sulphoxide and 10% foetal bovine serum.

Fig. 2.1 Phase-contrast microscopic appearance of human proximal tubule cells (PTC) in culture. Human PTC form a tight “cobblestone-like” monolayer following 8 days culture in serum-free hormonally defined media. Dome formation can be seen throughout the culture. Magnification: (a) × 200, (b) × 100, (c) × 40

2 Isolation and Primary Culture of Human Proximal Tubule Cells

3.2

23

Characteristics of Cultured PTC

1. Confluent cultures of PTC have a typical epithelial cobblestone appearance. Domes or hemicysts form when the cultures become highly confluent. These domes are the result of fluid transport below the cell monolayer, which slightly raises the cells off the culture surface (Fig. 2.1). 2. Using scanning and transmission electron microscopy, the cells can be seen to have a polarised morphology with numerous apical microvilli. The cells have a columnar shape with intercellular tight junctions and abundant mitochondria (9, 10). 3. When grown to confluence on semipermeable membrane inserts, PTC monolayers acquire trans-epithelial electrical resistance in the order of 100 Ω·cm2. 4. PTC cultures stain uniformly for cytokeratin, an epithelial marker, but not vimentin, a mesenchymal marker. 5. The activity of brush border enzymes such as alkaline phosphatase, alanine aminopeptidase, and gamma-glutamyl transpeptidase are strong in PTC cultures. 6. Cyclic AMP production is increased in PTC cultures when treated with parathyroid hormone but not vasopressin (9).

4

Notes

1. When KHS is prepared, the CaCl2 should be added slowly, with stirring, to the final solution. If it is added too quickly, a cloudy precipitate can form and the solution must be discarded and prepared again. 2. We routinely used KHS as a buffer during cell isolation. However, PBS or other buffered salt solutions or culture media can be used. 3. The collagenase should be dissolved in KHS at 1 mg/mL and filter sterilised before use. We use 2 mg of collagenase per gram of cortical tissue. 4. During the collagenase digestion, it is important that the solution contains calcium, which is necessary for optimum collagenase activity. 5. After each wash make sure the cell pellet is fully resuspended. If clumps are observed it is necessary to pipette the suspension up and down several time to encourage dissociation. 6. Make sure not to overload the Percoll gradient because this will make identification of the PTC band difficult. 7. This protocol yields proximal tubule fragments, cell aggregates, and single cells and thus making cell counting unreliable. 8. A small amount fibroblastic cell contamination may occur. Growth in defined serum free media will limit their growth. 9. Coating culture flasks, dishes, or inserts with collagen or laminin can promote cell attachment and growth. Acknowledgements The invaluable assistance provided by the PAH Tumour Tissue Bank and urologists of Princess Alexandra Hospital in the procurement of human renal tissue is gratefully acknowledged. Our research is funded in part by the National Health and Medical Research Council of Australia, the Juvenile Diabetes Foundation, and the Princess Alexandra Hospital Research and Development Foundation.

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References 1. Al Awqati Q, Schwartz GJ. (2004) A fork in the road of cell differentiation in the kidney tubule. J Clin Invest. 113, 1528–1530. 2. Phillips AO, Steadman R. (2002) Diabetic nephropathy. The central role of renal proximal tubular cells in tubulointerstitial injury. Histol Histopathol. 17, 247–252. 3. Nakagawa T, Kang DH, Ohashi R, et al. (2003) Tubulointerstitial disease: role of ischemia and microvascular disease. Curr Opin Nephrol Hypertens. 12, 233–241. 4. Handler JS, Perkins FM, Johnson JP. (1980) Studies of renal cell function using cell culture techniques. Am J Physiol. 238, F1–F9. 5. Montrose MH, Murer H. (1990) Polarity and kinetics of Na(+)-H+ exchange in cultured opossum kidney cells. Am J Physiol. 259, C121–C133. 6. Orosz DE, Woost PG, Kolb RJ, et al. (2004) Growth, immortalization, and differentiation potential of normal adult human proximal tubule cells. In Vitro Cell Dev Biol Anim. 40, 22–34. 7. Prozialeck WC, Edwards JR, Lamar PC, et al. (2006) Epithelial barrier characteristics and expression of cell adhesion molecules in proximal tubule-derived cell lines commonly used for in vitro toxicity studies. Toxicol In Vitro. 20, 942–953. 8. Vinay P, Gougoux A, Lemieux G. (1981) Isolation of a pure suspension of rat proximal tubules. Am J Physiol. 241, F403–F411. 9. Johnson DW, Brew BK, Poronnik P, et al. (1997) Transport characteristics of human proximal tubule cells in primary culture. Nephrology. 3,183–194. 10. Goligorsky MS, Hruska KA. (1986) Transcytosis in cultured proximal tubular cells. J Membr Biol. 93, 237–247.

Chapter 3

Propagation and Culture of Renal Fibroblasts Lauren Grimwood and Rosemary Masterson

Abstract In this chapter we describe a reliable and reproducible method for the selective propagation and culture of renal fibroblasts derived from explantation of renal cortical tissue in vitro. The chapter outlines how primary renal interstitial fibroblasts are derived from explants grown in medium supplemented with foetal calf serum. The subculture of confluent cells and their ultimate characterisation as fibroblasts through immunohistochemical and immunocytochemical techniques are described in detail. Keywords Propagation, Interstitium, Fibroblasts, Culture, Explants, Characterisation, Immunocytochemistry, Kidney

1

Introduction

Tubulointerstitial fibrosis is the final common pathway of all renal disease regardless of aetiology and is the most accurate histological predictor of disease progression (1). Although other cells may play a role in the pathogenesis of scarring, it is widely accepted that the renal interstitial fibroblast is the pivotal effector cell in the development of fibrosis. Given the central role played by these cells in the development of fibrocontractive disease, it is critical to be able to study their behaviour in isolation through an in vitro culture system. The study of factors affecting phenotypic change and growth of fibroblasts has typically involved the cultivation of established primary cell lines or alternatively the use of fibroblasts derived from outgrowth of primary explants of renal cortical tissue. Immortalised cell lines have been specifically selected for their ability to rapidly proliferate but present the problem of being virally mutated cells, which differ in behaviour to endogenous non-transformed cells. Conversely primary cultures of fibroblasts provide a system in which the growth of cells can be investigated without dissociating them totally from their normal environment. A variety of methodologies and techniques has been used to isolate renal interstitial fibroblasts from various segments of the kidney. These include differential From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_3, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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sieving, centrifugation, collagenase digestion, separation using magnetic beads, and outgrowth culture, amongst others (2–4). In this chapter, we describe a simplified method used in our laboratory for the preparation of selective culture of fibroblasts that has proven to be both efficient and reproducible (5, 6). The method is based on the outgrowth of cells from explanted renal tissue and it resembles established methodologies used in the culture of hepatic (7) and skin (8) fibroblasts. The technique involves dissecting the cortical tissue away from the whole kidney section, mincing it finely, and scratching the tissue explant into a gelatin-coated petri dish base. The tissue is then incubated in enriched medium for 10–14 days until the monolayer has covered 75% of the dish. A large proportion of the cells may initially display a cobblestone appearance typical of tubuloepithelial cells before being succeeded by the classic “fingerprint” pattern characteristic of a fibroblast monolayer. The cells are then lifted from the base of the petri dish by trypsinisation, before being re-suspended in medium and transferred to culture flasks until confluent. At this stage the origin of the resident cell populations is defined by using immunocytochemistry and immunofluorescent techniques, which identify specific cytoskeletal markers. Cells are characterised as fibroblasts on the basis of positive staining for alpha-smooth muscle actin, vimentin, and collagen I in conjunction with an absence of endothelial and pancytokeratin staining. Once it is established that the cell monolayer is phenotypically that of fibroblasts, the cells can be cryogenically stored following the second passage of their primary culture. The subsequent culture of these fibroblasts affords the opportunity to study their functional response following exposure to putative agonists and antagonists of fibroblast function, which may ultimately have therapeutic application in the treatment of fibrocontractive disease.

2 2.1 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Materials General Sterile Cell Culture Materials 100-mL and 200-mL media bottles (Schott, Duran, Germany). 60-mm2 cell culture Petri dishes (TPP, Trasadingen, Switzerland). 50-mL centrifuge tubes (TPP). 25-cm2 and 75-cm2 tissue culture flasks with vent caps (TPP). 1.8-mL cryogenic vials (TPP). 200-µL and 1,000-µL pipette tips. Borosilicate graduated glass 5-mL and 10-mL pipettes. Cell scraper (Corning Incorp, Costar, NY, USA). Disposable 60-mL syringe. Disposable 0.2-µm pore syringe filters (Millipore, Billerica, MA, USA). Reusable 300-mL filter unit and 0.2-µm pore cellulose acetate filters (Advantec MFS Inc., Dublin, CA, USA).

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12. Sterile containers for tissue collection and dissection (e.g. a basic sterile dressing pack). 13. No 22 scalpel blades (Swann-Morton Ltd, Sheffield, England). 14. Plastic Pasteur pipettes. 15. Waste bottle. 16. Deionised water (dH2O).

2.2 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

2.3

Specialist Cell Culture Equipment Biohazard hood. Inverted light microscope. Vacuum pump. Tissue culture incubator set at 37°C with 95% O2/5% CO2. Fluorescent microscope. Water bath. Autoclave. Pipet-Aid™ (BD Falcon, BD Biosciences, San Jose, USA) or equivalent for aspirating and changing media. Borosilicate glass test tubes (Chase-Scientific Glass, Rockwood, TN, USA). Haemocytometer. Mechanical tally counter. Cryogenic Dewar for storage of frozen cells. –70°C freezer.

Reagents

1. Sterile 0.01M Dulbecco A phosphate-buffered saline (PBS) pH 7.3 (Oxoid, Hampshire, England): prepare by reconstituting each PBS tablet in 1 L dH2O. Sterilise by autoclaving. 2. Matrix Coating Solution: 2% solution of Type B gelatin from bovine skin (Sigma–Aldrich, St. Louis, IL, USA). 3. Tissue Collection Buffer: combine 100 mL of 1× Hank’s Balanced Salt Solution (HBSS; SAFC Biosciences, Lenexa, Kansas, USA), 200 µL of 1 M Hepes Buffer (SAFC Biosciences), 2.5 µL gentamicin (10 mg/mL gentamicin as gentamicin sulphate; SAFC Biosciences) and 47 µL sodium bicarbonate 7.5% in solution (Sigma–Aldrich). Filter through a 0.2-µm filter using a 60-mL syringe. Prepare immediately prior to use and store on ice. 4. Enriched medium (Dulbecco’s Modified Eagle Medium [DMEM] with 20% serum): for each 400 mL of medium, combine 80 mL filter-sterilised fetal calf serum (FCS), 10 mL of 1 M Hepes (SAFC Biosciences), 4 mL of 200 mM glutamine (SAFC Biosciences), and 8 mL penicillin–streptomycin (formulated

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to contain 5,000 U/mL penicillin and 5,000 µg/mL streptomycin; Sigma– Aldrich), and make up to 400 mL with 1× DMEM (SAFC Biosciences). This solution is sterilised by passing through a sterile 0.2-µm cellulose acetate filter unit into a sterile 500-mL glass bottle by vacuum extraction. The bottle is capped and stored at 4°C (see Note 1). Maintenance medium: as above for enriched medium but supplemented with 10% FCS (40 mL FCS in 400 mL medium). Ethylenediaminetetraacetic acid (EDTA): 0.02% solution (Sigma–Aldrich). Enzyme dissociation solution: 1:10 dilution v/v of 1× trypsin (SAFC Biosciences) in 0.02% EDTA solution. Freezing medium: dimethyl sulphoxide (DMSO) in medium 10% v/v (1 mL DMSO in 9 mL of enriched medium supplemented with either 10% or 20% FCS). Liquid nitrogen (N2).

2.4

Immunoperoxidase and Immunofluorescent Cytochemistry

2.4.1

General Materials

1. 2. 3. 4. 5. 6. 7. 8.

Established primary renal cell line. Sterile glass cover slips (22×40 mm). 0.01 M PBS, pH 7.3. Borosilicate glass test tubes. Filter paper. Fixative e.g. methanol, acetone, or 4% paraformaldehyde (see Note 2). Glass microscope slides with frosted edges. Hydrophobic wax pen (Dako, Glostrup, Denmark).

2.4.2

Immunoperoxidase Cytochemistry

1. Primary antibody (Table 3.1). 2. Antibody diluent (Dako). 3. Peroxidase IgG kits containing normal serum and appropriate biotinylated secondary antibody (2°Ab) (Table 3.1) (Vectastain Kit; Vector Laboratories, Burlingame, CA, USA). 4. Avidin-biotin complex (ABC) Elite kit (Vector Laboratories). 5. Chromogen substrate: dissolve 1 × 3′3-diaminobenzidine (DAB) tablet (Dako) in 10 mL of 0.01 M PBS and filter to remove undissolved material. To prepare working solution, add 1.5 µL of 30% v/v H2O2 to 4 mL of DAB stock immediately prior to use (see Note 3). 6. Harris haematoxylin (Fronine, Melbourne, Victoria, Australia). 7. Aquamount™ (BDH Laboratories, Poole, UK).

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Table 3.1 Immunocytochemical anti-sera specifications Antibody

Clone

IgG Source

Dilution

Specificity

Company

α-SMA

1A4

Mouse

1:50

Dako (Glostrup, Denmark)

Cytokeratin Vimentin Rat endothelial cell antigen (RECA) Von Willebrand Factor (VWF) Collagen I

MNF116 V9 MCA970

Mouse Mouse Mouse

1:10 1:50 1:20

Smooth muscle isoform of actin Pan-keratin Vimentin RECA

Polyclonal

Rabbit

1: 100

VWF

Polyclonal

Goat

1:50

Collagen I

Collagen III

Polyclonal

Goat

1:100

Collagen III

Desmin

D33

Mouse

1:100

Desmin

Dako Dako Serotec (Oxford, England) Dako Southern Biotechnology (Birmingham, AL, USA) Southern Biotechnology Dako

8. Humidified chamber (e.g. plastic lunch box lined with blotting paper and soaked with 0.01 M PBS).

2.4.3

Immunofluorescent Cytochemistry

1. Unlabeled primary antibody or fluorochrome conjugated primary antibody (Table 3.1). 2. Aluminium foil. 3. Fluorescein isothiocyanate (FITC)-conjugated anti-IgG. 4. Fluoromount (Dako). 5. Fluorescent microscope.

3

Methods

The procedures described in Sects. 3.1–3.4 offer techniques to establish and characterise a primary fibroblast cell line cultivated from explanted renal cortical tissue (Fig. 3.1). The methods outline 1) preparation of cell culture reagents, 2) explanting of renal tissue, 3) subculturing of a cultivated primary cell line, and 4) characterisation of an established cell line.

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Fig. 3.1 Propagation of renal interstitial fibroblasts by explanting

tissue minced in ‘cross-wise’ fashion

kidney

cortex

Petri dish

cellular outgrowth

tissue culture flask

subcultured monolayer

3.1

Gelatin Coating of Petri Dishes

Petri dishes are coated with 1% gelatin solution prior to tissue explanting. This allows minced tissue to be adequately secured to the petri and provides a substrate for the initial cultivation of cell populations. 1. Dilute 2% gelatin solution with 0.01 M PBS (1:1 dilution) and filter sterilise into a centrifuge tube using a disposable syringe and 0.2-µm-pore filter unit. 2. Coat entire surface of Petri dish with 1 mL of 1% gelatin solution and incubate for 30 minutes at 37°C. 3. Remove excess gelatin solution and rinse Petri with 2 mL of 0.01 M PBS prior to explanting.

3.2

Propagation of Renal Interstitial Fibroblast Cultures from Kidney Explants

1. Collect kidney tissue in Tissue Collection Buffer and store at 4°C until required for explanting (see Note 4). 2. Set out surgical dressing kit onto the sterile field in biohazard hood.

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3. Pour out HBSS and kidney into a small sterile container (e.g. dressing pack tray) and use the forceps to gently transfer kidney tissue into a clean section of the tray. 4. Cover with warm enriched medium (1× DMEM/20% FCS) to keep moist and dissect with the forceps and scalpel blade. 5. Carefully peel the renal capsule back and use the scalpel blade to make a longitudinal incision. An anatomical division between the medulla and cortex should be visible. Cut off small sections of cortex, place into the biopsy tray, and dice until a minced consistency is achieved. 6. Transfer minced tissue into a gelatin-coated Petri dish. 7. Adhere tissue to the gelatin surface or Petri dish by making cross-wise scratches across surface of Petri dish. Continue until at least 75% of the dish is covered (5, 6) (see Note 5). 8. Cover explant with 2 mL of 1× DMEM/20% FCS using a plastic Pasteur pipette, label and date. 9. Incubate overnight at 37°C with 95% O2/5% CO2. 10. Supplement tissue with an additional 2 mL of 1× DMEM/20% FCS 24 h post explanting procedure. 11. Incubate explants for 72h to allow for initial cell populations to be established. 12. Aspirate explanting medium into a waste bottle using a 5-mL pipette and replenish with fresh 4 mL of 1× DMEM/20% FCS to remove cellular debris and non-adherent cells that may cause contamination. 13. Change medium twice weekly for 10–14 days or until the monolayer has covered approximately 75% of the dish surface (see Note 6).

3.3

Subculture of Primary Cell Cultures

Both primary and subcultured cells are lifted from their growing surface by treatment with a combination of EDTA and the enzyme trypsin. FCS-supplemented tissue culture medium routinely contains a mixture of divalent cations and proteins that inhibit trypsin. In each case cells are therefore first washed with Ca2+/Mg2+-free PBS to reduce the concentration of these trypsin inhibitors. EDTA is a calcium chelator that can be used to “mop” up any remaining divalent ions. Finally, after cells detach, the enzyme reaction is quenched by adding an excess of fresh medium to prevent over digestion and damage to the cells.

3.3.1

Lifting Explant Monolayer Cultures

1. Remove medium from the Petri dish and discard as biological waste. 2. Wash the monolayer twice with 2 mL of 0.01 M PBS to remove residual medium. 3. Wash cells with 1 mL of EDTA solution to assist with cell dissociation.

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4. Replace with 1 mL of cell dissociation solution (1× trypsin/EDTA). 5. Incubate for 10–15 min at 37°C. 6. Under an inverted light microscope, check that cells have detached from the dish surface. 7. Gently tap the base of the Petri dish to detach the remaining cells and add 2 mL of warm enriched medium (1× DMEM/20% FCS) to inhibit the trypsin reaction. 8. Gently scrape the explant with a cell scraper to assist the detachment of cells. 9. Transfer cell suspension into sterile 25-cm2 tissue culture flask. 10. Add 3 mL of enriched medium (1× DMEM/20% FCS). 11. Incubate overnight at 37°C with 95% O2/5% CO2. 12. Change medium twice weekly until cultures are 100% confluent (see Note 7).

3.3.2

Maintenance of Primary Renal Fibroblast Subcultures

1. Remove medium from 25-cm2 tissue culture flasks and discard the medium as biological waste. 2. Wash the monolayer twice with 2 mL of 0.01 M PBS. 3. To detach cells, add 1 mL of cell dissociation medium (1× trypsin/EDTA) and incubate flask at 37°C for 10–15 min. 4. Gently tap the base of the flask to separate cells. 5. Add 2 mL of warm maintenance medium (1× DMEM/10% FCS) to inhibit the trypsin reaction. 6. Gently aspirate cell suspension and transfer contents into a 75-cm2 tissue culture flask. 7. Add 10 mL of maintenance medium, label and date the flask. 8. Incubate the flask at 37°C with 95% O2/5% CO2 . 9. Change medium twice weekly until cultures are completely confluent (see Note 8).

3.4

Characterisation of Primary Renal Fibroblasts

Primary cell cultures isolated from renal tissue explants yield a mixed population of cells due to the complex composition of the kidney. It is therefore important to remember that explanting produces a heterogenous population of cell lineages consistent with a variety of anatomical locations (reviewed in refs. (9, 10)). Identification of fibroblasts per se is however somewhat problematic. The low immunogenicity of fibroblasts has meant that the production of fibroblast-specific antisera has been difficult. Fibroblasts are therefore usually characterised by a series of inclusion/exclusion criteria according to their distinctive morphological and biochemical features. This includes the exhibition of an elongated, spindleshaped cell in culture, “finger print” patterned monolayer when confluent, consistent immunocytochemical staining for mesenchymal markers, and negative staining for epithelial and endothelial makers (see Table 3.2 and Fig. 3.2) (11).

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Table 3.2 Cell characteristics and phenotype Markers

Phenotype

α-Smooth muscle actin (α-SMA)

Myofibroblasts, vascular smooth muscle cells, some mesangial cells Fibroblasts, mesangial cells, vascular smooth muscle cells Epithelial cells Vascular smooth muscle cells, mesenchymal cells, mesangial cells, some myofibroblasts Fibrogenic cells Endothelial cells

Vimentin Cytokeratin Desmin Collagen I Rat endothelial cell antigen (RECA)

Fig. 3.2 Staining characteristics of sub-cultured cells (passage 3); SMA, smooth muscle actin (reproduced from ref. (11) with permission from Elsevier) (see Color Plate 1)

3.4.1

Growth Characteristics

In the method described here, many of the initial outgrowths from renal cortical tissue explants display a cobblestone morphology consistent with an epithelial cell origin. Prolonged culture (14–20 days) results in the accumulation of a homogeneous population of fibroblastic cells, recognisable by their spindle shape. At confluence, monolayers of fibroblasts form a “finger print” pattern without the “hillocks” typical of mesangial cells.

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L. Grimwood, R. Masterson

Immunocytochemistry

Both immunoperoxidase and immunofluorescent techniques are commonly employed to characterise cells with the use of specific anti-sera. Generally, chromogen-based techniques are used for the quantitative analysis, while immunofluorescent labeling is better suited to qualitative analysis.

3.4.2.1

Immunoperoxidase Staining

1. Grow primary cell line overnight or until 70% confluent on sterile cover slips in a Petri dish supplemented with enriched medium at 37°C with 95% O2/5% CO2. 2. Remove medium and wash monolayer with 0.01 M PBS. 3. Fix cells for 5 min at 4°C (see Note 2), remove methanol, and air dry. 4. Outline cover slip with wax pen to conserve antibody. 5. Wash cover slips twice with 0.01 M PBS (5 min/wash). 6. Block non-specific binding sites by incubating sections with the appropriate diluted normal serum for 10 min. 7. Gently blot with tissue paper to remove excess serum solution. 8. Incubate for 60 min in a humid chamber with a 1:10–1:100 dilution of primary antibody in antibody diluent (Table 3.1). 9. Wash twice with 0.01 M PBS (5 min/wash). 10. Incubate with biotinylated secondary antibody (Vector) for 10 min. 11. Wash twice with 0.01 M PBS (5 min/wash). 12. Incubate with ABC Elite solution (Vector) for 15 min (12). 13. Prepare peroxidase chromogen substrate working solution (see Note 3). 14. Wash twice with 0.01 M PBS (5 min/wash). 15. Apply chromogen substrate to sections for 2–5 min. Monitor staining on wet slides using a light microscope. 16. Wash in H2O for 5 min to terminate reaction. 17. Counterstain with Harris haematoxylin for 1 min. 18. Wash in H2O for 5 min. 19. Mount cover slips onto glass microscope slides with AquamountTM (BDH) and examine using a light microscope (Table 3.2).

3.4.2.2

Immunofluorescent Labeling Technique

1. Grow and fix, and pre-treat cells on sterile cover slips as above (Sect. 3.4.2.1, steps 1–7). 2. Incubate with primary antibody diluted with 0.01 M PBS for 60 min (Table 3.1). 3. Wash twice with 0.01 M PBS (5 min/wash). 4. Incubate with FITC conjugate/0.01 M PBS for 30 min. Wrap in aluminium foil during FITC incubation to minimise light exposure (see Note 9). 5. While wrapped in foil, wash twice with 0.01 M PBS (5 min/wash).

3 Propagation and Culture of Renal Fibroblasts

35

6. Use Fluoromount (Dako) to mount cover slips on microscope slides. 7. Visualise under a fluorescent microscope set with appropriate filters (495– 525 nm). 8. Determine cell phenotype (Table 3.2, Fig. 3.2).

3.5

Freezing Cell Cultures for Cryogenic Storage

Once the monolayer is 100% confluent in a 75-cm2 flask, the cells can be passaged and frozen in 10% DMSO-supplemented medium for cryogenic storage. DMSO is added to the cells as a cryoprotectant. DMSO protects cells in two ways. Firstly, by penetrating the cell and binding to water molecules, DMSO prevents the loss of water from the cytoplasm during freezing, thus preventing cellular dehydration and maintaining intracellular physiology. Secondly, by slowing the rate of freezing, DMSO prevents the formation of ice crystals within the cell. 1. Remove medium from monolayer and discard as biological waste. 2. Wash monolayer twice with 5 mL of 0.01 M PBS. 3. Dislodge cells with 2 mL of dissociation solution (1× trypsin/EDTA) and incubate flask at 37°C for 10–15 min. 4. Gently tap the base of the flask to separate the cells. 5. Add 5 mL of warm maintenance medium (1× DMEM/10% FCS) to inhibit the trypsin reaction. 6. Gently aspirate cell suspension and transfer contents into a 50-mL centrifuge tube. 7. Centrifuge the cell suspension for 5 min at 700×g. 8. Remove the supernatant from the cell pellet and discard. 9. Resuspend the cell pellet in 1.8 mL of cold freezing medium and transfer into a sterile cryogenic vial (see Note 10). 10. Label and date the vial with appropriate details using a grey lead pencil. 11. Store at −70°C overnight. 12. Transfer vial to a Dewar filled with liquid nitrogen for long-term cryogenic storage.

3.6

Thawing of Cryogenically Stored Cells

1. Set up a biohazard hood with warm medium, sterile pipettes, centrifuge tube, and flask. 2. Remove the cryogenic vial from liquid nitrogen and quickly thaw it in a 37°C water bath. 3. Transfer the contents to a 50-mL centrifuge tube. 4. Slowly add 10 mL of warm 1× DMEM/20% FCS medium in a drop-wise fashion to the cell suspension and simultaneously gently agitate the container.

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L. Grimwood, R. Masterson

5. Aspirate the suspension to separate cells and transfer the cells into a 75-cm2 cell culture flask. 6. Vent the cap and incubate the cells at 37°C overnight to allow the cells to adhere to the flask surface. 7. Remove the DMSO-containing medium from the cell layer after 24 h and replace it with 10 mL of fresh medium and return the flask to the incubator. 8. Change medium twice weekly and maintain until ready for experimentation.

4

Notes

1. It is important that all cell culture reagents especially growth medium is preheated to 37°C prior to use in order to maintain homeostasis for cellular integrity and viability. 2. The objective of fixation is to preserve cells and tissue constituents in as close a life-like state as possible. However, fixation itself constitutes a major artefact. Two main types of fixatives are used in cytochemistry: precipitating (e.g. methanol, acetone) and cross-linking (e.g. paraformaldehyde). Although fixation by precipitation may destroy fewer antibody-binding sites, is does not preserve the three-dimensional organisation of specimens, and is therefore not recommended for the study of cytoskeletal proteins. 3. DAB is a carcinogen and must be handled and disposed of in accordance with the Material Safety Data Sheet (MSDS). DAB stock solution can be stored in a light-proof container for up to 3 days at 4°C. 4. Explanting can be used to propagate fibroblasts from both human and rodent kidneys. In the case of mice and rats, collect the kidney with the capsule intact. For human kidneys, it is necessary to excise approximately 1-cm2 portions of human tissue. 5. Approximately 1.0–1.5 mm3 of minced tissue per Petri dish is adequate to achieve successful cultivation of initial outgrowths. 6. Primary cell populations emerge from tissue within 24–72 h of explanting depending on the maturity of the kidney as well as the individual explanting technique. Initially these small clusters consist of a mixed population of epithelial and mesenchymal cells. A large proportion of these cells display a cobblestone appearance that resemble epithelial morphology, while a smaller fraction appear as spindle-shaped fibroblasts. Eventually fibroblastic cells proliferate significantly to become the predominant resident cell type and exhibit the classical “fingerprint” pattern monolayer. At this point primary cell populations from an explant are ready for enzymatic disaggregation using trypsin and are then re-seeded into a fresh vessel to become a secondary culture. 7. When initial primary renal fibroblasts are 100% confluent, the monolayer should be passaged using the trypsinisation method. Transfer cells into larger culture vessels to establish subcultures. 8. Once a secondary passage of primary fibroblast cultures is achieved, cells may be characterised for experimentation and/or cryogenically stored (10). 9. When using directly conjugated antibodies, apply the antibodies immediately after blocking with dilute normal serum (see Sect. 3.4.2.2, step 7) and incubate for 1 h, with the container wrapped in foil. Proceed as per protocol with 0.01 M PBS washes and mount with Fluoromount before examining the cells with a fluorescent microscope. 10. An exothermic reaction that can damage the cell membrane is produced when DMSO is added to warm medium. Therefore, before trypsinising the monolayer, prepare 10% DMSO medium and refrigerate until cold. Ensure that the cell pellet is evenly resuspended in cold freezing medium and that no clumps are visible. Transfer to a sterile cryogenic vial as per protocol.

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References 1. Bohle, A., Mackensen-Haen, S., and von Gise, H. (1987) Significance of tubulointerstitial changes in the renal cortex for the excretory function and concentration ability of the kidney: a morphometric contribution. Am J Nephrol. 7, 421–433. 2. Sharpe, C. C., Dockrell, M. E. C., Noor, M. I., Monia, B. P., and Hendry, B. M. (2000) Role of ras isoforms in the stimulated proliferation of human renal fibroblasts in primary culture. J. Am. Soc. Nephrol. 11, 1600–1606. 3. Sommer, M., Schaller, R., Funfstuck, R., Bohle, A., Bohmer, F. D., Muller, G. A., and Stein, G. (1999) Abnormal growth and clonal proliferation of fibroblasts in an animal model of unilateral ureteral obstruction. Nephron. 82, 39–50. 4. Clayton, A., Steadman, R., and Williams, J. D. (1997) Cells isolated from the human cortical interstitium resemble myofibroblasts and bind neutrophils in an ICAM-1 dependent manner. J. Am. Soc. Nephrol. 8, 604–615. 5. Hewitson, T. D., Martic, M., Kelynack, K. J., Pedagogos, E., and Becker, G. J. (2000) Pentoxifylline reduces in vitro renal myofibroblast proliferation and collagen secretion. Am. J. Nephrol. 20, 82–88. 6. Kelynack, K. J., Hewitson, T. D., Nicholls, K. M., Darby, I. A., and Becker, G. J. (2000) Human renal fibroblast contraction of collagen I lattices is an integrin-mediated process. Nephrol. Dial. Transplant. 15, 1766–1772. 7. Blazejewski, S., Preaux, A. M., Mallat, A., Brocheriou, I., Mavier, P., Dhumeaux, D., Hartmann, D., Schuppan, D., and Rosenbaum, J. (1995) Human myofibroblastlike cells obtained by outgrowth are representative of the fibrogenic cells in the liver. Hepatology. 22, 788–797. 8. Gabbiani, G., Ryan, G. B., and Majne, G. (1971) Presence of modified fibroblasts in granulation tissue and their possible role in wound contraction. Experientia. 27, 549–550. 9. Freshney, R. I. (1992) Animal Cell Culture: A Practical Approach. Oxford University Press. NY. USA. pp. 1–14.Second Ed. pp. 1–14. 10. Grupp, C., and Muller, G.A. (1999) Renal fibroblast culture. Exp. Nephrol. 7, 377–385. 11. Hewitson, T. D., Tait, M. G., Kelynack, K. J., Martic, M., and Becker, G. J. (2002) Dipyridamole inhibits in vitro renal fibroblast proliferation and collagen synthesis. J. Lab. Clin. Med. 140, 199–208. 12. Hsu, S. M., Raine, L., and Fanger, H. (1981) Use of avidin-biotin peroxidase complex (ABC) in immunoperoxidase technique: a comparison between ABC and unlabeled antibody (PAP) procedures. J. Histochem. Cytochem. 29, 577–580.

Chapter 4

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

Abstract Animal models of renal disease have provided valuable insights into the pathogenesis of acute and chronic kidney disease. Extension of these models to the mouse has become increasingly important with the development of gene knockout and transgenic animals. In this review we discuss a range of models that can be used to mimic the mechanisms of human renal disease. While not perfect, the careful and ethical use of these models offers the opportunity to examine individual mechanisms in an accelerated time frame. Keywords Kidney disease, Animal models, Nephrotoxins, Progression.

1

Introduction

Our understanding of human renal disease depends largely on studies of animal models. What do experimental models offer? Decades may elapse between injury and clinical manifestations in human renal disease. The rapid onset of experimental models therefore offers a more feasible means of studying pathogenesis and possible therapies. Furthermore, in many cases the pathology of human disease is multifactorial. The careful use of animal models affords the opportunity to study individual mechanisms, such as interstitial fibrosis without confounding glomerular pathology (1). An animal model can therefore be used to delineate mechanisms and test putative interventions in a disease process (2).

2

Ethical and Moral Issues

Compliance with statutory regulations and satisfying the concerns of animal ethics committees is an important part of medical research. A thorough understanding of animal welfare is therefore an essential prerequisite for commencing these studies.

From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_4, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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Because legal constraints vary widely around the world, a detailed discussion of these is beyond the scope of this review. However, it is necessary at the outset for all scientists to acknowledge that some in the community regard any animal experimentation as an unjustified means of pursuing knowledge about human disease (3). The community as a whole therefore has the right to expect that scientists will do their utmost to minimize the use of animal models where possible. For example the cell culture protocols outlined in Part I of this book offer an ideal means to initially test hypotheses.

3

Species, Strain and Gender

Many animals including monkeys, dogs, sheep, rabbits, and particularly rats, have been used to mimic human disease. Although mouse models are less well established, we increasingly recognize that the genetic modification of mice provides exciting opportunities to examine the influence of gene deletion and over expression in the pathogenesis of renal disease. Not surprisingly then, as in other fields of medicine, attention has recently focused on the use of the mouse. In this review we discuss a range of models that can be used to mimic the mechanisms of human renal disease. Our emphasis is on artificial rather than spontaneous genetic models that may be difficult to source. We deliberately emphasize mouse models, since they are currently most relevant to genetic modifications. With all animal studies it is important to obtain mice from a thoroughly reputable source, to ensure their genetic background as required, and that they are free from parasites and other animal disease that can obscure the results. Unfortunately in all animal studies, strain and species mutation is problematic and pilot studies are usually required to determine the optimal experimental conditions, or sometimes the optimal species/strain. Observational studies in humans have suggested that the rate of progression of renal disease is influenced by gender. Deterioration of renal function in patients with chronic renal disease is generally more rapid in men than in women, independent of differences in blood pressure or serum cholesterol levels. Likewise, potentially important gender differences exist in experimental models. Female rats are more resistant to acute renal ischemia (4) while male mice survive better than female mice after cisplatin administration (4). Although the reasons for this remain unclear, several groups are examining the role of estrogen and testosterone.

4

Surgical Models of Acute and Chronic Renal Failure

Surgical manipulations are widely used to model human disease. These may reproduce the injury that causes renal disease (e.g. unilateral ureteric obstruction [UUO]) or reproduce the consequences of renal injury (e.g. renal ischaemia, reduction in renal

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mass). Outbred but uniform rat strains are most commonly used. Although technically demanding, unlike immune-mediated disease, many surgical models can be easily adapted to the mouse.

4.1

Ureteric Obstruction

UUO has become one of the most popular models of chronic kidney disease. Although it is not one of the most common causes of kidney disease in humans, interest in obstructive renal disease has been kindled by renewed interest in the mechanisms of tubulointerstitial disease. The model has the advantages of reproducibility (hence inter-animal variation is not a problem), a short time-course, and it is easy to perform, even in the mouse (5). The immediate consequence of urinary tract obstruction is an increase in pressure within the urinary tract, proximal to the cause of obstruction. This results in dilation of the renal pelvis. Increased renin–angiotensin system activity is an early renal hemodynamic consequence resulting in renal vasoconstriction. The subsequent interstitial inflammatory response is initially characterized by macrophage infiltration (Fig. 4.1), tubular dilation, and renal tubular apoptosis leading to tubular atrophy (6). A major deficiency of this model is the inability to accurately measure changes in renal function since the remaining unobstructed kidney compensates for much of the loss of function in the obstructed kidney. Hence, only modest changes occur in such markers of decreased overall function such as blood creatinine or urea

Fig. 4.1 Interstitial accumulation of monocytes–macrophages after unilateral ureteric obstruction (UUO). Micrograph shows immunoperoxidase staining with the monoclonal antibody clone ED-1, 3 days post-UUO (see Color Plate 2)

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concentrations. If such data is important, the contralateral kidney can be removed, usually prior to temporary obstruction. Variations of this model have included the obstruction of kidneys in neonatal rats (7) and the relief of obstruction after a few days (8). The latter has proved a particularly useful model to study regeneration; the period of obstruction being inversely related to the capacity for repair. While in rats short term UUO appears to be completely reversible, UUO for greater than 72 hours leads to a permanent loss of function (8). This rapid loss of function is one of the disadvantages of UUO. It is so rapid, severe, and irreversible that it limits the testing of possible ameliorating interventions where profound effects are not expected.

4.2

Sub-total Nephrectomy or Infarction

Sub-total nephrectomy, 5/6 nephrectomy, is probably the most established method of modelling the progressive renal failure seen with loss of renal mass. It is most commonly performed in the rat (9). Rabbit and mouse nephrectomy models have been less extensively studied, and appear less reliable. Rather than mimicking a renal disease per se, sub-total nephrectomy reproduces the consequences of reducing functional renal mass. The predominant pathological abnormalities are glomerulosclerosis and tubulointerstitial fibrosis. It is important to appreciate that two very different models are encompassed by the expression “5/6 nephrectomy”. In the ligation model, one kidney is removed (uninephrectomy) and then ligation of polar branches of the renal artery interrupts blood flow to the poles of the remaining kidney. Typically 2/3 branches of the renal artery in the rat are ligated. The natural history of this infarction model is typically associated with severe hypertension. In the ablation model, one kidney is removed, and approximately 50% of the remaining kidney is removed by polar excision 1–2 weeks later. Severe hypertension is not a usual feature of this model. The rate of progression to renal failure is very closely related to the amount of tissue excised. As a guide, a 70% total reduction in renal mass in the rat results in renal failure in about 4 months. It is necessary to carefully calculate the proportion of excised tissue. If the weight of the first kidney is recorded at the time of uninephrectomy, then the excised tissue weighed after the subsequent surgery provides a reasonable estimate of the total reduction of renal mass (60–80%), minimizing variation in extent of nephrectomy. The variability of the model however still poses problems for interventionist experiments. It is difficult to run sufficiently powered experiments without pairing for renal function at a certain post-operative time point. Reducing renal mass by uninephrectomy is also often used as a means of accelerating other models of renal disease. For instance uninephrectomy exacerbates the effects of a high-fat diet on kidney structure and function (10). In these cases, the left kidney is usually removed because it is lower and more surgically accessible.

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4.3

45

Ischemic Acute Renal Failure

Ischemic acute renal failure (IARF) remains the most common cause of acute renal failure in the adult human population. Its importance is underscored by the fact that morbidity and mortality associated with acute renal failure have not appreciably declined in recent years (11), despite attempts to develop ameliorating strategies or molecules. Ischemia/reperfusion is a cause of organ damage in diverse clinical circumstances. Three different models of IARF are commonly employed. Bilateral clamping of the renal pedicle has been used by many groups to simulate IARF induced by acute severe hypotension (12). Clamping of a single renal artery induces a unilateral disease, valuable because comparison with changes seen in the contralateral kidney are useful, and overall survival is more reliable than with bilateral clamping. Finally, removal of a kidney prior to occlusion of the contralateral renal artery avoids the confusion caused by a functioning and non-functioning kidney, this more closely mimics the situation often occurring in renal transplantation (13). The consequences of IARF are proportional to the period of ischemia, and have to be determined for the individual rat strain and age. As a guide, 30–50 min of occlusion is generally used in uninephrectomized rats. However, the functional effect remains highly variable, with even small differences in ischemia time significantly changing the characteristics of the model. Again these differences can make comparative studies problematic.

5

Nephrotoxic Models

The ability of kidneys to concentrate and metabolize chemicals makes them particularly prone to toxic damage. The high vascularization of the kidney, the presence of specific transport mechanisms increasing cellular uptake, the concentrating ability of the kidney and the high metabolic rate of tubular cells all exacerbate toxicity. Molecules can be toxic to the kidney in a variety of ways, including having deleterious effects on the vasculature, glomerular cells, and tubular epithelium. While many drugs affect renal function, in this review we outline some commonly used models where the molecules are directly toxic to intrinsic renal cells. In most cases, outcomes are directly related to the period of insult. Single exposure to a nephrotoxin results in a model of acute renal disease, with resolution and regeneration. Conversely, repeat exposure models the chronicity seen in progressive renal disease.

5.1

Podocyte Toxins

The nephrotoxic properties of the adriamycin, and the puromycin fragment puromycin aminonucleoside (PAN), are frequently used to model nephrotic syndrome.

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5.1.1

T.D. Hewitson

Adriamycin

Adriamycin is a commonly used antineoplastic antibiotic that inhibits DNA replication. The clinical use of adriamycin is however associated with nephrotic syndrome characterized by heavy proteinuria, albuminuria, hypoalbuminaemia, and hyperlipidaemia (14, 15). Both rat (16, 17) and mouse (15) models of adriamycin nephropathy are well characterized. The principal site of action is the podocyte with ultrastructural studies showing foot process fusion in glomerular epithelial cells. The model is progressive with focal glomerulosclerosis, interstitial infiltration of macrophages and T-lymphocytes, and after about 6 weeks, interstitial fibrosis (15). BalbC but not C57Bl mouse stains are susceptible to a single intravenous (IV) injection of 6–12 mg/kg (15). However, even BalbC mice display a wide range of outcomes with small changes in dose (15), highlighting the necessity to undertake pilot studies.

5.1.2

PAN Nephrosis

In rats, administration of PAN results in nephrotic range proteinuria within days. Again the underlying pathophysiology is a toxic insult to glomerular podocytes. With a single injection, effacement of foot processes and proteinuria resolves (18), although glomerulosclerosis has been seen when the animals are followed for several months (2). Repeat injections result in rapid fibrotic lesions involving both the glomerulus and interstitium (19). The acute model has been used to study the mechanisms of proteinuria, and in the case of repeat injections, to study the pathogenesis of glomerulonephritis and interstitial fibrosis. Mice are not susceptible to PAN.

5.2

Tubular Toxins

5.2.1

Cisplatin

While cisplatin is a highly effective anti-tumor agent, approximately one third of patients develop cytotoxic effects (20). Acute renal failure is inducible in rats or mice by a single intraperitoneal (IP) injection of 6–20 mg cisplatin per kilogram body weight (4, 21). Direct tubular nephrotoxicity is the predominant pathophysiologic process, with cisplatin being absorbed and accumulated in proximal tubule cells (especially the S3 segment). Immediate consequences include apoptosis and necrosis of epithelial cells. Recent reports also indicate that tubular expression of inflammatory chemokines and cytokines has an important role (22). The model is simple and reproducible with pathology and tubular dysfunction comparable to that seen in humans. Repeat injections in the rat result in inflammatory and fibrotic processes (23).

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The declining use of cisplatin however means that its relevance is more as a generic model than a clinical analogue.

5.2.2

Mercuric Chloride

Exposure to mercury is an environmental cause of heavy metal-induced renal toxicity in humans. Injection of rats with 100 µmol/kg of methyl mercury chloride results in acute proximal tubule injury within 2–3 days, followed by a regenerative process (24). Stain and species specific differences have been attributed to the distribution of apical γ-glutamyltranspeptidase in the proximal tubule (25).

5.2.3

Gentamicin

Aminoglycoside antibiotics act on bacteria by penetrating bacterial membranes and binding to the ribosome to prevent protein synthesis. Despite improvements in prescription practices, nephrotoxicity remains a major side effect, the occurrence of which is directly proportional with duration of treatment (26). Proximal but not distal tubules concentrate aminoglycosides and are the site of nephrotoxicity. How aminoglycosides damage tubules is far from certain, but seems to involve lysosyme and mitochondrial dysfunction rather than alterations of protein synthesis (26). A model of gentamicin nephrotoxicity is established in the rat. Animals are given IP gentamicin at a dose generally ranging from 100 to 200 mg/kg body weight for 3–6 consecutive days (27). The model is reversible, although the dose required in animals is far higher than that used in humans. The model is highly relevant clinically and displays myeloid bodies in proximal tubule cells, a characteristic morphologic feature of gentamicin exposure in humans.

5.3

Tubular Obstruction

5.3.1

Adenine

Long-term feeding of adenine (vitamin B4), a major component of DNA and RNA, produces histopathological abnormalities consistent with progressive kidney disease. Although the underlying cause of renal disease is unknown, it is speculated that byproducts formed in the metabolism of adenine are responsible. Orally administered adenine is metabolized to 2,8-dihydroxyadenine, which precipitates and crystallizes in proximal tubular epithelia, accumulation of crystals causing degeneration of tubules. Standard rat chow supplemented with 0.75% adenine is the established model. At this dose, loss of renal function is reversible if feeding is ceased within 2 weeks. Feeding for 4–6 weeks results in interstitial inflammatory infiltration and fibrosis, dilation of tubules, and calcification of tubular basement

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membranes. The model has been described in the rat (28), with one report in the mouse (29).

5.3.2

Folic Acid

Folic acid crystals rapidly appear in tubules after IP administration of folic acid in mice (24, 30). The consequences are acute tubular necrosis and patchy interstitial fibrosis (30). It has often been assumed that the formation of crystals obstructs tubules. A modification of the model uses co-administration of sodium bicarbonate to increase the alkalinity of the urine (24). Although this decreases crystal formation, tubular lesions still occur, suggesting that folic acid also has a direct toxic effect (30).

6

Immune-Mediated Models

Experimental animal models have given valuable insights into the pathophysiology and underlying immunological mechanisms of renal disease in general and glomerular disease in particular.

6.1

Thy-1 Nephritis

Mesangial cell proliferation and expansion of extracellular matrix are important components of the pathophysiological changes in various glomerular diseases. Anti-thymocyte serum (ATS)-induced rat nephritis is recognised as an experimental model of mesangioproliferative glomerulonephritis (MsPGN). The model is often referred to as Thy-1 nephritis (31) as ATS contains antibodies against Thy-1, an antigen originally found on thymocytes and later identified on rat glomerular mesangial cells. To induce Thy-1 nephritis, male rats are intravenously administered a rabbit ATS or a mouse anti-Thy 1 monoclonal antibody (e.g. commercially available OX-7 antibodies at a dose of 1 mg/kg body weight) by a single injection through a tail vein. Thy-1 nephritis is characterized initially by mesangiolysis due to necrosis and fibrin deposition with monocyte/macrophage infiltration (32). Nephritis is followed by subsequent mesangial cell proliferation and mesangial matrix expansion (Fig. 4.2), similar to that seen in human variants of MsPGN such as immunoglobulin A (IgA) nephropathy and Henoch–Schönlein purpura nephritis. However, Thy-1 nephritis only partially models human MsPGN, as there is no evidence of IgA deposits, the defining feature of IgA nephropathy. While extracapillary crescent formation is not diffusely found, some crescent formation accompanies Thy-1 nephritis (33), resembling active human MsPGN.

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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 (see Color Plate 3)

The mesangial cell proliferation peaks at about 1 week in the original Thy-1 nephritis model, and spontaneous repair of the glomerular tufts then occurs, together with the formation of capillary network and a return to normal rates of cell proliferation. Mesangial cell apoptosis is the major mechanism for resolution of glomerular hypercellularity (34). A combination of Thy-1 injection and uninephrectomy has been used to produce a progressive model, characterised by the development of glomerulosclerosis (35), perhaps more in keeping with progressive MsPGN in humans.

6.2

Anti-Glomerular Basement Membrane Nephritis

Human anti-glomerular basement membrane (GBM) nephritis presents a clinical course of rapidly progressive glomerulonephritis with diffuse crescent formation. During progression, breaks in the glomerular capillary basement membrane and tissue factor expression may contribute to the fibrin precipitation in Bowman’s space, and thus promote extracapillary cellular proliferation with features of delayed-type hypersensitivity, including upregulated T cell function (36). This type of nephritis overlaps with lung haemorrhage in Goodpasture’s disease, involving the production of autoantibodies to the basement membranes of glomeruli and alveoli (37). Principal targets for anti-GBM antibodies are the NC1 domains of the alpha-3 chain of type IV collagen (38). A number of animal models of anti-GBM nephritis have been developed. One is induced by the immunization of animals with either homologous or heterologous GBM in complete Freund’s adjuvant (CFA) (39). Originally described in sheep, the

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Steblay nephritis model (40) is also easily induced in rabbits, less reliably in rats. An alternative model, first described in the 1930s, is passively induced by the IV administration of heterologous anti-GBM antibodies (41) (Fig. 4.3). Anti-GBM nephritis can be induced in both mice and rats, although great variation in susceptibility among strains has been reported. Many of the most common mouse strains are not readily susceptible to anti-GBM sera (Table 4.1), suggesting that renal susceptibility to immune nephritis may be genetically programmed. The less susceptible strains may develop more severe immune nephritis when the dose of anti-GBM serum is increased (42). The genetic background is thus an important consideration in designing experiments with gene knockout mice (43). In a similar manner, the Wistar Kyoto (WKY) strain of rats is highly susceptible, with more obvious crescentic glomerulonephritis than Lewis (LEW) rats (44). To prepare rabbit anti-GBM antiserum against mice in the passive induction model, glomeruli from the mouse renal cortex are isolated through a series of sieves and sonicated. The GBM is collected by centrifugation, emulsified with CFA, and administered to rabbits by repeated immunization (43). For the induction of anti-GBM nephritis, mice are intraperitoneally injected with rabbit IgG emulsified with CFA. Five days later they are given an injection with anti-GBM antiserum or purified

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

Table 4.1 Relative susceptibility of various mouse strains to anti-GBM reactive sera Susceptible

Nonsusceptible

NZW, BUB/BnJ, DBA/1J, and 129/svJ

AKR/J, C3H/HeJ, DBA/2J, MRL/MpJ, NOD/LtJ, P/J, SJL/J, and SWR/J

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anti-GBM immunoglobulin into the tail vein (43). Severe proliferative glomerulonephritis with crescent formation and proteinuria develop within 2 to 3 weeks.

7

Metabolic Models

Diabetic nephropathy is now the single most common cause of kidney failure. In the USA for instance it accounts for over 25% of patients commencing dialysis with end stage renal failure and there is a worldwide increase in prevalence. Diabetes causes unique changes in kidney structure, underlying the importance of animal models, all of which depend on persistent hyperglycaemia. Mouse models of diabetes have recently been reviewed in detail (45, 46).

7.1

Models of Type 1 Diabetes

In type 1 diabetes these is an absolute deficiency of insulin production. A number of toxins can directly destroy pancreatic islet cells and induce insulin-dependent diabetes mellitus. This property has been exploited to develop experimental models of type 1 diabetes in a number of animals (Figs. 4.4 and 4.5). The most widely used model is streptozotocin (STZ)-induced β-cell destruction. STZ is a nitrosourea derivative that was first isolated from Streptomyces achromogenes.

Fig. 4.4 Increased glomerular staining for collagen IV in a mouse model of streptozotocin (STZ) diabetes, 16 weeks after STZ injection

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Fig. 4.5 Glomerular sclerosis and interstitial fibrosis in a uninephrectomized rat model of STZ diabetes, 26 weeks after STZ injection. Silver-Masson’s trichrome-stained section

Structural similarity between glucose and STZ results in STZ being transported into pancreatic β-cells by GLUT2, the low affinity glucose transporter. The destruction of the pancreatic β-cells is caused by direct DNA damage and nicotinamide adenine dinucleotide depletion. STZ sensitivity is dependent on a number of factors including GLUT2 expression by pancreatic β-cells, with significant variation in both inter- and intra-species sensitivity to the drug (47) (Fig. 4.4). Unfortunately, GLUT2 is also expressed by renal and hepatic cells. The cytotoxicity of STZ is thus not restricted to pancreatic β-cells but also affects kidney and liver cells, hence the need for carefully controlled experimental design. In rats, diabetes can be induced by a single tail vein injection of STZ in 0.1 M citrate buffer at a dose of 55 mg/kg. Mice are more resistant and are usually injected IP on consecutive days at 100–125 mg/kg. Inter-animal variation is considerable (Fig. 4.6), even with inbred strains, hence many investigators are forced to discard animals that do not become sufficiently hyperglycaemic. Considerable batch variation can exist in STZ, so many investigators avoid using multiple batches in a single study. Self-evidently, STZ is highly toxic and requires great care in handling. It is also unstable when wet so should be stored under desiccated conditions. The STZ model is therefore one in which pilot studies to optimize hyperglycemia are required. In each animal the development of diabetes is monitored by consecutive measurement of blood glucose concentrations, with diabetes usually defined as a blood glucose concentration ≥16 mmol/L. Those animals not achieving this within 2 weeks of STZ injection are either culled or used as technical controls. Measurement of glycated hemoglobin (HbA1c) provides a more useful measure of longer term hyperglycemia than individual blood glucose levels per se. However, it should be remembered that the relatively short half-life of red cells in rodents means (48) that HbA1c levels are often less than those seen in diabetic humans.

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Fig. 4.6 Relative susceptibility of various mouse strains to STZ. Reproduced from ref. (47) with permission of The American Physiological Society

Unfortunately experimental models of diabetes only reproduce some of the pathophysiology of human diabetic nephropathy (3). A number of accelerated models have been developed in an attempt to more closely mimic the human condition, including uninephrectomized or hypertensive animals. The transgenic (mRen-2)27 rat over-expresses the murine renin gene, with elevated angiotensin II activity (49, 50). STZ administration to the mRen-2 rat is an established rodent model of diabetic nephropathy and cardiomyopathy (49, 50), the major advantage being that it does develop functional changes and structural pathology closely mimicking that seen in advanced human diabetic nephropathy (49) and cardiomyopathy (51). Rats are made diabetic by tail-vein injection of 55 mg/kg STZ diluted in 0.1 M citrate buffer (pH 4.5) (47). In a similar manner, spontaneously hypertensive rats (SHR) made diabetic with STZ replicate the confounding hypertension seen in diabetic nephropathy. However despite a similar rise in blood pressure, unlike the diabetic mRen-2 rat, these animals do not usually develop progressive renal failure. As in other models, uninephrectomy may also be used to exacerbate pathology (Fig. 4.5).

7.2

Models of Obesity and Type 2 Diabetes

Type 2 diabetes accounts for the vast majority of patients with diabetic nephropathy. A combination of obesity and insulin resistance are major components of type 2 diabetes in humans, since insulin usually continues to be produced by pancreatic islet cells. It thus has considerable metabolic differences from type 1 diabetes, accordingly requiring animal models in which insulin resistance is present. Animal models of obesity have provided valuable insights into the human condition (3). A high-fat diet is commonly used to induce obesity and insulin resistance in C57BL6 mice (45).

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Much about type 2 diabetes has been learnt from models with single gene mutations. Mutations of the leptin receptor have resulted in the db/db mouse. The db/db mouse develops hyperglycaemia, glomerular hypertrophy, and sclerosis with age (52). The mutant was originally recognized on the C57BLKS/J strain but has been backcrossed to a pure C57BL/6 strain. The pathophysiology of db/db on the C57BLKS/J mice is more severe than that seen with C57BL/6 mice, where plasma glucose seems to normalize with age (45). Mutations of the leptin receptor are however a rare cause of type 2 diabetes in humans, and the direct relevance to the human condition may thus be argued. Obesity and hyperlipidemia develop at an early age in Zucker rats. In addition, obese Zucker rats are characterized by mild glucose intolerance and peripheral insulin resistance similar to that found in humans with type 2 diabetes. These abnormalities precede the development of albuminuria and glomerular injury. Lean litter mates have normal serum lipids and normal renal structure and function. The Zucker rat is readily available commercially, and has been extensively studied in the past (53).

8

Concluding Remarks

Although an animal model can never be a perfect match for human disease (2), adequate, although not yet perfect, animal models have provided valuable insights into the mechanisms of renal disease. The current explosion of techniques to induce genetic modifications in the mouse, and more recently the rat, offer exciting prospects for dissecting individual genes and their related effects in renal disease (24, 54). Concurrently however, adopting models generated in other species or strains can be challenging in these expensive and sometimes fragile animals.

References 1. De Heer, E. and Bruijn, J. A. (1996) Advantages and shortcomings of experimental models of immune-mediated glomerular and tubulointerstitial diseases. Exp. Nephrol. 4, 193–200. 2. Furness, P. N. and Harris, K. (1994) An evaluation of experimental models of glomerulonephritis. Int. J. Exp. Pathol. 75, 9–22. 3. Rees, D. A. and Alcolado, J. C. (2005) Animal models of diabetes mellitus. Diabet. Med. 22, 359–370. 4. Wei, Q., Wang, M. H., and Dong, Z. (2005) Differential gender differences in ischemic and nephrotoxic acute renal failure. Am. J. Nephrol. 25, 491–499. 5. Hewitson, T. D., Mookerjee, I., Masterson, R., Zhao, C., Tregear, G. W., Becker, G. J., and Samuel, C. S. (2007) Endogenous relaxin is a naturally occurring modulator of experimental renal tubulointerstitial fibrosis. Endocrinology 148, 660–669. 6. Chevalier, R. L. (2006) Obstructive nephropathy: towards biomarker discovery and gene therapy. Nat. Clin. Pract. Nephrol. 2, 157–168. 7. Haralambous-Gasser, A., Chan, D., Walker, R. G., Powell, H. R., Becker, G. J., and Jones, C. L. (1993) Collagen studies in newborn rat kidneys with incomplete ureteric obstruction. Kidney Int. 44, 593–605.

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8. Cochrane, A. L., Kett, M. M., Samuel, C. S., Campanale, N. V., Anderson, W. P., Hume, D. A., Little, M. H., Bertram, J. F., and Ricardo, S. D. (2005) Renal structural and functional repair in a mouse model of reversal of ureteral obstruction. J. Am. Soc. Nephrol. 16, 3623–3630. 9. Strauch, M. and Gretz, N. (1988) Animal models to induce renal failure: A historical survey. Contrib. Nephrol. 60, 1–8. 10. Eddy, A. A., Liu, E., and McCullock, L. (1998) Interstitial fibrosis in hypercholesterolemic rats: Role of oxidation, matrix synthesis, and proteolytic cascades. Kidney Int. 53, 1182–1189. 11. Molitoris, B. A. (1998) Ischemic acute renal failure: exciting times at our fingertips. Curr. Opin. Nephrol. Hypertens. 7, 405–406. 12. Azuma, H., Nadeau, K., Takada, M., and Tilney, N. L. (1997) Initial ischemia/reperfusion injury influences late functional and structural changes in the kidney. Transplant Proc. 29, 1528–1529. 13. Forbes, J. M., Jandeleit-Dahm, K., Allen, T. J., Hewitson, T. D., Becker, G. J., and Jones, C. L. (2001) Endothelin and endothelin A/B receptors are increased after ischaemic acute renal failure. Exp. Nephrol. 9, 309–316. 14. Bertani, T., Rocchi, G., Sachi, G., Mecca, G., and Remuzzi, G. (1986) Adriamycin-induced glomerulosclerosis in the rat. Am. J. Kid. Dis. 7, 12–19. 15. Wang, Y., Wang, Y. P, Tay, Y.-C., and Harris, D. C. H. (2000) Progressive adriamycin nephropathy in mice: Sequence of histologic and immunohistochemical events. Kidney Int. 58, 1797–1804. 16. Okuda, S., Oh, Y., Tsuruda, H., Onoyama, K., Fujimi, S., and Fujishima, M. (1986) Adriamycin-induced nephropathy as a model of chronic progressive glomerular disease. Kidney Int. 29, 502–510. 17. Javaid, B., Olson, J. L., and Meyer, T. W. (2001) Glomerular injury and tubular loss in adriamycin nephrosis. J Am Soc Nephrol 12, 1391–1400. 18. Jones, C. L., Buch, S., Post, M., McCulloch, L., Liu, E., and Eddy, A. A. (1992) Renal extracellular matrix accumulation in acute puromycin aminonucleoside nephrosis in rats. Am. J. Pathol. 141, 1381–1396. 19. Jones, C. L., Buch, S., Post, M., McCulloch, L., Liu, E., and Eddy, A. A. (1991) Pathogenesis of interstitial fibrosis in chronic purine aminonucleoside nephrosis. Kidney Int. 40, 1020–1031. 20. Razzaque, M. S. (2007) Cisplatin nephropathy: is cytotoxicity avoidable? Nephrol. Dial. Transplant. 22, 2112–2116. 21. Deng, J., Kohda, Y., Chiao, H., Wang, Y., Hu, X., Hewitt, S. M., Miyaji, T., McLeroy, P., Nibhanupudy, B., Li, S., and Star, R. A. (2001) Interleukin-10 inhibits ischemic and cisplatininduced acute renal injury. Kidney Int. 60, 2118–2128. 22. Lee, S., Kim, W., Moon, S. O., Sung, M. J., Kim, D. H., Kang, K. P., Jang, Y. B., Lee, J. E., Jang, K. Y., and Park, S. K. (2006) Rosiglitazone ameliorates cisplatin-induced renal injury in mice. Nephrol. Dial. Transplant. 21, 2096–2105. 23. Yamate, J., Ishida, A., Tsujino, K., Tatsumi, M., Nakatsuji, S., Kuwamura, M., Kotani, T., and Sakuma, S. (1996) Immunohistochemical study of rat renal interstitial fibrosis induced by repeated injection of cisplatin, with special reference to the kinetics of macrophages and myofibroblasts. Toxicol. Pathol. 24, 199–206. 24. Anders, H. and Schlondorff, D. (2000) Murine models of renal disease: possibilities and problems in studies using mutant mice. Exp. Nephrol. 8, 181–193. 25. Tanaka-Kagawa, T., Suzuki, M., Naganuma, A., Yamanka, N., and Imura, N. (1998) Strain difference in sensitivity of mice to renal toxicity of inorganic mercury. J. Phramacol. Exp. Ther. 285, 335–341. 26. Palmer, B.F., and Henrich, W.L. (2004) Toxic nephropathy. In: Brenner and Reactor’s The Kidney (7th edn.), Brenner BM. Saunders, Philadelphia, PA, 1625–1658. 27. Spiegel, D. M., Shanley, P. F., and Molitoris, B. A. (1990) Mild ischemia predisposes the S3 segment to gentamicin toxicity. Kidney Int 38, 459–464.

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28. Tamaki, K., Yuan, Q., Ohkawa, H., Imazeki, I., Moriguchi, Y., Imai, N., Sasaki, S., Takeda, K., and Fukagawa, M. (2006) Severe hyperparathyroidism with bone anormalities and metastatic calcification in rats with adenine-induced uraemia. Nephrol Dial Transplant 21, 651–659. 29. Minami, T., Nakagawa, H., Ichii, M., Kadota, E., and Okazaki, Y. (1994) Nephrotoxicity induced by a single dose of adenine: effects of 4-aminopyrazolo [3,4-d] pyrimidine and allopurinol. Biol. Pharm. Bull. 17, 201–206. 30. Yuan, H. T., Li, X. Z., Pitera, J. E., Long, D. A., and Woolf, A. S. (2003) Peritubular capillary loss after mouse acute nephrotoxicity correlates with down-regulation of vascular endothelial growth factor-A and hypoxia-inducible factor-1 alpha. Am. J. Pathol. 163, 2289–2301. 31. Ishizaki, M., Masuda, Y., Fukuda, Y., Sugisaki, Y., Yamanaka, N., and Masugi, Y. (1986) Experimental mesangioproliferative glomerulonephritis in rats induced by intravenous administration of anti-thymocyte serum. Acta Pathol. Jpn. 36, 1191–1203. 32. Mosley, K., Collar, J., and Cattell V. (2000) Mesangial cell necrosis in Thy 1 glomerulonephritis—an ultrastructural study. Virchows Arch 436, 567–573. 33. Liu, N., Makino, T., Nogaki, F., Kusano, H., Suyama, K., Muso, E., Honda, G., Kita, T., and Ono, T. (2004) Coagulation in the mesangial area promotes ECM accumulation through factor V expression in MsPGN in rats. Am. J. Physiol. Renal Physiol. 287, F612–F620. 34. Baker, A. J., Mooney, A., Hughes, J., Lombardi, D., Johnson, R. J., and 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. 35. Wada, Y., Morioka, T., Oyanagi-Tanaka, Y., Yao, J., Suzuki, Y., Gejyo, F., Arakawa, M., and Oite, T. (2002) Impairment of vascular regeneration precedes progressive glomerulosclerosis in anti-Thy 1 glomerulonephritis. Kidney Int. 61, 432–443. 36. Erlich, J. H., Holdsworth, S. R., and Tipping, P. G. (1997) Tissue factor initiates glomerular fibrin deposition and promotes major histocompatibility complex class II expression in crescentic glomerulonephritis. Am. J. Pathol. 150, 873–880. 37. Pusey, C. D. (2003) Anti-glomerular basement membrane disease. Kidney Int. 64, 1535–1550. 38. Hellmark, T., Johansson, C., and Wieslander, J. (1994) Characterization of anti-GBM antibodies involved in Goodpasture’s syndrome. Kidney Int. 46, 823–829. 39. Reynolds, J., Mavromatidis, K., Cashman, S. J., Evans, D. J., and Pusey, C. D. (1998) Experimental autoimmune glomerulonephritis (EAG) induced by homologous and heterologous glomerular basement membrane in two substrains of Wistar-Kyoto rat. Nephrol. Dial. Transplant. 13, 44–52. 40. Steblay, R. W. (1962) Glomerulonephritis induced in sheep by injections of heterologous glomerular basement membrane and Freund’s complete adjuvant. J. Exp. Med. 116, 253–272. 41. De Vriese, A. S., Endlich, K., Elger, M., Lameire, N. H., Atkins, R. C., Lan, H. Y., Rupin, A., Kriz, W., and Steinhausen, M. W. (1999) The role of selectins in glomerular leukocyte recruitment in rat anti-glomerular basement membrane glomerulonephritis. J. Am. Soc. Nephrol. 10, 2510–2517. 42. Xie, C., Sharma, R., Wang, H., Zhou, X. J., and Mohan, C. (2004) Strain distribution pattern of susceptibility to immune-mediated nephritis. J. Immunol. 172, 5047–5055. 43. Hisada, Y., Sugaya, T., Yamanouchi, M., Uchida, H., Fujimura, H., Sakurai, H., Fukamizu, A., and Murakami, K. (1999) Angiotensin II plays a pathogenic role in immune-mediated renal injury in mice. J. Clin. Invest. 103, 627–635. 44. Reynolds, J., Albouainain, A., Duda, M. A., Evans, D. J., and Pusey, C. D. (2006) Strain susceptibility to active induction and passive transfer of experimental autoimmune glomerulonephritis in the rat. Nephrol. Dial. Transplant 21, 3398–3408. 45. Breyer, M. D., Bottinger, E., Brosius, IIIF. C., Coffman, T. M., Harris, R. C., Heilig, C. W., and Sharma, K. (2005) Mouse models of diabetic nephropathy. J. Am. Soc. Nephrol. 16, 27–45. 46. Tesch, G. H. and Nikolic-Paterson, D. J. (2006) Recent insights into experimental mouse models of diabetic nephropathy. Nephron Exp. Nephrol. 104, e57–e62.

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47. Gurley, S. B., Clare, S. E., Snow, K. P., Hu, A., Meyer, T. W., and Coffman, T. M. (2006) Impact of genetic background on nephropathy in diabetic mice. Am. J. Physiol. Renal Physiol. 290, F214–F222. 48. Magnani, M., Rossi, L., Stocchi, V., Cucchiarini, L., Piacentini, G., and Fornaini, G. (1988) Effect of age on some properties of mice erythrocytes. Mech. Ageing Dev. 42, 37–47. 49. Kelly, D. J., Wilkinson-Berka, J. L., Allen, T. J., Cooper, M. E., and Skinner, S. L. (1998) A new model of diabetic nephropathy with progressive renal impairment in the transgenic (mRen-2)27 rat (TGR). Kidney Int. 54, 343–352. 50. Mifsud, S. A., Skinner, S. L., Cooper, M. E., Kelly, D. J., and Wilkinson-Berka, J. L. (2002) Effects of low-dose and early versus late perindopril treatment on the progression of severe diabetic nephropathy in (mREN-2)27 rats. J. Am. Soc. Nephrol. 13, 684–692. 51. Martin, J., Kelly, D. J., Mifsud, S. A., Zhang, Y., Cox, A. J., See, F., Krum, H., WilkinsonBerka, J., and Gilbert, R. E. (2005) Tranilast attenuates cardiac matrix deposition in experimental diabetes: role of transforming growth factor-beta. Cardiovasc. Res. 65, 694–701. 52. Chow, F. Y., Nikolic-Paterson, D. J., Ozols, E., Atkins, R. C., and Tesch, G. H. (2005) Intercellular adhesion molecule-1 deficiency is protective against nephropathy in type 2 diabetic db/db mice. J. Am. Soc. Nephrol. 16, 1711–1722. 53. Stevenson, F. T., Kaysen, G. A. (1999) Hyperlipidemia and renal disease: the use of animal models in understanding pathophysiology and approaches to treatment. Wien Klin Wochenschr. 111, 307–314. 54. Bascands, J. L. and Schanstra, J. P. (2005) Obstructive nephropathy Insights from genetically engineered animals. Kidney Int. 68, 925–937.

Chapter 5

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

Abstract Glomerular filtration rate (GFR) is an important index of renal function and routinely used in patient care and basic research to evaluate progression of renal diseases or test the efficacy of novel therapeutic strategies. Determination of GFR in mouse models has been mostly practiced in anesthetized animals, which is not suitable for serial monitoring of GFR in the individual mouse. In this chapter, we outline two approaches for determining GFR in conscious mice including 1) determination of urinary excretion of fluoresceinlabelled inulin (FITC–inulin), and 2) determination of plasma FITC–inulin decay following a single bolus injection. The GFR values determined using these two methods are comparable. The sensitivity of the methods in reflecting renal function was validated in nephrectomized mice and early stage diabetic mice. The effects of inbred mouse genetic background on GFR values are also discussed in this chapter. Keywords Hemodynamics, Glomerular filtration rate, Nephrectomy, Diabetes mellitus, Salts

1

Introduction

Chronic kidney disease (CKD) has become epidemic, affecting over 10% of Americans (1, 2). Mouse models resembling human kidney disease provide important tools that should facilitate our understanding of the pathogenesis of CKD and the testing of therapeutic potential of drugs. Chronic renal disease is characterized by a progressive decline in glomerular filtration rate (GFR). Thus, determination of GFR is an important practice in both patient care and basic research using rodent models. Of the methods which have been used to estimate GFR, inulin clearance is considered the gold standard for GFR determination (3–5). Inulin is a polysaccharide that is filtered freely through glomeruli without reabsorption or secretion by the renal tubules, thus its clearance from the kidney is governed by glomerular filtration capacity. The measurement of From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_5, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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Inulin clearance in mice has typically been conducted in anesthetized animals, however, this approach is not suitable for studies requiring repeated determination of GFR in the same mouse. Furthermore, recent studies suggest the GFR values obtained in conscious rodents might be different from those found in anesthetized animals (6, 7). In this chapter, we outline two approaches for determining GFR in conscious mice: firstly measuring the urinary excretion rate of FITC–inulin using timed urine collections, and secondly, based on the plasma clearance kinetics after a single bolus injection of FITC–inulin. The GFR values determined using these two approaches were comparable in conscious mice (8).

2 2.1

Materials General

1. Fluorescein isothiocyanate (FITC)–inulin (Sigma–Aldrich, St. Louis, MO, USA; cat no F3272). 2. Dialysis membrane with a 1,000-Da cutoff (Spectra/Pro 6; Spectrum Laboratories, Rancho Dominguez, CA, USA). Rinse the membrane in 0.9% NaCl for approximately 30 min prior to use. 3. Dialysis membrane clamps (Universal Closure; Spectrum Laboratories). 4. Aluminium foil. 5. 500 mM HEPES buffer (pH 7.4): dissolve HEPES (FW 238.3; Sigma–Aldrich) in distilled water, and adjust to pH 7.4. 6. Fluoroscan Ascent FL (FIN-00811; Labsystems, Helsinki, Finland). 7. Heparinized capillary tubes (Microvette CB300; Sarstedt, Nümbrecht, Germany). 8. 0.22-µm syringe filter (Costar, Corning, NY, USA). 9. Mouse restraint, e.g. a 50-mL conical centrifuge tube with breathing holes drilled in the end. 10. 1.5-mL microfuge tubes (Eppendorf, Westbury, NY).

2.2

Urinary Inulin Clearance Method

1. Micro-osmotic pump: Alzet™, model 1007D (Durect Corporation, Cupertino, CA, USA), with a release rate of 0.5 µL/h for 7 days. 2. Anesthetic: ketamine and xylazine. 3. Surgical instruments and suture. 4. Surgical iodine, ethanol, and gauze. 5. Metabolic cage (Braintree Scientific, Braintree, MA, USA).

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2.3

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Single-Bolus Injection Method

1. Isoflurane (Baxter Pharmaceutical Products, Deerfield, IL, USA). 2. Normal mouse serum. 3. GraphPad Prism™ (GraphPad Software, San Diego, CA, USA).

3

Methods

3.1

GFR Measurement Based on Urinary Inulin Clearance

This method utilizes intraperitoneally implanted micro-osmotic pumps that continuously release FITC–inulin to achieve a steady level of plasma inulin concentration. GFR is determined based on urinary FITC–inulin excretion rate. This method requires housing mice in metabolic cages to collect urine over 24 h.

3.1.1

Preparation and Dialysis of FITC–Inulin

1. 5% (w/v) FITC–inulin solution is made by dissolving FITC–inulin in 0.9% NaCl. This requires heating the solution in boiling water to fully dissolve the inulin. 2. To remove residual unbound FITC, the FITC–inulin solution (5%) is placed in a 1,000-Da cutoff dialysis membrane (Spectra/Pro 6; Spectrum Laboratories) and clamped at each end of the membrane using dialysis membrane clips (Spectrum Laboratories). The membrane is then placed in 1,000 mL of 0.9% NaCl with a magnetic stir bar for dialysis overnight at room temperature. To minimize photo bleaching, the container is wrapped with aluminium foil. 3. Before use, filter the dialyzed FITC–inulin through a 0.22-µm filter.

3.1.2

Implantation of Micro-osmotic Pumps

1. Mice are anesthetized with ketamine (50 µg/g body wt) and xylazine (5 µg/g body weight) and placed on a warming pad. The midline fur is shaved, and the skin is sterilized with iodine and ethanol. 2. Two micro-osmotic pumps (Alzet™, model 1007D), each filled with ∼100 µL of dialyzed FITC–inulin, are inserted into the peritoneal cavity through a small midline incision (∼0.5 cm) (see Note 1). 3. The incision is closed with sterilized suture.

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Urine Collection Using Metabolic Cage

1. Three days later, following complete recovery from surgery, mice with implanted micro-osmotic pumps are individually housed in metabolic cages. 2. On day 7, urine is collected over 24 h. 3. To harvest the residual fluorescence on the wall, we rinse the cage twice with 10 mL of 500 m M HEPES (pH 7.4) (see Note 2 ) (8) .The fluorescence reading in the rinse solution is measured and added to that from the urine collected.

3.1.4

Blood Collection

To determine blood FITC–inulin concentration in these mice, sample blood via the saphenous vein at the end of the 24 h urine collection. 1. Conscious mice are restrained in a 50-mL centrifuge tube with air holes drilled in the tip. The inner thigh is closely shaved and sterilized using iodine followed by 75% ethanol, revealing the saphenous vein. 2. The vein is punctured using a sterilized 23-gauge syringe needle, and approximately 100 µL of blood is collected using a heparinized capillary tube. 3. Centrifuge the blood at 1,000×g for 10 min using a desktop centrifuge. Transfer 40 µL of plasma to a clear 1.5-mL microfuge tube.

3.1.5

Measurement of FITC–Inulin in Plasma and Urine

It has been demonstrated the pH significantly affects the fluorescence reading of FITC in solution, and the FITC fluorescence readings is linearly correlated with FITC–inulin concentration at pH 7.4 (9), thus we buffer both plasma and urine samples to pH 7.4 using 500 mM HEPES (pH 7.4). 1. To do this, mix 40 µ L of plasma or urine with 10 µL of 500 m M HEPES (pH 7.4) (see Note 3). 2. The titrated samples (total 50 µL) are loaded onto a 96-well plate. 3. Fluorescence is determined with 485-nm excitation and reading at 538-nm emission (Fluoroscan Ascent FL; Labsystems).

3.1.6

Calculation of GFR

A linear relationship between fluorescence reading and inulin concentration is confirmed using a set of serially diluted FITC–inulin solutions. Using fluorescence as an index of inulin concentration in both blood and urine, the GFR is calculated using the equation: GFR = (urinary fluorescence concentration) × (urine volume) / (plasma fluorescence concentration).

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To validate this measurement of GFR, we compared it with that calculated using an additional equation: GFR = (inulin infusion rate) / (steady-state blood inulin concentration), as previously described (10, 11). The infusion rate was estimated based on the manufacturer’s pre-set release rate of the micro-osmotic pump. The GFR values calculated were comparable using these two methods (8).

3.2

Measurement of GFR with a Single Bolus Injection of FITC–Inulin

This method calculates GFR using a two-compartment model (10, 11). This model is widely used to determine clearance rate of a drug or tracer (e.g. inulin) first distributing from blood (central compartment) into the extracellular fluid (peripheral compartment) after single-bolus intravenous injection, followed by its elimination from the central compartment (e.g. via the kidney). In this model, the plasma disappearance of the drug or indicator fits to bi-exponential characteristics. Figure 5.1a represents the plasma disappearance of FITC–inulin in five

Fluorescence (counts ml plasma)

a

7.5

b 6.0

A

4.5

B

Y = Ae -atx + Be -b tx + Plateau

Plateau 3.0

Time

1.5

0 0

25

50

75

100

125

150

175

200

Fig. 5.1 Representative plasma clearance kinetics of FITC–inulin in five healthy male C57BL/6 mice (a). Fluorescence is plotted versus time on an arithmetic scale. The plasma fluorescence was also examined at 180 min post injection in two healthy mice. The inset (b) provides the formula used for non-linear regression curve fitting and the model. For these studies the plateau was set to equal zero. (Reproduced from ref. (8) with permission from The American Physiology Society)

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healthy male C57BL/6J mice, indicating the clearance of inulin in mice fit to the two-compartment model. In the two-compartment model used, depicted in Fig. 5.1b, the initial, rapid decay phase represents redistribution of the tracer from the intravascular compartment to the extracellular fluid. Systemic elimination also occurs, but the distribution process is relatively dominant during this initial phase. During the later phase, slower decay in tracer concentration predominantly reflects systemic clearance from the plasma. At any given time (tX), the plasma concentration of the tracer (Y) equals Ae− tx + Be− tx + Plateau (http://curvefit.com/id205.htm) (10, 11). The clearance can be calculated using the equation: GFR = I/(A/α + B/β ) The “I” is the amount of FITC–inulin delivered by the bolus injection. The A and B are the y-intercept values of the two decay rates, and α and β are the decay constants for the distribution and elimination phases, respectively (10, 11).

3.2.1

Introduction of FITC–Inulin and Blood Collection

1. Dialyzed FITC–inulin (3.74 µL/g body weight) is injected into the tail vein or retroorbitally under light anesthesia induced using isoflurane. The anesthesia lasts approximately 20 s. 2. Approximately 25 µL venous blood is collected at 3, 7, 10, 15, 35, 55, and 75 min after FITC–inulin injection, yielding 10 µL of plasma for the determination of FITC concentration by fluorescence. Care must be taken not to exceed ∼200 µL total blood loss, to minimize volume depletion of the mouse (see Note 4). 3.2.2

Measurement of FITC–Inulin in Plasma

As discussed above, pH significantly affects FITC fluorescence value (9). 1. Buffer the plasma to pH 7.4, by mixing 40 µL of 500 mM HEPES solution (pH 7.4) with 10 µL plasma obtained at each time point and load the titrated samples (total 50 µL) onto a 96-well plate. 2. To do this, mix 4 µL of the FITC–inuli n solution with 156 µL of 500 mM HEPES solution, and then divide the solution equally into four 1.5-mL Eppendorf tubes. In addition to the sample plasma, the fluorescence of the FITC–inulin solution used for bolus injection is determined. 3. 10 µL of mouse plasma (commercially available) is added into each tube (final volume: 50 µL/tube containing 1 µL FITC–inulin). 4. The average fluorescence reading (from 1 µL of the FITC–inulin solution) multiplied by the volume injected into the mouse equals the total amount of fluorescence injected (the “I” in the above-described GFR calculating equation).

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3.2.3

67

Calculation of GFR Based on Plasma FITC–Inulin Decay

The plasma fluorescence data at predetermined time points following the single bolus injection of FITC–inulin is fit to a two-phase exponential decay curve using non-linear regression (GraphPad Prism™, GraphPad Software). Our preliminary studies have shown that the plasma fluorescence level decreases to nearly zero at 180 min post-injection (Fig. 5.1a), thus the plateau is set to zero. The GFR is calculated using the equation GFR = I/ (A/α + B/β) as described above. In the Prism GraphPad™ program, the “SPAN2” is equivalent to “A”, the “SPAN1” to “B”, “K2” equals “α”, and “K1” equals“β”. The units of A and B are fluorescence counts/10 µL, and α and β equal 0.7/T1/2 (minutes) (10). The GFR calculated using this single bolus injection method is comparable to that using a micro-osmotic pump.

3.3

Validation that the Single Bolus Injection Method Sensitively Reflects Renal Function

To examine whether the single bolus method reflects renal function, we have determined GFR in mouse models with total or 5/6 nephrectomy, and in diabetic mice. The nephrectomy was conducted in C57BL/6J male mice. The mice were anesthetized using ketamine (50 µg/g body weight) and xylazine (5 µg/g body weight), to achieve the desired depth of anesthesia for approximately 45 min. A 0.5-cm incision parallel to spinal column was cut on each flank of the back under aseptic conditions. For total nephrectomy, both kidneys were removed through these incisions. For 5/6 nephrectomy, the right kidney and upper and lower thirds of the left kidney were removed (12). The incisions were sutured (6.0 thread, Ethicon Inc., Somerville, NJ, USA) and the mice allowed to recover from anesthesia. Determination of GFR was conducted 2 weeks following 5/6 nephrectomy or immediately after full recovery from anesthesia in mice with total nephrectomy. The diabetic mice were generated using C57BL/6J male mice that had received streptozotocin injection intraperitoneally (50 mg/kg, made freshly in 0.1 M citrate buffer, pH 4.5) each day for 5 consecutive days. Fasting levels of blood glucose were examined weekly with a B-Glucose Analyzer. GFR was measured following 5 weeks of documented hyperglycemia. The inulin clearance calculated using the two-compartment model was dramatically decreased in 5/6 nephrectomized mice, and totally lost in mice following the removal of both kidneys, while GFR in early stage diabetic mice was higher than control mice (Fig. 5.2). Furthermore, we have recently found that the GFR determined using the single bolus injection approach is significantly correlated with glomerulosclerosis scores in diabetic mice (unpublished data). These studies support the use of the single-bolus FITC–inulin method in determining GFR in mice.

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Fig 5.2 GFR determined by single bolus injection approach in male C57BL/6 mice following total nephrectomy (n=4), 5/6 nephrectomy (n=5), or early stages of diabetes mellitus (n=4) (a). Nephrectomy abolished or dramatically decreased GFR compared with control (n=6). Attenuated elimination of FITC–inulin fluorescence was also observed in mice with 5/6 nephrectomy (b). In contrast, mice with early diabetes induced by STZ exhibited significantly increased GFR. *P<0.005 compared with control; #P< 0.05 compared with 5/6 nephrectomy. (Reproduced from ref. (8) with permission from The American Physiological Society)

3.4

GFR Values in Mice

Table 5.1 summarizes previously published GFR in mice. It appears that significant variation in GFR exists in mice and this may be due to several factors. Gender difference in GFR has been previously reported in many species including humans and mice (13–16). Mouse strain appears to be another factor influencing the levels of GFR. For example, C57BL/6 mice have previously been found to exhibit a relatively low GFR as compared with other inbred strains (17). GFR obtained using 51 Cr-EDTA elimination kinetics in conscious C57BL/6 male mice was only 57.9% of that in B6D2F1 hybrid (Table 5.1), a difference which could not be explained by difference in body or kidney weight between these strains (17). Other previous studies examined GFR in anesthetized animals. Anesthesia may significantly influence GFR. The values of GFR in conscious rats (7) were only approximately 60% of that obtained in anesthetized rats by the same group (6). The method used to calculate renal clearance may also result in different GFR values. For example, a one-compartment model has been used to estimate GFR (14, 17–19). This model calculates clearance rate of a drug or tracer by considering the body as a single compartment, and assuming the drug is rapidly distributed in the compartment and elimination is first order. This single compartment model has been more recently shown to overestimate GFR in rats (8, 11).

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Table 5.1 Published GFR values in mice Status

GFR (µL/ min) BW (g)

Conscious Conscious Conscious Conscious Conscious Conscious Conscious Conscious Conscious Conscious Conscious Conscious

237 140 330 570 497 388 495 201 407 315 40.8 65

Conscious Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized Anesthetized

175 229 288 294 303 325 252 246 385 275 229 422 460

27.2 22.3 27.5 32.7 36.3 31.4 40.9 27.7 22.5 35.6 19.2

M F M M M F M M M F M&F

C57BL/6J C57BL/6J C57BL/6 B6D2F1 Swiss Webster Swiss Webster Swiss Webster BABL/c BALB/c BALB/c MF1xCD1 BKSmLeprdb/++ BALB/c C57BL/6 C57BL/6 C57BL/6 C57BL/6 C57BL/6 C57BL/6 CD-1 C57x129 C57x129 C57x129 129xBS C57x129

Tracer

Reference

FITC–inulin FITC–inulin 51 Cr-EDTAa 51 Cr-EDTAa 51 Cr-EDTAa 51 Cr-EDTAa FITC–inulin 51 Cr-EDTAa 51 Cr-EDTAa 51 Cr-EDTAa Creatinine Creatinine

(8) (8) (17) (17) (14) (14) (8) (24) (15) (15) (25) (18)

Creatinine (26) FITC–inulin (27) 14 20–25 C-inulin (28) 14 C-inulin (29) 19.4 24.8 Polyfructosan (30) 26.1 FITC–inulin (16) 21.3 FITC–inulin (16) 3 30.5 H-inulin (31) 3 47 H-inulin (32) 3 H-inulin (33) 3 24 H-inulin (34) 31.1 FITC–inulin (35) 125 22.3 F I-iothalamate (36) Summarized data from the published literature. Some of the parameters are not available in the publications. a One-compartment model. (Reproduced from ref. (8) with permission from The American Physiology Society)

3.5

20–25

Gender Strain

M M M M M M F M M M&F F

Creatinine Clearance in Mice

Creatinine clearance is widely used to estimate GFR in humans. Several studies indicate the Jeffe alkaline picrate method significantly overestimates plasma creatinine level in mice (20, 21). Recent studies show a HPLC method is more suitable to determine plasma creatinine concentration in mice, and the creatinine clearance based on plasma and urinary concentration of creatinine determined using HPLC correlates with inulin clearance in mice (21). Additionally, the enzymatic creatininase method coupled with a peroxidase indicator more closely agrees with HPLC methods (22). Others have utilized tandem mass spectrometry to determine creatinine in mice (23).

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Conclusion

GFR can be accurately determined in conscious mice using FITC–inulin. The single bolus injection method is preferable for serial GFR measurement in the same mouse without surgery or housing mice in metabolic cages. The osmotic pump method may be more suitable for studies examining electrolyte metabolism, but because of the surgery required, is less suitable for long-term monitoring.

4

Notes

1. The rationale for using two micro-osmotic pumps is based on pilot studies showing a plasma fluorescence level two to six times higher than background could only be achieved by using two minipumps. 2. For the micro-osmotic pump method, rinsing the cage to collect residual FITC–inulin is an important step for accurately estimating GFR. We found the harvested fluorescence by this step accounts for 26.2±2.2% and 36.7±5.0% of total urinary fluorescence detected in male and female C57BL/6J mice, respectively (8). 3. Pilot studies have indicated that 10 µL of 500 mM HEPES (pH 7.4) is sufficient to buffer 40 µL of mouse plasma or urine to pH 7.4. 4. For single bolus injection approach, a successful blood collection at each time point is critical for the measurement. To preserve the vessel, the pressure applied to stop bleeding after each sampling should be appropriate. Pressing too hard may cause vessel closure and make it difficult to get blood at the next time point.

References 1. Coresh, J., Byrd-Holt, D., Astor, B. C., Briggs, J. P., Eggers, P. W., Lacher, D. A., and Hostetter, T. H. (2005). Chronic kidney disease awareness, prevalence, and trends among U.S. adults, 1999 to 2000. J Am Soc Nephrol 16, 180–188. 2. Hallan, S. I., Coresh, J., Astor, B. C., Asberg, A., Powe ,N. R., Romundstad, S., Hallan, H. A., Lydersen, S., and Holmen, J. (2006). International comparison of the relationship of chronic kidney disease prevalence and ESRD risk. J Am Soc Nephrol 17, 2275–2284. 3. Fischer, P. A., Bogoliuk, C. B., Ramirez, A. J., Sanchez, R. A., and Masnatta, L. D. (2000). A new procedure for evaluation of renal function without urine collection in rat. Kidney Int 58, 1336–1341. 4. Prescott, L. F., Freestone, S., and McAuslane, J. A. (1991). Reassessment of the single intravenous injection method with inulin for measurement of the glomerular filtration rate in man. Clin Sci (Lond) 80, 167–176. 5. Levey, A. S. (1990). Measurement of renal function in chronic renal disease. Kidney Int 38, 167–184. 6. Ahloulay, M., Dechaux, M., Laborde, K., and Bankir, L. (1995). Influence of glucagon on GFR and on urea and electrolyte excretion: direct and indirect effects. Am J Physiol 269, F225–235. 7. Bouby, N., Ahloulay, M., Nsegbe, E., Dechaux, M., Schmitt, F., and Bankir, L. (1996). Vasopressin increases glomerular filtration rate in conscious rats through its antidiuretic action. J Am Soc Nephrol 7, 842–851.

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8. Qi, Z., Whitt, I., Mehta, A., Jin, J., Zhao, M., Harris, R. C., Fogo, A. B., and Breyer, M. D. (2004). Serial determination of glomerular filtration rate in conscious mice using FITC–inulin clearance. Am J Physiol Renal Physiol 286, F590–596. 9. Lorenz, J. N. and Gruenstein, E. (1999). A simple, nonradioactive method for evaluating single-nephron filtration rate using FITC–inulin. Am J Physiol 276, F172–177. 10. Catlin, D.H. (1983). Pharmacokinetics. In: Essentials of Pharmacology, edited by J.A. Bevan and J.H. Thompson. 3rd edn., Harper & Row, Publishers, Inc., Philadelphia. 11. Sturgeon, C., Sam, A. D., 2nd, and Law, W. R. (1998). Rapid determination of glomerular filtration rate by single-bolus inulin: a comparison of estimation analyses. J Appl Physiol 84, 2154–2162. 12. Ma, L. J. and Fogo, A. B. (2003). Model of robust induction of glomerulosclerosis in mice: importance of genetic background. Kidney Int 64, 350–355. 13. Goldfarb, D. A., Matin, S. F., Braun, W. E., Schreiber, M. J., Mastroianni, B., Papajcik, D., Rolin, H. A., Flechner, S., Goormastic, M., and Novick, A. C. (2001). Renal outcome 25 years after donor nephrectomy. J Urol 166, 2043–2047. 14. Hammond, K. A. and Janes, D. N. (1998). The effects of increased protein intake on kidney size and function. J Exp Biol 201 (Pt 13), 2081–2090. 15. Messow, C., Gartner, K., Hackbarth, H., Kangaloo, M., and Lunebrink, L. (1980). Sex differences in kidney morphology and glomerular filtration rate in mice. Contrib Nephrol 19, 51–55. 16. Noonan, W. T. and Banks, R. O. (2000). Renal function and glucose transport in male and female mice with diet-induced type II diabetes mellitus. Proc Soc Exp Biol Med 225, 221–230. 17. Hackbarth, H. and Hackbarth, D. (1981). Genetic analysis of renal function in mice. 1. Glomerular filtration rate and its correlation with body and kidney weight. Lab Anim 15, 267–272. 18. Cohen, M. P., Clements, R. S., Hud, E., Cohen, J. A., and Ziyadeh, F. N. (1996). Evolution of renal function abnormalities in the db/db mouse that parallels the development of human diabetic nephropathy. Exp Nephrol 4, 166–171. 19. Gartner, K. (1978). Glomerular hyperfiltration during the onset of diabetes mellitus in two strains of diabetic mice (c57bl/6j db/db and c57bl/ksj db/db). Diabetologia 15, 59–63. 20. Breyer, M. D., Bottinger, E., Brosius, F. C., 3rd, Coffman, T. M., Harris, R. C., Heilig, C. W., and Sharma, K. (2005). Mouse models of diabetic nephropathy. J Am Soc Nephrol 16, 27–45. 21. Dunn, S. R., Qi, Z., Bottinger, E. P., Breyer, M. D., and Sharma, K. (2004). Utility of endogenous creatinine clearance as a measure of renal function in mice. Kidney Int 65, 1959–1967. 22. Jung, K., Wesslau, C., Priem, F., Schreiber, G., and Zubek, A. (1987). Specific creatinine determination in laboratory animals using the new enzymatic test kit “Creatinine-PAP.” J Clin Chem Clin Biochem 25, 357–361. 23. Takahashi, N., Boysen, G., Li, F., Li, Y., and Swenberg, J. A. (2007). Tandem mass spectrometry measurements of creatinine in mouse plasma and urine for determining glomerular filtration rate. Kidney Int 71, 266–271. 24. Haines, H. and Farmer, J. N. (1991). Glomerular filtration rate and plasma solutes in BALB/c mice infected with Plasmodium berghei. Parasitol Res 77, 411–414. 25. Aizman, R., Asher, C., Fuzesi, M., Latter, H., Lonai, P., Karlish, S. J., and Garty, H. (2002). Generation and phenotypic analysis of CHIF knockout mice. Am J Physiol Renal Physiol 283, F569–577. 26. Park, K. M., Chen, A., and Bonventre, J. V. (2001). Prevention of kidney ischemia/reperfusion-induced functional injury and JNK, p38, and MAPK kinase activation by remote ischemic pretreatment. J Biol Chem 276, 11870–11876. 27. Wang, W., Jittikanont, S., Falk, S. A., Li, P., Feng, L., Gengaro, P. E., Poole, B. D., Bowler, R. P., Day, B. J., Crapo, J. D., and Schrier, R. W. (2003). Interaction among nitric oxide, reactive

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30. 31. 32.

33.

34.

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36.

Z. Qi, M.D. Breyer oxygen species, and antioxidants during endotoxemia-related acute renal failure. Am J Physiol Renal Physiol 284, F532–537. O’Donnell, M. P., Burne, M., Daniels, F., and Rabb, H. (2002). Utility and limitations of serum creatinine as a measure of renal function in experimental renal ischemia-reperfusion injury. Transplantation 73, 1841–1844. Kawada, N., Imai, E., Karber, A., Welch, W. J., and Wilcox, C. S. (2002). A mouse model of angiotensin II slow pressor response: role of oxidative stress. J Am Soc Nephrol 13, 2860–2868. Cervenka, L., Mitchell, K. D., and Navar, L. G. (1999). Renal function in mice: effects of volume expansion and angiotensin II. J Am Soc Nephrol 10, 2631–2636. Luippold, G., Pech, B., Schneider, S., Osswald, H., and Muhlbauer, B. (2002). Age dependency of renal function in CD-1 mice. Am J Physiol Renal Physiol 282, F886–890. Cullen-McEwen, L. A., Kett, M. M., Dowling, J., Anderson, W. P., and Bertram, J. F. (2003). Nephron number, renal function, and arterial pressure in aged GDNF heterozygous mice. Hypertension 41, 335–340. Wulff, P., Vallon, V., Huang, D. Y., Volkl, H., Yu, F., Richter, K., Jansen, M., Schlunz, M., Klingel, K., Loffing, J., Kauselmann, G., Bosl, M. R., Lang, F., and Kuhl, D. (2002). Impaired renal Na(+) retention in the sgk1-knockout mouse. J Clin Invest 110, 1263–1268. Brown, R., Ollerstam, A., Johansson, B., Skott, O., Gebre-Medhin, S., Fredholm, B., and Persson, A. E. (2001). Abolished tubuloglomerular feedback and increased plasma renin in adenosine A1 receptor-deficient mice. Am J Physiol Regul Integr Comp Physiol 281, R1362–1367. Lorenz, J. N., Baird, N. R., Judd, L. M., Noonan, W. T., Andringa, A., Doetschman, T., Manning, P. A., Liu, L. H., Miller, M. L., and Shull, G. E. (2002). Impaired renal NaCl absorption in mice lacking the ROMK potassium channel, a model for type II Bartter’s syndrome. J Biol Chem 277, 37871–37880. Sun, D., Samuelson, L. C., Yang, T., Huang, Y., Paliege, A., Saunders, T., Briggs, J., and Schnermann, J. (2001). Mediation of tubuloglomerular feedback by adenosine: evidence from mice lacking adenosine 1 receptors. Proc Natl Acad Sci U S A 98, 9983–9988.

Chapter 6

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

Abstract Kidney tissue laser capture microdissection (LCM) is of great clinical relevance since genome wide studies on total kidney messenger RNA (mRNA) potentially miss important factors involved in the pathogenesis of the disease in glomeruli and tubules. This technique is readily applicable to study mRNA from isolated glomeruli and tubules of human kidney biopsy material. In this chapter we present a “cook-book” practical approach of utilizing LCM in combination with RNA isolation technique in downstream applications in nephrology. Keywords Glomerular isolation, Tubular dissection, RNA quality and quantity

1

Introduction

Studies of human renal tissue using microarray technology confirmed that a molecular based classification of renal disease is possible (1–4). However, this task is made difficult by the fact that renal disease is characterized by interrelated mechanisms of inflammation, repair, scarring, and atrophy affecting over 20 different renal cell types (4), and by the heterogeneity of the gene expression across kidney compartments (2). While the approach of global screening of gene expression in whole kidney of chronic progressive experimental renal disease and/or human renal biopsies has been highly successful in identifying numerous novel candidate genes that correlate with pathological and/or developmental outcomes (5–7), it falls short of identifying compartmental or cellular culprits of altered candidate gene expression. Laser capture microdissection (LCM) has allowed molecular interrogation of individual cells and structures of frozen biopsy tissues (8–10), and feasibility of renal gene expression analysis of renal tissue isolated by laser microdissection has been validated (11). This chapter describes a method for preparing material for the quantitative analysis of gene expression in different compartments of the kidney. The technique uses LCM to isolate RNA from glomeruli and tubules for subsequent analysis of gene expression with real-time polymerase chain reaction (PCR). By comparing whole From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_6, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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kidney RNA (total RNA) and RNA in captured tissue, useful insights into relative expression can be obtained.

2 2.1 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

2.2

Materials Tissue Collection Cryomold. Liquid nitrogen. 100% ethanol. OCT™ embedding compound (Tissue-Tek™ ; Fisher, Pittsburgh, PA, USA). Freezing microtome (cryostat) (Leica, Wetzlar, Germany). Microslide box. Silica desiccant. Dry ice. HistoGene™ Frozen Section Staining Kit (Arcturus Engineering, Mountain View, CA, USA). RNase Zap™ (Sigma–Aldrich, St. Louis, MO, USA). Plastic slide jars or 50-mL centrifuge tubes (Falcon, BD Biosciences, San Jose, CA, USA). Kimwipes™ (Kimberly-Clark, Roswell, GA, USA). Staining rack.

Whole Kidney (Total) RNA Isolation

RNeasy™ purification kit (Qiagen, Hilden, Germany).

2.3

Laser Capture Microscopy

1. CapSure™ Macro LCM Caps (Arcturus Engineering). 2. Laser capture microscope (Pixcell™; Arcturus Engineering).

2.4

Captured Tissue RNA Isolation and Analysis

1. PicoPure™ RNA isolation kit (Arcturus Engineering). 2. RNase-Free DNase Set (Qiagen). 3. Lab-on-chip technology (Agilent 2100 Bioanalyzer; Agilent Technologies, Palo Alto, CA, USA).

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4. PicoRNA™ chip (Agilent). 5. Freezer (−80°C) for storage.

3

Methods

This section contains detailed protocols for tissue processing, LCM, and RNA isolation.

3.1

Processing and Collection of Kidney Tissue

In summary, kidney tissue (2-mm coronal slices of whole kidney or 5-mm longitudinal slice of kidney biopsy with 16-gauge needle) embedded in OCT™ (Tissue Tek™) is cryosectioned into 7-µm sections at −20°C using a cryomicrotome (Leica). Cryosections are then attached to RNase free glass slides (Arcturus Engineering). The slides should be processed utilizing an RNase-free technique, and stained using a HistoGene™ kit (Arcturus Engineering). The HistoGene stain has been especially developed to stain tissue sections that are going to be used as a source of RNA. It is a fast penetrating stain that provides good contrast by differential staining of nuclei (purple) and cytoplasm (light pink) while preserving RNA integrity. By providing contrast, it enables the operator of a LCM to identify areas of interest for excision. HistoGene™ staining is performed using a modified rapid protocol for LCM, as previously described (8). The slides are defrosted, fixed in 75% ethanol for 1 min, stained with HistoGene™ (Arcturus Engineering), and dehydrated in 75% ethanol, 95% ethanol, 100% ethanol, and xylene. Thirty glomeruli dissected under direct visualization with a laser capture microscope (Pixcell™) are sufficient for downstream analysis (Fig. 6.1) (12). 3.1.1

Specimen Freezing

Wear clean disposable gloves throughout the specimen freezing procedure. Change gloves after each sample. Use clean RNase-free instruments. 1. 2. 3. 4. 5. 6.

Place the cryomold on dry ice. Collect the dissected tissue specimen. Place the tissue specimen in the cold cryomold from step 1. Add OCT™ (Tissue-Tek™) to the cold cryomold until the specimen is covered. Place the bottom of the cryomold in liquid nitrogen. Wait for the OCT™ to turn white, then submerge the cryomold completely until frozen. 7. Store the frozen specimen in the cryomold in a −80°C freezer until further processing.

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Fig. 6.1 Cryosection of single glomerulus of 4-week-old wild-type mice before (a), and after (b) laser capture with laser spots of 15 µm. Recovery of single glomerulus (c) on transfer film (magnification × 40). Slide of the renal cortex (d) after laser microdissection of several glomeruli (magnification × 10)

3.1.2

Slide Preparation

Wear double disposable clean gloves throughout the slide preparation procedure. 1. Precool the cryostat to the temperature recommended by the manufacturer for the specimen you are preparing (–20°C for kidney tissue). 2. Remove and discard the old blade. Wipe down the knife holder and anti-roll plate in the cryostat with RNase Zap™ (Sigma) and 100% ethanol to avoid sample cross-contamination. Do not use the 100% ethanol solution provided in the HistoGene™ Frozen Section Staining Kit for this step. 3. Install a new disposable blade in the cryostat. 4. Place a microslide box containing silica desiccant (Sigma) on dry ice near the cryostat. 5. Move the cryomold containing your specimen from the –80°C freezer or dry icebox to the cryostat. 6. Wait 10 min for your specimen to reach an appropriate cutting temperature. 7. Cut 7-µm sections from your specimen. If you are cutting more than one specimen, repeat step 2 above. 8. Collect each section onto a room-temperature LCM slide and place immediately into a microslide box on dry ice. Do not allow the sections to air dry. 9. Discard slides with folded or wrinkled sections. 10. Store at –80°C for up to 2 weeks.

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77

Staining and Dehydration

Change all solutions in the plastic slide jars between each batch of slides to avoid cross contamination. Do not reuse solutions. Do not transfer solutions back into their original bottles. If you plan to reuse jars, discard all solutions upon completion of the staining and clean the jars. 1. Label seven plastic slide jars or clean 50-mL Falcon tubes as follows: (a) 75% ethanol, (b) distilled water, (c) distilled water, (d) 75% ethanol, (e) 95% ethanol, (f) 100% ethanol, and (g) xylene. 2. Using the LCM Certified RNase-free solutions provided with your HistoGene™ Frozen Section Staining Kit (Arcturus Engineering), fill the labeled plastic slide jars with 25–40 mL of the appropriate solution. 3. Remove two slides from the slide box on dry ice or from the –80°C freezer, and allow to thaw for no more than 30 s. 4. Place the slides in plastic slide jar “a” containing 75% ethanol for 30 s. Use cover glass forceps (first clean the forceps with RNase Zap™) to transfer slides from jar to jar. 5. Transfer the slides to plastic slide jar “b” containing distilled water for 30 s. 6. Place the slides on a Kimwipe™ or a horizontal staining rack. 7. Using an RNase-free pipette tip, apply approximately 100 µL of the HistoGene™ Staining Solution so that it covers the section. Stain for 20 s. 8. Place the slides in plastic slide jar “c” containing distilled water for 30 s. 9. Transfer the slides to plastic slide jar “d” containing 75% ethanol for 30 s. 10. Transfer the slides to plastic slide jar “e” containing 95% ethanol for 30 s. 11. Transfer the slides to plastic slide jar “f ” containing 100% ethanol for 30 s. 12. Transfer the slides in to plastic slide jar “g” containing xylene for 5 min. 13. Place the slides on a Kimwipe™ to dry in the hood for 5 min. 14. Place all slides in a desiccator or slide box containing fresh desiccant. 15. Remove one slide and perform LCM, keeping the remainder in the desiccator until ready for LCM. 16. Discard all used staining and dehydration solutions according to standard procedures.

3.1.4

Slide Scraping and Total RNA Isolation (RNeasy Purification Kit)

Total RNA is extracted using published methodology with modifications (9, 10, 12). Wear double disposable clean gloves throughout the RNA isolation. 1. Using a RNase-free scalpel blade, scrape a stained and dehydrated 7-µm kidney section (as above) from one slide, into 350 µL of RLT buffer (Qiagen) with added β-mercaptoethanol (β-ME) (10 µL β-ME per 1 mL Buffer RLT) (RNeasy™ purification kit; Qiagen). 2. Vortex for 1–2 min. 3. Add 1 volume (usually 350 µL or 600 µL) of 70% ethanol to the cleared lysate, and mix immediately by pipetting. Do not centrifuge.

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4. Apply up to 700 µL of the sample, including any precipitate that may have formed, to an RNeasy™ (Qiagen) mini column placed in a 2-mL collection tube (supplied). Close the tube gently, and centrifuge for 15 s at ≥ 8,000 × g. Discard the flow-through.

3.1.5

On-Column DNase Digestion with the RNase-Free DNase Set

1. Pipette 350 µL Buffer RW1 into the RNeasy™ mini column, and centrifuge for 15 s at ≥ 8,000 × g to wash. Discard the flow-through. Reuse the collection tube in step 4. 2. Add 10 µL DNase I stock solution (see above) to 70 µL Buffer RDD. Mix by gently inverting the tube. Buffer RDD is supplied with the RNase-Free DNase Set (see Note 1). 3. Pipette the DNase I incubation mix (80 µL) directly onto the RNeasy™ silicagel membrane, and place on the bench top (20–30°C) for 15 min (see Note 2). 4. Pipette 350 µL Buffer RW1 into the RNeasy™ mini column, and centrifuge for 15 s at ≥ 8,000 × g. Discard the flow-through. Continue with the first Buffer RPE wash step in the relevant protocol. 5. Transfer the RNeasy™ column into a new 2-mL collection tube. Pipette 500 µL Buffer RPE onto the RNeasy™ column. Close the tube gently, and centrifuge for 15 s at ≥ 8,000 × g to wash the column. Discard the flow-through. Reuse the collection tube in step 8 (see Note 3). 6. Add another 500 µL Buffer RPE to the RNeasy™ column. Close the tube gently, and centrifuge for 2 min at ≥ 8,000 × g to dry the RNeasy™ silica-gel membrane. It is important to dry the RNeasy™ silica-gel membrane since residual ethanol may interfere with downstream reactions. This centrifugation ensures that no ethanol is carried over during elution (see Note 4). 7. Place the RNeasy™ column in a new 2-mL collection tube (not supplied), and discard the old collection tube with the flow-through. Centrifuge in a microcentrifuge at full speed for 1 min. 8. To elute, transfer the RNeasy™ column to a new 1.5-mL collection tube (supplied). Pipette 30 µL RNase-free water directly onto the RNeasy™ silica-gel membrane. Close the tube gently, and centrifuge for 1 min at ≥ 8,000 × g to elute.

3.2

Laser Capture Microdissection

Arrange a microscope time slot in advance. Check the microscope head, computer, and monitor connections before slide staining and dehydration. Have 50 µL of RNA Extraction Buffer (XB) in 0.5-mL tube ready for the CapSure™ Macro LCM Cap (Arcturus Engineering). 1. Place the sample slide on the microscope. 2. Locate the target area.

6 Laser Capture Microdissection of Kidney Tissue

3. 4. 5. 6. 7. 8.

9. 10. 11.

12.

79

Place the LCM Cap (CapSure™; Arcturus Engineering). Pulse the laser. Lift the cap. Pipette 50 µL Extraction Buffer (XB) into a 0.5-mL microcentrifuge tube. Insert the CapSure™ Macro LCM Cap onto the microcentrifuge tube using an LCM Cap Insertion Tool. Invert the CapSure Cap–microcentrifuge tube assembly. Tap the microcentrifuge tube to ensure all Extraction Buffer (XB) is covering the CapSure™ Macro LCM Cap. Incubate the assembly for 30 min at 42–50°C in an oven or sand heater. Centrifuge the assembly at 800 × g for 2 min to collect cell extract into the microcentrifuge tube. After centrifugation, the microcentrifuge tube contains the cell extract required to complete the protocol. Remove the CapSure™ Macro LCM Cap and save the microcentrifuge tube with the cell extract in it. Proceed with the RNA isolation protocol or freeze the cell extract at –80°C.

3.3

Isolation of RNA from Laser-Captured Tissue

Captured tissue RNA (e.g., glomerular RNA) may be extracted using a PicoPure™ RNA Isolation Kit (Arcturus Engineering). To avoid genomic DNA contamination, all the RNA samples should be treated with DNase (Qiagen), according to the manufacturer’s protocol. 3.3.1

RNA Isolation

1. Precondition the RNA Purification Column: (i) Pipette 250 µL Conditioning Buffer (CB) onto the purification column filter membrane. (ii) Incubate the RNA Purification Column with Conditioning Buffer for 5 min at room temperature. (iii) Centrifuge the purification column in the provided collection tube at 16,000×g for 1 min. 2. Pipette 50 µL of 70% ethanol into the captured tissue RNA. Mix well cell extract from Part 1 (RNA Extraction). Mix well by pipetting up and down. Do not centrifuge. 3. Pipette the cell extract and ethanol mixture into the preconditioned purification column. The cell extract and ethanol will have a combined volume of approximately 100 µL. 4. To bind RNA to the column, centrifuge for 2 min at 100 × g , immediately followed by a centrifugation at 16,000 × g for 30 s to remove flow-through. 5. Pipette 100 µL Wash Buffer (W1) into the purification column and centrifuge for 1 min at 8,000 × g.

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R.P. Woroniecki, E.P. Bottinger

DNase Treatment (RNase-Free DNase Set)

1. Pipette 5 µL DNase I Stock Solution to 35 µL Buffer RDD (provided with RNaseFree DNase Set). Mix by gently inverting. For isolations from larger samples such as cell pellets, pipette 10 µL DNase I stock solution to 30 µL Buffer RDD. 2. Pipette the 40 µL DNase incubation mix directly into the purification column membrane. Incubate at room temperature for 15 min. 3. Pipette 40 µL PicoPure™ RNA Kit Wash Buffer 1 (W1) into the purification column membrane. Centrifuge at 8,000 × g for 15 s. 4. Pipette 100 µL Wash Buffer 2 (W2) into the purification column and centrifuge for 1 min at 8,000 × g. 5. Pipette another 100 µL Wash Buffer 2 (W2) into the purification column and centrifuge for 2 min at 16,000 × g. Check the purification column for any residual wash buffer. If wash buffer remains, centrifuge again at 16,000 × g for 1 min. 6. Transfer the purification column to a new 0.5-mL microcentrifuge tube. 7. Pipette 11 µL Elution Buffer (EB) directly onto the membrane of the purification column. Gently touch the tip of the pipette to the surface of the membrane while dispensing the EB to ensure maximum absorption of EB into the membrane. 8. Incubate the purification column for 1 min at room temperature. 9. Centrifuge the column for 1 min at 1,000 × g to distribute EB in the column, then for 1 min at 16,000 × g to elute RNA. The isolated RNA is now ready for use in downstream applications such as reverse transcription. The entire sample may be used immediately or stored at –80°C until use.

3.4

RNA Quantity and Quality

We routinely use the Agilent 2100 Bioanalyzer and lab-on-chip technology (RNA 6000 Pico LabChip kit; Agilent) to verify the quantity and quality of RNA. The Agilent bioanalyzer is a microfluidics-based platform that is ideally suited to checking the quality and quantity of RNA in LCM as it uses less material and is more rapid than comparable gel electrophoresis techniques. A PicoRNA™ chip should be used for captured (glomerular) and total RNA. Use 1 µL of the above sample to examine for integrity of 18S and 28S eukaryotic ribosomal RNA, and to determine RNA concentration in each sample. Complete analysis in accordance with the manufacturer’s directions. Subsequently, RNA is stored at –80°C for further analysis. The storage period ideally should not exceed 2–3 months (13, 14).

3.5

Downstream Analysis

The material prepared above is available for subsequent analysis using reverse transcription (RT)-PCR. Comparisons between RNA expression in whole kidney sections and captured material provide a quantitative measure of expression in renal

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compartments relative to the whole kidney (12). We routinely perform quantitative RT-PCR using SYBR-Green PCR and the ABI PRISM™ 7900HT Sequence Detection System (Applied Biosystems, Foster City, CA, USA), in accordance with published protocols.

4

Notes

1. DNase I is especially sensitive to physical denaturation. Mixing should only be carried out by gently inverting the tube. Do not vortex. 2. Make sure to pipet the DNase I incubation mix directly onto the RNeasy ™ silica-gel membrane. DNase digestion will be incomplete if part of the mix sticks to the walls or the Oring of the RNeasy™ column. 3. Buffer RPE is supplied as a concentrate. Ensure that ethanol is added to Buffer RPE before use. 4. Following the centrifugation, remove the RNeasy™ mini column from the collection tube carefully so the column does not contact the flow-through, as this will result in carryover of ethanol.

References 1. Henger A, Kretzler M, Doran P, et al. Gene expression fingerprints in human tubulointerstitial inflammation and fibrosis as prognostic markers of disease progression. Kidney Int. 65:904– 917, 2004. 2. Higgins JP, Wang L, Kambham N, Montgomery K, Mason V, Vogelmann SU, Lemley KV, Brown PO, Brooks JD, van de Rijn M. Gene expression in the normal adult human kidney assessed by complementary DNA microarray. Mol Biol Cell. 15(2):649–56, 2004. 3. Kretzler M, Cohen CD, Doran P, et al. Repuncturing the renal biopsy: strategies for molecular diagnosis in nephrology. J Am Soc Nephrol. 13:1961–1972, 2002. 4. Henger A, Schmid H, Kretzler M. Gene expression analysis of human renal biopsies: recent developments towards molecular diagnosis of kidney disease. Curr Opin Nephrol Hypertens. 13(3):313–318, 2004. 5. Schwab K, Witte DP, Aronow BJ, Devarajan P, Potter SS, Patterson LT. Microarray analysis of focal segmental glomerulosclerosis. Am J Nephrol. 24(4):438–447, 2004. 6. Devarajan P, Mishra J, Supavekin S, Patterson LT, Potter SS. Gene expression in early ischemic renal injury: clues towards pathogenesis, biomarker discovery, and novel therapeutics. Mol Genet Metab. 80(4):365–376, 2003. 7. Patterson LT, Potter SS. Profiling gene expression in kidney development. Nephron Exp Nephrol. 98(4):e109–113, 2004. 8. Peterson KS, Huang JF, Zhu J, D’Agati V, Liu X, Miller N, Erlander MG, Jackson MR, Winchester RJ. Characterization of heterogeneity in the molecular pathogenesis of lupus nephritis from transcriptional profiles of laser-captured glomeruli. J Clin Invest. 113(12):1722– 1733, 2004. 9. Bonner RF, Emmert-Buck M, Cole K, Pohida T, Chuaqui R, Goldstein S, Liotta LA. Laser capture microdissection: Molecular analysis of tissue. Science. 278:1481–1483, 1997. 10. Fend F, Emmert-Buck MR, Chuaqui R, Cole K, Lee J, Liotta LA, Raffeld M. Immuno LCM: Laser capture microdissection of immunostained frozen sections for mRNA analysis. Am J Pathol. 154:61–66, 1999. 11. Kohda Y, Murakami H, Moe OW, Star RA. Analysis of segmental renal gene expression by laser capture microdissection. Kidney Int. 57:321–331, 2000.

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12. Woroniecki RP, Schiffer M, Shaw AS, Kaskel FJ, Bottinger EP. Glomerular expression of transforming growth factor-beta (TGF-beta) isoforms in mice lacking CD2-associated protein. Pediatr Nephrol. 21(3):333–338, 2006. 13. Kalluri R, Neilson EG. Epithelial-mesenchymal transition and its implications for fibrosis. J Clin Invest. 112(12):1776–1784, 2003. 14. Tanji N, Ross MD, Cara A, Markowitz GS, Klotman PE, D’Agati VD. Effect of tissue processing on the ability to recover nucleic acid from specific renal tissue compartments by laser capture microdissection. Exp Nephrol. 9(3):229–234, 2001.

Chapter 7

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

Abstract Quantitative gene expression analysis is fundamental to many experimental protocols and hypothesis testing in scientific research. The most popular currently used method to measure the expression level of specific genes in biological samples is real-time quantitative polymerase chain reaction (PCR). The method itself has become routine in many laboratories however stringent protocols and careful planning are required to for the generation of meaningful data. Many variations to these protocols are described in the literature. We describe here the methods used in our laboratory that have been compiled following many hours of troubleshooting and in our view they are robust protocols, providing solid data. The protocols are applicable to tissue culture cells where acute changes in gene expression are routinely observed following exposure to chemical or environmental stimuli, as well as tissue samples where gene expression is altered as a result of disease processes and interventions. Keywords Quantitative real-time PCR, Polymerase chain reaction, RNA extraction, cDNA synthesis, Cell culture, Normalisation, Primer design, Genomic DNA, Gene expression,

1

Introduction

A number of methods have been used to measure messenger RNA (mRNA) levels of specific genes. The earliest relied on Northern analysis where RNA was extracted from samples, fractionated by agarose gel electrophoresis, electrophoretically transferred to nitrocellulose membrane, and finally detected by a radioactively labelled complementary probe specific for the gene of interest. The resulting signal was captured on x-ray film. Several variations of this theme have been used including the use of non-radioactive detection methods, but all suffered from problems related to RNA quality, poor sensitivity and the inability to be used where a limited amount of RNA is available. Real-time quantitative PCR has overcome many of these issues and is amenable to high throughput, accurate and specific From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_7, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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quantitation of mRNA, large dynamic range, and sensitivity. The technique has even been used successfully with RNA isolated with single cells and with laser capture technology, and for the quantitation of rare mRNAs such as low abundance transcription and growth factors. Although real-time PCR has many benefits over traditional methods of measuring RNA levels of specific genes, mastering the technique and underlying concepts can be challenging. As a result a number of “quirks” in methodology and special “tricks” have made their way into protocols with the underlying scientific rationale remaining somewhat obscure. We describe here the protocols we have successfully used in our laboratory with cell lines, tissues, and even RNase-rich pancreatic tissue. A problem that still confronts many workers in this area is that of a suitable control to normalise between samples for the many steps in the process. These steps include the isolation of the biological sample and the resulting RNA integrity, isolation of RNA, removal of genomic DNA (gDNA) from RNA preparations, complementary (cDNA) synthesis, real-time quantitative PCR, and analysis. Finally, PCR is a “bulk-analysis” tool and any experiments using this technique should not be over- nor under-interpreted. We have observed on a number of occasions a modest reduction or increase in the expression of a specific gene when measured by PCR. On further analysis using other techniques such as immunofluorescence and in situ hybridisation, it was evident that that a significant change in the pattern of gene expression had occurred between the control and diseased kidney (1). This sort of information is lost with PCR methodology unless laser-capture (see Chap. 6) or tissue microdissection techniques are employed to isolate cells from different regions of the same organ. Similarly, it is not uncommon in tissue culture cells to see the expression of genes to remain largely unaltered following a treatment where dramatic changes have occurred in subcellular localisation (2).

2 2.1

Materials Tissue Storage and RNA Isolation

1. TrizolTM (Invitrogen, Carlsbad, CA, USA). 2. SolD, otherwise known as solution D, is made up according to the recipe of Chomcznski and Sacchi (3): 4 M guanidinium thiocyanate (hazardous), 25 mM sodium citrate (pH 7.0), and 0.5% (v/v) Sarkosyl. Add 0.36 mL mercaptoethanol per 50 mL solution D before use. Store up to 3 months at room temperature (RT) or longer at 4°C. 3. RNAlaterTM (Invitrogen). 4. Polytron PT2100 Homogeniser system and probes (Kinematica, Lucerne, Switzerland).

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5. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco/BRL, Bethesda, MD, USA) supplemented with 10% fetal bovine serum. 6. RNase cleaner, e.g. RNaseZapTM (Ambion) or RNaseOutTM (GenoTech, St. Louis, MO, USA). 7. Sterile H2O. 8. 70% ethanol. 9. 50-mL conical centrifuge tubes (Falcon; BD Biosciences, San Jose, CA, USA). 10. Chloroform. 11. Isoamyl alcohol. 12. 1.5-mL microfuge tubes (Eppendorf, Westbury, NY, USA). 13. Formamide.

2.2

Additional Materials for Cell Culture Experiments

1. 3-mL syringes with 21-gauge needles. 2. Isopropanol.

2.3

Quantitation

Quartz cuvettes and spectrophotometer (Nanodrop Technologies Inc, Wilmington, DE, USA).

2.4

DNase Treatment

1. RNase-free DNase-I (Epicentre Biotechnologies, Madison, WI, USA) is aliquoted and stored at –20°C, to avoid contamination. 2. DNA-free Kit (Ambion, Austin, TX, USA).

2.5

cDNA Synthesis

1. Μ-MLV reverse transcriptase (Invitrogen). 2. Random hexamers, 50 ng/µL, aliquoted and frozen at –20°C (Takara, Tokyo, Japan). 3. dNTPs, 10 mM each, aliquoted and frozen at –20°C, (Takara). 4. RNase inhibitor: Superase-InTM (Ambion, Austin, TX, USA).

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2.6

Quantitative Real-Time PCR

1. PCR machine: ABI 7500 Fast PCR (ABI; Applied Biosystems, Foster City, CA, USA). 2. Taqman Fast Universal master mix, SYBR Green PCR Master Mix, PCR plastic ware, eukaryotic 18s rRNA MGB endogenous control probe, and all other realtime MGB probes (ABI). 3. Oligonucleotides were purchase from (GeneWorks, Adelaide, Australia), standard synthesis and purity. 4. Software to design probes and primers (Primer Express; ABI). 5. Chromatin Immunoprecipitation kit and anti acetyl histone 3 antibody were purchased from Upstate (now part of Millipore, Charlottesville, VA, USA). 6. Data are analysed by ANOVA using Statview. Post-hoc comparisons are made among the various groups using Fishers least significant difference method. Data is expressed as mean ± SEM unless otherwise specified. P≤0.05 is considered to be statistically significant.

3

Methods

Many methods are employed for the isolation of RNA. Commercial products, both chemical and kit form abound and whether it is better to isolate mRNA versus total RNA is often debated. The needs of each individual laboratory may dictate a different emphasis. In our laboratory a number of criteria were used, including a standardised protocol across all sample types, yield and quality of RNA, scalability, cost, and safety, and in our case we adopted the view that total RNA is better than mRNA because ribosomal RNA and transfer RNA (tRNA) act as a buffer against RNase degradation. Where columnbased kits have been used we generally find the yield around 20–30% less compared with our in-house method. This is mainly due to the elimination of tRNA and other low molecular weight species from the RNA sample by column-based protocols. We routinely use an organic solvent-based method, which requires extra care in the laboratory, but meets our requirements on all other levels. We have adapted this method for use with tissue samples (e.g. kidney) as well as cells from tissue culture. It is scalable and with a little experience, the cost per sample is significantly cheaper than with other methods. The RNA isolated using the protocols below have been used for gene expression profiling, Northern analysis, as well as quantitative real-time PCR (QRT-PCR). Many variations to the methods reported below are found in the published literature.

3.1

Tissue Isolation and Storage

3.1.1

Harvesting Tissue

Animals are sacrificed by any one of a number of standard methods. Speed is the essence when isolating tissue for RNA, especially when dealing with tissues rich in

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endogenous RNases such as pancreatic tissue. In these cases, every attempt should be made to isolate these tissues/organs first from the animals. Where possible, the same section of the organ/tissue from each animal/specimen should be collected in order to make valid comparisons between treatment groups. For example, if the expression of genes in kidney cortex is under investigation, isolating cortical tissue from the same part of the kidney in each animal is important. Differential gene expression between the different regions (S1–S3) of the kidney is well documented. If the isolation of RNA from the tissue is to be delayed, the tissue needs to rapidly be stored such that RNA integrity is not compromised.

3.1.2

Storage of Tissue

After tissue is excised from animals, there are essentially two storage options. 1. Snap freeze in liquid nitrogen. Dissected tissue is inserted into small re-sealable, labelled plastic bags and dropped into liquid nitrogen to snap freeze the sample. When all the tissues have been processed in this way they are carefully retrieved from the liquid nitrogen and placed at -80°C for longer-term storage. Good quality RNA has been isolated from frozen tissue 4 or more years later. Ideally the RNA should be extracted as soon as possible and stored appropriately. 2. Storage in RNAlaterTM. A much safer and more reliable way to store tissue for RNA extraction is to immerse the tissue in RNAlaterTM (Invitrogen). This solution promotes the precipitation of both nucleic acid and proteins, thus protecting the nucleic acid and inactivating the enzymes that can potentially degrade it (see Note 1). The excised tissue is quickly cut so that no one dimension is more than 3- to 5-mm thick and subsequently dropped in 10 volumes RNAlaterTM in prelabelled sterile tubes. The recovered RNA is of excellent quality when tissue is stored in this way, even from RNase-rich pancreatic tissue. We have not used tissue stored in RNAlaterTM for any protein or immunohistochemistry studies, however a number of reports in the literature demonstrate that this can be done successfully (4).

3.2

Homogenisation of Tissues

1. The work environment and equipment where RNA work is to be conducted must be clean and work in the laboratory involving RNase should be confined. We use “RNase-cleaners” such as RNaseZapTM (Ambion) or RNaseOUTTM (GenoTech) to wipe down equipment and benches to remove potential RNase contamination. The following instructions assume the use of a Polytron Homogeniser with appropriately sized probes and tubes as discussed below. Samples should be kept on ice during this process. 2. The homogenisation part of the work is carried out in a fume hood to minimise exposure to phenol-containing aerosols that may be generated during the homogenisation process.

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3. Prior to and after use, the probes should be thoroughly cleaned with mild detergent and water. Once assembled on the homogeniser, the probe is washed by running the homogeniser at 15,000 rpm for 2–3 s each in the following four solutions, subsequently referred to as the “standard wash”: sterile water, 70% ethanol ×2, followed by sterile water, in 50-mL centrifuge tubes (Falcon). In between the homogenisation of samples, a squirt bottle with water is used to rinse away residual material and the probe is then used at 20,000 rpm for 5 s in a 250-mL beaker with distilled water to ensure minimal carryover between samples. The probe is then subjected to the standard wash and dabbed dry with a lint-free tissue prior to the next sample. After a number of samples are processed, usually 8–10 samples, the water in the beaker and the first of the water washes may have to be renewed. 4. Once the wash solutions and equipment is ready, tissue stored in RNAlaterTM is thawed from -80°C. The tissue is removed from RNAlaterTM to a clean tube containing 1 mL TrizolTM (see Note 2) per 100 mg tissue (see Note 3) and homogenised for 10–15 s at 25,000 rpm (see Note 4). For this step we use tubes that have a conical bottom into which the homogeniser probe fits snugly, with the tissue sample trapped below. This creates a vacuum effect when the homogeniser is switched on such that the sample is rapidly drawn into the blades of the homogeniser. Once the sample is drawn in to the probe, the tube is moved up and down carefully keeping the tip of the probe submersed in TrizolTM while mixing the contents of the tube to ensure adequate disruption of the tissue. The homogenate is placed on ice for 1 min and then inspected to determine if tissue disruption has been complete. This is determined by visual inspection of the homogenate to ensure no clumps of tissue remain. If tissue clumps are present, the process is repeated (see Note 5). 5. If the sample was snap frozen in liquid nitrogen, it is rapidly thawed, transferred to the tube containing TrizolTM (1 mL per 100 mg tissue) and straight to the homogeniser probe for disruption. It is important to disrupt the tissue rapidly in order to allow the TrizolTM to protect the RNA and inactivate RNases. The homogenisation process is the same as that described above for tissues stored in RNAlaterTM. 6. Once the sample is homogenised, the RNA is relatively safe but should be always kept on ice and the subsequent steps carried out quickly.

3.3

RNA Extraction from Tissue Homogenates

1. To isolate RNA, 0.2 volumes of chloroform/isoamyl alcohol (49:1) is added to each tube containing homogenate, the samples are vortexed for 15 s and then placed on ice for 10 min to allow for phases to begin separating (see Note 6). Samples are then centrifuged for 10 min at 4°C at >15,000×g or at maximum speed in larger tubes (around 4,000 rpm) for 30 min. The longer spinning time required for larger centrifuge tubes is to compensate for the lower speed. Following centrifugation, the upper aqueous phase (see Note 7) is carefully removed to new tubes (see Note 8).

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2. At this point, all aqueous supernatants are transferred to microfuge tubes to facilitate the pelleting of RNA precipitates at higher centrifugation speed. To each tube 0.6 volumes of isopropanol (0.6 mL per 1 mL of TrizolTM used initially) is added, tubes are briefly vortexed, and left at −20°C for 2 h, preferably overnight (see Note 9). Samples are then centrifuged at >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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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.

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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<0.01) compared with normal control kidney (1.09±0.44).

3.10.2

A Case Study with RNA from Rat Proximal Tubular Cells

NRK52E cells were treated 5 days with in high glucose (25 mM) with or without TGFβ (50 ng/mL), similar to our previously published study in this area (2). RNA was isolated and cDNA was synthesised as described in the protocols above. Multiplex reactions were carried out using 18S probe for the housekeeping gene against a number of FAM-labelled probes for makers of fibrosis and growth factors involved in epithelial to mesenchymal transition (EMT). The experimental results are summarised in the Table 7.4 and presented graphically in Fig. 7.5.

3.10.3

Case Study with gDNA from a ChIP Experiment Using SYBR Green

A chromatin immunoprecipitation (ChIPTM) experiment was carried out using the Upstate (now part of Millipore, Charlottesville, VA, USA) ChIPTM assay kit to establish whether TGF β 1 in the NRK52E cells causes a down-regulation in E-cadherin expression via a change in the acetylation status of histone 3 on the E-cadherin promoter (see Note 30). NRK52E cells cultured in DMEM medium were treated with TGFβ1 (10 ng/mL) for 3 days and chromatin was cross-linked with formaldehyde as per the manufacturer’s recommendations. Following the harvesting of cells and shearing of chromatin, immunoprecipitation was performed with anti acetyl-histone 3 antibody. The precipitated chromatin was isolated, the cross-links

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Table 7.4 Comparison of FI reveals TGFβ1 treatment causes an elevation of collagen I and IV levels (2.875±0.065; p< 0.0001 and 2.391±0.260; p<0.002, respectively) compared with control cells (1.008±0.07 and 1.008±0.071, respectively) Treatment Control

Gene Mean STDv SEM col I 1.008 0.140 0.070 col IV 1.008 0.142 0.071 CTGF 1.042 0.306 0.153 TGFβ1 1.012 0.174 0.087 TGFβ-II R 1.011 0.166 0.083 TGFβ1 col I 2.875 0.130 0.065 col IV 2.391 0.521 0.260 CTGF 3.135 0.395 0.197 TGFβ1 2.780 0.198 0.198 TGFβ-II R 0.896 0.079 0.040 Interestingly, connective tissue growth factor (CTGF) was also elevated (3.135 ± 0.197; p <0.0001) compared with control as was TGFβ1 (2.780 ± 0.198; p<0.0002), but not the TGFβ-II receptor (0.896 ± 0.04; p=0.259). The data is graphed in Fig. 7.4

control TGFβ1 (50ng/ml)

mRNA Fold Induction

3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0

collagen I

collagen IV

CTGF

TGFβ1

TGFβ-IIR

Fig. 7.5 Fold Induction of a number of genes in NRK52E cells following 5-day exposure to TGF (50 ng/mL) compared with control. The experiment shows triplicate samples for each treatment. The expression of all genes was increased significantly compared with control (p< 0.002), except for TGFβ-IIR, which was not significant

reversed and the gDNA fraction purified. This gDNA fraction was used to measure the level of E-cadherin promoter in control versus TGFβ1-treated cells. Given that in a typical EMT experiment, TGFβ1 results in decreased E-cadherin expression (2), TGFβ1 should also result in decreased acetylation of histone 3 on the E-cadherin promoter, which would result in decreased E-cadherin transcription in these cells compared with control. Therefore, one would predict less E-cadherin promoter gDNA to be immunoprecipitated from TGFβ1-treated cells than control cells.

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To quantify the amount of E-cadherin promoter in gDNA fractions, quantitative PCR is carried out using the SYBR green dye rather than a FAM-labelled MGB probe. In this reaction, specific primers are designed as described earlier, but the amplicons are detected by the SYBR green dye. This dye is added to the PCR reaction and it fluoresces once it is bound to double-stranded DNA. As more PCR products are generated in each PCR cycle, the level of fluorescence increases (see Note 31). 1. The following represents the ingredients in a standard PCR reaction for each sample to be analysed. Primer concentrations need to be optimised as discussed earlier with the recommended final concentration in the range of 50–300 nM: 2 × SYBR Green PCR Master Mix Forward primer (5 µM stock) Reverse primer(5 µM stock) Template Final volume with water

10 µL 0.6 µL 0.6 µL 0.5–4 µL 20 µL

2. A master mix of all the above reagents, excluding the template, is prepared and added into the wells of a PCR plate taking into account the volume of template to be used. We would typically use 1 µL of template, in which case the volume of master mix added to each well is 19 µL. At least 2 NTCs should be included in each run. 3. Templates are added to wells containing the master mix. 4. Given that there is no multiplexing with a housekeeping gene, the same volume of template should be added to each well, 0.5–4 µL, and each sample is analysed in triplicate. The volume of water in the master mix in step 1 can be adjusted to accommodate different sample volumes. In the system we use (ABI), the passive ROX dye will correct for differences in total volume. 5. The plate is centrifuged, holding at 1,000×g for 5 s before the amplification reaction on the real-time PCR machine. 6. Following the run, a single Ct is taken for each well. An average Ct is then taken of the three replicates for each sample and this is converted to a linear scale with the formula below. This value gives an approximation of the target gene’s abundance, which is compared with the value of the control group: 1 2C t . In this case study, Fig. 7.6 depicts the bound acetylated histone 3 on the E-cadherin promoter as the difference of bound to no-antibody control, as a ratio to input DNA. Less-acetylated histone 3 was bound to the E-cadherin promoter following TGFβ1 treatment compared with control cells (p = 0.0108). This is a preliminary study representing three real-time PCR analyses of a single ChIPTM assay. The present descriptions of both probe-based and SYBR real-time PCR methods are referred to as relative quantitation. Absolute quantitation is also possible, however, it is beyond the scope of this article and unnecessary for our laboratory’s purposes.

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Bound Ac-Histone 3

1.00 0.75 0.50 0.25 0.00 Control

TGFb1

Fig. 7.6 Relative abundance of bound acetylated histone 3 on the E-cadherin promoter in control and TGF 1-treated NRK52E cells. This is a preliminary study and the error bars represent three separate real-time PCR analyses of a single ChIPTM assay. In each analysis, every sample was amplified in triplicate (p = 0.0108, unpaired, two-tailed t-test)

4

Notes

1. Tissues stored in RNAlaterTM can be left at room temperature till all samples are collected, put at 4°C for weeks, or stored at -20°C or -80°C indefinitely. The only slight difficulty we have noticed using this method is that the tissue appears to get a little rubbery and some extra care needs to be taken when extracting RNA from minute tissue samples (1–2 mg). When required for RNA isolation, the cells are pelleted and treated as per the standard protocol. 2. An alternative to TrizolTM involves the procedure of Chomcznski and Sacchi (3). 3. The volume of TrizolTM used depends on the amount of tissue to be homogenised or the number of cells being used. We usually use around 1 mL TrizolTM per 100 mg of tissue, but do not go lower than 200 µL for very small pieces of tissue because of the way the homogeniser probe works. For tissue culture cells we use 0.8 mL TrizolTM for up to 5×106 cells. Larger numbers of cells can be used however the purity is subsequently affected as the gDNA makes the lysed cells viscous and is co-purified with the RNA such that it forms a substantial proportion of the nucleic acid in the RNA preparation. For the same reason, small volumes of TrizolTM for tissues should be avoided where possible. 4. The homogeniser probes use the solution they are immersed in for lubrication. For this reason they should never be used without the probe tip submersed in solution. When using the homogeniser, allow the probe to sit in TrizolTM with the sample before switching on the unit. Once switched on, wind up to the appropriate speed, being careful to always keep the tip of the probe (where the slits are) submerged. When moving the tube up and down to ensure total disruption of the tissue sample, ensure that the probe tip never exits the TrizolTM. Wind down the speed to the lowest setting before switching off the homogeniser. Wait until the probe stops before lowering the tube with the sample. 5. If repeat homogenisation is required, ensure the sample is chilled on ice between each repeat to prevent overheating of the sample. 6. During the 10-minute incubation on ice, organic and aqueous phases should begin to separate. If this does not occur, centrifugation may clarify the phases but it is also possible that the amount of cellular debris is too much load for the volume of TrizolTM and the sample may have to be diluted. This involves the further addition of 0.5 volumes of TrizolTM and 0.1 volume of chloroform/isoamyl alcohol (49:1), followed by vortexing and placing on ice a further 10 min. If the problem persists, repeat the dilution.

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7. Occasionally, the aqueous phase following the addition of chloroform and centrifugation collects below the phenol phase (yellow-coloured phenol on top). This situation occurs when the density of the aqueous phase is higher than the organic phase, for example when too much RNAlaterTM is carried over with the tissue. To avoid this problem, make sure the tissue sample is relatively “dry” of RNAlaterTM. If it does occur, add an equal volume of TrizolTM and chloroform as per the standard protocol to dilute out the higher density sample. Sometimes, depending on the density of the sample, the sample may have to be diluted twofold to fourfold. 8. Removal of the aqueous phase to fresh tubes is relatively straightforward except when the gDNA has not been sheared properly, or where too much sample has been used for the volume of TrizolTM. Care should be taken at this point to avoid the interface that contains proteins and some gDNA. On occasions the organic phase appears to be drawn up with the aqueous phase. This is due to inadequate shearing of gDNA that consists of long strands, which can disturb the interface when the aqueous phase is being removed. If this occurs, the gDNA may have to be sheared by passing the sample through a 21-gauge needle a few times before centrifuging to clarify samples. To visualise the interface more clearly after centrifugation, especially for extractions from cell lines, tubes can be placed in a cold rack sitting on ice. The cold temperature will make the interface appear whitish in colour. Chilling the sample too much will make both lower and upper phases appear cloudy. If this happens, bring the sample gently to room temperature to reverse the situation. 9. Precipitation of nucleic acids depends on a number of variables. In this work we need to consider the concentration of RNA, salt and alcohol, the temperature of incubation, and centrifugation speed and duration. We find it convenient to leave the samples at -20°C overnight and collect the RNA precipitate the following day, but 1 h at -20°C is adequate. It is important to NOT precipitate the RNA at -80°C as this extremely low temperature results in increased viscosity of the solution and in fact freezes the solution, preventing the formation of the RNA precipitate. If very low concentrations of RNA are expected (<2 µg/mL), the subsequent centrifugation step should be increased to 30 min. 10. When collecting RNA precipitate by centrifugation, it is advisable to spin the Eppendorf tubes with the hinge outermost so that the location of the RNA pellet can be easily identified. This is particularly important if low yields of RNA are anticipated. On occasions the RNA pellet may be hard to visualise while in isopropanol/ethanol. Usually when the supernatant is removed, the pellet becomes more visible. Also, if the tubes are mixed (inverted) just prior to centrifugation, the pellet often may be harder to see as it will form a speckled layer along the side of the tube and not a distinct pellet at or near the bottom. 11. When redissolving RNA it is important to ensure the RNA pellet is totally dissolved. Large pellets may appear to go into solution but closer investigation may reveal they become translucent “lenses”. In such cases some coaxing may be required with heating for 15 min at 50°C, pipetting, and mixing. If the pellets are not over dried or spun too hard in the ethanol wash, RNA will dissolve relatively easily. 12. For dissolving RNA, we use sterile, nuclease-free water, non-DEPC treated, since any residual DEPC can interfere with the reverse transcriptase enzyme in the cDNA synthesis reaction. This grade of water can be purchased from a number of suppliers and removes the potential for contamination in laboratories where much DNA is processed. 13. The Solution D method of Chomcznski and Sacchi (3) is known to many laboratories. It essentially involves the lysis of cells in guanidinium thiocyanate, followed by the addition of sodium acetate and acid phenol in separate steps, whereas the main protocol we use combines these reagents in a single step. An added advantage of the TrizolTM method is that mercaptoethanol is not required. 14. Spectrophotometric analysis of DNA is routine in all molecular laboratories. The use of the Nanodrop spectrophotometer (Nanodrop) is favoured as this instrument requires 1 µL of sample, is quick, very accurate, easy to use, and does not require sample dilution or quartz cuvettes. The Nanodrop also provides a spectral analysis of each sample, clearly displaying RNA integrity. For quantitation of RNA in other spectrophotometers, quartz cuvettes that use 80 µL volumes are best, as they require less sample per quantitation. Using this cuvette, 1 µL

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of RNA sample is added to 79 µL water (or 2–10 µL of sample is diluted in 0.5- or 1-mL cuvettes) and the absorbance at 260 and 280 nm is measured. Cuvettes should be RNase-free. This can be accomplished by washing in 0.1 M NaOH, followed by rinsing in clean water. The threshold of detection for the instrument used needs to be kept in mind for confidence in the measurements. 15. For quantitation of RNA, the absorbance at 260/280 is measured following dilution of RNA in water and water is used as the blank. The measurement of RNA purity is affected by pH and therefore the 260/280 ratio needs to be determined in TE buffer. A ratio of 1.8–2.1 indicates good purity. Since samples are often limiting, we routinely measure the sample in water and only use samples that have a 260/280 ratio >1.5 when using cuvettes, or >1.9 with the Nanodrop. 16. The concentration of RNA in water is determined as follows: an absorbance of 1 at 260 nm is equivalent to 40 µg of RNA (the Nanodrop does the calculations automatically). Thus for a sample of 10 µL of RNA with an A260 of 0.18 which has been diluted 1:80 in water, the following formula is used. Note that the calculations are only valid for samples in water: Concentration of RNA (µg/µL)

Total RNA yield

= 40 × A260 × dilution factor = 40 × 0.18 × 80 = 576 µg/mL = concentration of RNA × volume of sample (mL) = 576 µg/mL × 0.01 mL = 5.76 µg

17. RNA samples can be stored at –80°C for a number of years. Samples can also be stored at –20°C in a manual defrost freezer for up to 1 year without appreciable loss of RNA integrity. For longer-term storage, the RNA should be dissolved in formamide and stored at –80°C. For precious samples, cDNA should be synthesised and stored as well as it is more stable than RNA. 18. gDNA contamination is often visible as a high molecular weight band near the top of the gel when RNA is analysed on an agarose gel. gDNA contamination is present even if it is not visible on a gel. To minimise gDNA contamination, larger volumes of TrizolTM should be used where possible and the interface between aqueous and organic phases should be avoided during the RNA extraction procedure. To assess gDNA contamination, PCR is carried out on DNase-treated RNA with primers known to amplify from gDNA. 19. The limiting reagent in the DNA-free kit of Ambion is the RNase-free DNase. We have since found that a number of other sources of RNase-free DNase work equally well and are inactivated by the inactivation reagent. Avoid overloading the reaction with RNA, as this will limit the removal of gDNA from the sample. We usually do not use more that 6 µg of RNA per DNase digestion reaction. 20. The recommended volume of inactivation reagent suggested by Ambion is 0.1 volumes. This is 1 µL for a standard 10 µL reaction volume but we never use less than 2 µL. The reagent has to be resuspended well before pipetting because it settles out very quickly. We find it easier in practice to use 2 µL per reaction. In all cases, the inactivation reagent should be thoroughly mixed before being added to the sample. 21. The inactivation reagent should be pelleted and removed from RNA samples according the manufacturer’s recommendations. However we have encountered no problems when leaving the beads in the sample as long as the samples are centrifuged to pellet beads before removal of an RNA sample for subsequent reactions. 22. The volume of DNase-treated RNA can be varied and the water adjusted to make up the final reaction volume. We tend to aim for around 1 µg RNA per cDNA reaction for consistency across samples and within experiments. The reaction works well with less RNA but we avoid using much more than 1.5 µg as this affects the kinetics of the reaction and the representation of genes expressed at lower levels in the RNA sample. 23. The incubation of RNA at 70°C is to ensure denaturation of RNA to enable the random hexamers to anneal.

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24. Some of the reaction components are very labile (RNase inhibitor, MMLV) and should therefore only be removed from the freezer when ready for use, and should be returned to the freezer as quickly as possible. 25. To test for sequence specificity, we BLAST our amplicon sequence using the NCBI website. Since much of our work involves the use of different models, we attempt to design a single set of primers and probes that will work on human, mouse, and rat sequences where possible. Some of our work results in transfecting genes of one species into cells of another and in these experiments we attempt to design primers to distinguish between the endogenous gene and the transfected. This is not always easy or possible. 26. Much care needs to be taken with handling and storage of primers and probes. The dyes on probes are light sensitive and should therefore not be left out for extended periods of time. Aliquoting and storage minimises the risk of over exposure to light, but also reduces the potential for contaminating the entire stock or either primers or probes. Both primers and probes are fairly stable when it comes to freeze–thawing aliquots. However we aliquot in 0.5mL Eppendorf tubes, the size of the aliquot depending on the size of the average experiment in order to avoid multiple freeze–thaw cycles. 27. Explaining the steps required for setting up the PCR machine is beyond the scope of this review. The details are adequately explained in the instrument manual. We normally run the PCR reactions for 50 cycles as genes expressed at low levels often appear late. For example the AT2R usually appears at around cycle 34 and in some experimental protocols the expression is repressed such that it appears at cycle 38–39. 28. It is always advisable to include control reactions on the PCR plate to establish the background conditions. These may include a number of different controls however we generally use NTCs in the cDNA synthesis and PCR reactions to establish if any contamination is present during these steps. To minimise the possibility of contamination, plasmid and conventional PCR work should be restricted to a confined area in the laboratory. Often dedicated pipettes, filter tips, and PCR workstations are the best option in laboratories where nucleic acid work is the norm. Prevention is better than trying to establish from where contamination has come. 29. The Ct value of 18S–VIC can be used as a guide as to the quality of the cDNA. Typically, Ct values ranging between 8 and 15 indicate relatively good quality cDNA. We prefer the range 8–12 in our experiments. Furthermore, large variations in 18S–VIC Ct values within the same group of samples indicates inefficient cDNA synthesis and even poor RNA quality, both of which would dramatically affect gene expression data. 30. A discussion on epigenetics and the regulation of gene expression is beyond the scope of this review. The example used here is the way QRT-PCR can be used to measure levels of any nucleic acid, RNA (or cDNA) in the earlier case studies, and gDNA here using the SYBR green method. 31. When using SYBR green dye to quantify PCR products, it is imperative that the specificity of the amplification be assessed. This can be done in two ways. Firstly, a dissociation curve is produced of the final reaction product. This function is performed on the real-time PCR machine where a graph of change-in-fluorescence is plotted against temperature. The derivative plot will show a single large peak in the case where mainly one amplicon has been produced in the reaction. The presence of non-specific amplicons cause additional peaks to be detected. The second method of assessing specificity involves the size-separation of PCR products on an agarose gel, which are then visualised under ultraviolet light with dye that fluoresces on binding double-stranded DNA e.g. SYBR, ethidium bromide, or GelRedTM (Biotium Inc). The presence of a single band of the predicted size indicates the specific amplification of the desired product. Both of these methods will also detect the undesired production of primer dimers, which results from the partial pairing of primers and subsequent elongation by the DNA polymerase to yield small PCR products. We use the ABI Primer Express software to design primers as it reduces the chance of primer–dimer problems. We design a primer and probe set to the standard specifications as probe-based real-time PCR, but only order and use the primers. In cases where the detection by SYBR is not specific enough, one need only order the probe and switch to probe-based real-time PCR.

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References 1. Thomas Μ.C., Tikellis C., Burns W.C., Thallas V., Forbes J.Μ., Cao Z., Osicka T.Μ., Russo L. M., Jerums G., Ghabrial H., Cooper M.E., and Kantharidis P. (2003) Reduced tubular cation transport in diabetes: prevented by ACE inhibition. Kidney Int. 63, 2152–2161. 2. Burns W.C., Twigg S.M., Forbes J.M., Pete J., Tikellis C., Thallas-Bonke V., Thomas M.C., Cooper M.E., and Kantharidis P. (2006) Connective tissue growth factor plays an important role in advanced glycation end product-induced tubular epithelial-to-mesenchymal transition: implications for diabetic renal disease. J. Am. Soc. Nephrol. 17, 2484–2494. 3. Chomczynski P. and Sacchi N. (1987) Single step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162, 156–159. 4 Florell S.R., Coffin C.M., Holden J.A., Zimmermann J.W., Gerwels J.W., Summers B.K., Jones D.A., Leachman S.A. (2001) Preservation of RNA for functional genomic studies: a multidisciplinary tumor bank protocol. Mod. Pathol. 14, 116–128. 5 Livak K.J., Flood S.J., Marmaro J., Giusti W., Deetz K.(1995) Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization. PCR Methods Appl. 4,357–352. 6 de Kok J.B., Roelofs R.W., Giesendorf B.A., Pennings J.L., Waas E.T., Feuth T., Swinkels D.W., and Span P.N. (2005) Normalization of gene expression measurements in tumor tissues: comparison of 13 endogenous control genes. Lab. Invest. 85, 154–159. 7. Livak K. (1997) PRISM, ABI 7700 Sequence Detection System User Bulletin Number. 2, pp. 3–10.

Chapter 8

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

Abstract Leukocytes mediate some forms of glomerulonephritis, particularly severe proliferative and crescentic forms. The renal glomerulus is one of the few sites within the microvasculature in which leukocyte recruitment occurs in capillaries. However, due to the difficulty of directly visualising the glomerulus, the mechanisms of leukocyte recruitment to glomerular capillaries are poorly understood. To overcome this, a murine kidney can be rendered hydronephrotic, by ligating one ureter, and allowing the mouse to rest for 12 weeks. This allows the visualisation of the glomerular microvasculature during inflammatory responses. In inflammation, in this example induced by anti-glomerular basement membrane (GBM) antibody, leukocytes can be observed undergoing adhesion in glomerular capillaries using intravital microscopy. Leukocyte adhesion can be quantitated using this approach. An observation protocol involving few, limited periods of epifluorescence avoids phototoxicity-induced leukocyte recruitment. The process of hydronephrosis does not alter the ability of anti-GBM-antibody to induce a glomerular inflammatory response. This approach allows detailed investigation of the mechanisms of leukocyte recruitment within glomeruli. Keywords Glomerulus, Capillary, Leukocyte, Inflammation, Intravital microscopy

1

Introduction

Glomerulonephritis, inflammation of glomeruli, is a leading cause of end-stage renal failure worldwide. As part of this disease, leukocyte subsets, particularly neutrophils, monocytes, and T lymphocytes, are recruited to glomeruli (1). Experimental models of glomerulonephritis have demonstrated that these leukocytes play central roles in mediating tissue damage and renal dysfunction, prompting a number of studies investigating the molecular basis for this recruitment. The process of leukocyte recruitment in general requires a sequence of interactions between leukocytes in the mainstream of blood flow and endothelial cells lining the microvasculature of the inflamed tissue (2). Due to the dynamic nature From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_8, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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of these interactions, much of what we know about the process of leukocyte recruitment has been learnt from approaches that allow visualisation of leukocyte–endothelial cell interactions under conditions that maintain normal perfusion in the microvasculature of experimental animals (3). This technique, known as intravital microscopy, was originally used in analysis of vessels in transparent tissues, such as the mesentery and cremaster muscle. However, application of this approach to the glomerulus has been limited due to the kidney’s opaque structure. There is a growing body of evidence that the mechanisms of leukocyte recruitment in the glomerulus do not follow the conventional paradigm. In most tissues, leukocyte–endothelial cell interactions are restricted to the postcapillary venules. In contrast, the glomerular capillaries readily support leukocyte recruitment during inflammatory responses. In addition, leukocytes generally have to undergo deformation to pass through these narrow vessels, bringing into question the necessity for the rolling step observed in larger post-capillary venules (4). Furthermore, stimuli that induce expression of the specific adhesion molecules that mediate leukocyte rolling in venules fail to induce their expression in capillary endothelial cells (5, 6). Together, these observations imply that the process whereby leukocytes undergo recruitment in glomeruli has several unique characteristics. To fully understand the process of glomerular leukocyte recruitment, it was necessary to develop a technique to allow visualisation of this process in intact glomeruli in vivo. This was achieved using the process of experimental hydronephrosis (7, 8). Obstruction of urine outflow via ureteric ligation leads to a progressive atrophy of the renal tubulointerstitium. In mice, 12 weeks after ureteric ligation, the kidney is reduced to a thin rim of cortex capable of being transilluminated for visualisation. This tissue retains perfused glomeruli that are amenable to intravital microscopy for the analysis of leukocyte recruitment. When combined with experimental models of glomerular inflammation, this approach allows analysis of the dynamic and transient interactions of leukocytes with the glomerular endothelial surface. This article provides detail on the techniques for performing ureteric ligation and intravital microscopy of the hydronephrotic kidney. This technique was recently used to demonstrate the unique nature of the process of leukocyte recruitment to glomeruli (9).

2 2.1

Materials Animals

Mice ideally should undergo ureteric ligation between 4 and 6 weeks of age. In general, gene-deficient or transgenic mice can be used in this procedure, according to the aims of each experiment.

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1. Inhalation anesthetic such as Ethrane (Abbott Australasia, Kurnell, NSW, Australia). 2. Small chamber for the induction of anesthesia—preferably clear. 3. Nose-cone for administration of inhalation anesthetic during surgical procedure. 4. Perspex board for surgical procedure. 5. Electric clippers to shave abdomen. 6. Surgical tape. 7. 10/0 non-absorbable silk (Dynek Sutures; Dynek Pty Ltd, Hendon, Australia). 8. 4/0 non-absorbable silk with attached needle (Dynek Sutures; Dynek Pty Ltd). 9. Post-operative analgesic, e.g. buprenorphine (Subutex; Reckitt Benckiser, West Ryde, Australia). 10. Surgical instruments including #5 jewellers’ forceps (for handling ureter), retractors (× 2), scissors. 11. Chlorhexidine or other topical antiseptic agent. 12. Dissecting microscope.

2.3

Model of Glomerular Inflammation—Anti-Glomerular Basement Membrane Antibody (See Note 1)

1. Anti-glomerular basement membrane (GBM) antibody, raised against a murine renal or glomerular membrane fraction in a sheep or a rabbit, using sera that has been decomplemented, then purified by saturated ammonium sulfate precipitation and dissolved in phosphate-buffered saline (PBS) (10, 11). 2. Control non-immune antibodies from the same species, purified in an identical manner.

2.4

Instruments and Equipment for Intravital Microscopy

1. Anesthetic: 150 mg/kg ketamine hydrochloride (Pfizer, West Ryde, Australia) and 10 mg/kg xylazine (Troy Laboratories, Smithfield, Australia). 2. Instruments: Graefe forceps, self-closing forceps (Dumont N5), micro fine-tip ophthalmic cautery unit. 3. Catheter for i.v administration of additional anesthetic, antibodies, etc. (PE10 tubing or equivalent). 4. Optically clear viewing platform and thermo-controlled heat pad.

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5. Upright intravital microscope (e.g. Axioplan 2 Imaging; Carl Zeiss, Australia), equipped with a water immersion × 20 objective (× 20/0.50 NA) and light source for fluorescence imaging. 6. High-sensitivity CCD video camera suitable for imaging fluorescence (e.g. Dage-MTI IR-1000). 7. Videocassette recorder or digital recording device. 8. Bicarbonate-buffered saline (pH 7.4; at 37°C) for superfusion of exteriorised kidney. 9. Rhodamine 6G (0.05% in saline).

3 3.1

Methods Unilateral Ureteric Ligation

1. Mice (4–6 weeks old) are anesthetised via inhalation of Ethrane. During the procedure, anesthesia is maintained by Ethrane delivered via a nose-cone. The level of anesthesia must be continually and vigilantly assessed by monitoring the respiration rate and responsiveness to tail pinch. The degree of anesthesia is adjusted by altering the position of the nose cone closer (if mouse starts to become responsive) or further away (if respiration ceases or is greatly reduced) from the mouse, as required. 2. The mouse is prepared for surgery by securing the mouse to a board in a supine position using surgical tape on all four limbs, shaving the ventral abdomen, and swabbing the area with chlorhexidine. 3. Using scissors, a 1.5-cm midline incision is made in the skin of the lower abdomen, followed by a similar incision in the abdominal wall. The abdominal incision is held open using retractors. The abdominal organs are gently moved to the right of the abdominal cavity using Q-tips. 4. Using a dissecting microscope, the left ureter is identified. It is located at the rear of the peritoneal cavity, and identified as a thin, pale tubular structure adjacent to the peritoneal wall. Care should be taken to avoid traction on the ureter as this can induce renal subcapsular hemorrhage or hemorrhage into the renal pelvis, rendering the kidney unsuitable for intravital microscopy. The ureter is then ligated twice with 10/0 non-absorbable silk sutures and divided. 5. The organs are returned to their original positions and buprenorphine (10 µg) administered into the abdomen to provide post-operative analgesia. The incisions in the abdominal wall and skin are closed in two layers with 4/0 non-absorbable silk sutures. Anesthesia is removed and the mouse placed in a warm environment on its own to recover. After mice are fully awake, groups of mice that have undergone the same procedure can be housed together. 6. Post-operative care: mice should be checked a few hours after surgery and the following day to ensure sutures remain in place. Generally, this procedure is well tolerated by the mice and they survive the 12 weeks required for complete hydronephrosis without incident (see Note 2).

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3.2 Preparation of Mice for Intravital Microscopy 1. 12 weeks after ureteric ligation, the mouse is anesthetised via intraperitoneal injection of 150 mg/kg ketamine hydrochloride and 10 mg/kg xylazine. A jugular vein is cannulated with the PE10 catheter to allow administration of further anesthetic and other reagents, including fluorescent dyes (see Notes 2 and 3). 2. Place the mouse on a transparent viewing platform equipped with a thermocontrolled heat pad to maintain the core temperature at 37°C. 3. Exteriorise the hydronephrotic kidney through a lateral abdominal incision through the skin and abdominal wall, such that the kidney lies over the coverslip of the viewing platform. Ideally, the incision in the abdominal wall is made using a micro-cautery unit to avoid bleeding. Application of gentle pressure on the opposite side of the abdomen will result in the kidney being exteriorised through the incision. 4. Drain the hydronephrotic kidney of urine using a syringe with 30-gauge needle inserted into the side of the kidney, entering the kidney through a relatively avascular region. The volume of urine removed from the kidney should be in the range 0.3–1.0 mL. To achieve the optimal visualisation of the renal microvasculature, it is advisable to leave a small amount of urine in the renal pelvis to prevent the kidney from collapsing. 5. Using self-closing forceps to grasp the periphery of the kidney, two to three lengths of 4/0 silk are tied to the kidney. These are used to gently extend the kidney over the clear viewing platform (Fig. 8.1). Care should be taken not to stretch the kidney, as this can disrupt blood flow. 6. Superfuse the kidney with a bicarbonate-buffered saline solution (pH 7.4, 37°C), then cover the preparation with a large coverslip held in place with vacuum grease (Fig. 8.1). If using an immersion objective lens, it is advisable to include a barrier on the coverslip to prevent the immersion fluid (saline) from running off the coverslip. This can be achieved using a line of vacuum grease on the edge of the coverslip adjacent to the mouse. 7. The renal microvasculature can then be observed with an intravital microscope. Both wide-field transillumination and epifluorescence microscopy can be used.

3.3

Imaging the Hydronephrotic Kidney

1. Upon initial examination of the kidney, scan the tissue using transillumination microscopy. We have found that transillumination imaging of the hydronephrotic kidney is aided by use of a green filter in the beam path. Find a clear area of tissue populated with several well-perfused glomeruli. Typically, three glomeruli can be imaged simultaneously in one field (Fig. 8.2). Once a suitable area is found, imaging is switched to epifluorescence. 2. In order to visualise recruitment of endogenous leukocytes, rhodamine 6G is used. Mice receive 50 µL of rhodamine 6G (0.05% in saline) administered i.v.

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Fig. 8.1 An anesthetised mouse displaying a hydronephrotic kidney prepared for intravital microscopy. The kidney has been exteriorised, drained of urine, extended over the viewing platform by attaching sutures to the periphery, and covered by a coverslip held in place with vacuum grease

a minute or two prior to imaging. This labels all circulating leukocytes and platelets, and will also label parenchymal cells over time (Fig. 8.2). Rhodamine 6G-associated fluorescence is visualised by epi-illumination at 520–560 nm, using a 590 nm emission filter. (NB: Care should be taken with rhodamine 6G due to its potential toxicity). Microscopic images are projected onto a monitor (Sony PVM-20N5E) using a low-light video camera (DageMTI IR-1000) and recorded for playback analysis using a videocassette recorder (Panasonic NV-HS950) (see Note 4). Recordings of 30-s duration are made of the chosen field (~three glomeruli) at intervals throughout the experiment and subsequently analysed for numbers of leukocytes passing through each glomerulus (cells per minute) and leukocyte adhesion (cells per glomerulus). A leukocyte is defined as adherent if it remains stationary within the glomerulus for at least 30 s. If the imaging protocol extends over an hour, administer an additional bolus of rhodamine 1–2 min prior to each subsequent visualisation period. Example images are shown in Fig. 8.2. Restriction of fluorescence illumination is of key importance in glomerular intravital microscopy (see Note 5). 3. Glomerular inflammation is then induced according to the model under examination (see Note 1), and recordings of glomerular leukocyte recruitment made at specified subsequent time points. In the case of the anti-GBM antibody model, the kidney is imaged 1–2 hours after antibody administration, as histological analyses have shown that leukocyte recruitment in this model is maximal at this time (12) (see Note 6).

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Fig. 8.2 Representative captured images from intravital microscopy of the hydronephrotic kidney. a Image of the transilluminated hydronephrotic kidney, demonstrating the glomeruli and associated microvasculature. b and c Images of glomeruli 2 hours after sheep globulin (b) or antiGBM antibody (c) administration, showing recruited leukocytes (arrows) within the glomerular vasculature following anti-GBM antibody administration

4

Notes

1. Inducing an inflammatory response in the glomerulus requires specific stimuli. We have found that non-specific stimuli that induce inflammation in most vascular beds, e.g. lipopolysaccharide (LPS), do not induce leukocyte recruitment to glomeruli (13). However, alternative models of glomerular inflammation include antibodies against murine myeloperoxidase (13), circulating immune complexes (14), and systemic autoimmune disease (15). The imaging protocol should be adapted to suit the model under examination. 2. The mice should be imaged 11–13 weeks after ureteric ligation (12 weeks is optimal). Prior to 11 weeks, it is highly likely that the kidney will be incompletely hydronephrotic and unsuitable for glomerular imaging. 3. Effects of hydronephrosis: a consideration in the use of this approach is that glomeruli in hydronephrotic kidneys may be altered from their original state. We assessed this possibility in our recent study by comparing parameters such as degree of sheep globulin deposition, leukocyte recruitment, P-selectin expression, and complement deposition between glomeruli from the contralateral (”healthy”) and hydronephrotic kidneys (9). Using image

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analysis of immunohistochemically stained sections, these parameters did not differ significantly in the two types of kidneys. These findings suggest that aspects of the glomerular inflammatory response are unaltered by the process of hydronephrosis. However, care should be taken to analyse parameters of relevance to each experimental design to exclude this possibility. 4. This chapter does not deal with new imaging modalities and digital image recording processes that have become readily available over recent years. For conventional microscopy, analogue video output can be recorded digitally using a DVD recorder or similar device. Similarly, newgeneration high-sensitivity digital video cameras are available. These cameras generate images in TIF sequence or AVI style formats with, in general, better resolution than analogue VHS video cameras. Finally, single- and multi-photon confocal microscopy is also being applied in intravital imaging of the kidney (16, 17). Application of these novel technologies should allow new questions regarding the mechanisms of glomerular inflammation to be addressed. 5. Epifluorescence intravital microscopy and the associated use of fluorochromes for cell labelling are recognised as having the potential to induce phototoxicity, i.e. cellular injury due to the energy associated with fluorescence (18, 19). We have found that in the hydronephrotic kidney preparation, lack of care with fluorescence illumination can induce artefactual leukocyte adhesion. We determined this by comparing leukocyte adhesion induced by anti-GBM antibody in mice in which the kidney had been exteriorised prior to administration of antibody and undergone several periods of fluorescence illumination, with that in which the kidney was exteriorised and imaged 1 hour after antibody administration. In light of this, our standard approach is to ensure that the exposure of the tissue to fluorescence is restricted, by reducing the length and number of individual recordings, and attenuating the intensity of the fluorescent beam via neutral density filters. To compensate for a reduction in fluorescence, the gain on the CCD camera can be increased. Using these steps, we were able to ensure that the increased adhesion observed was due solely to the anti-GBM antibody. In our hands, this problem seems to occur in rhodamine 6G-treated mice, but much less so in experiments using fluorochromelabelled isolated cell populations. However it is advisable to routinely design experiments to restrict the number of fluorescent observations. 6. This technique is also amenable for alternative approaches for visualising specific cell sub-populations. For example, to investigate the occurrence of rolling during glomerular inflammation, we used bone marrow-derived neutrophils labelled with carboxyfluorescein succinimidyl ester (CFSE) (20). In these experiments, avoiding the use of systemic rhodamine 6G and restriction of fluorescence to the neutrophil population resulted in the absence of fluorescent staining in the glomerulus and allowed for improved clarity of leukocyte imaging (9). However, in order to achieve sufficiently high numbers of cells delivered to the kidney, it is necessary to infuse them into the arterial circulation via a catheter inserted in the carotid artery in a retrograde direction. A similar approach can be used for analysis of accumulation of T lymphocytes, whereas platelet accumulation can be assessed following intravenous transfer of fluorochrome-labelled platelets. In addition, the advent of mice expressing fluorescent marker proteins under the control of lineage-specific promoters should allow analysis of recruitment of specific leukocyte subsets using this approach (21).

References 1. Holdsworth, S. R., Kitching, A. R., and Tipping, P. G. 1999. Th1 and Th2 T helper cell subsets affect patterns of injury and outcomes in glomerulonephritis. Kidney Int 55: 1198–1216. 2. Springer, T. A. 1994. Traffic signals of lymphocyte recirculation and leukocyte emigration: The multistep paradigm. Cell 76: 301–314. 3. Mempel, T. R., Scimone, M. L., Mora, J. R. and von Andrian, U. H. von 2004. In vivo imaging of leukocyte trafficking in blood vessels and tissues. Curr Opin Immunol 16:406–417.

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4. De Vriese, A. S., Endlich, K., Elger, M., Lameire, N. H., Atkins, R. C., Lan, H. Y., Rupin, A., Kriz, W., and Steinhausen, M. W. 1999. The role of selectins in glomerular leukocyte recruitment in rat anti-glomerular basement membrane glomerulonephritis. J Am Soc Nephrol 10: 2510–2517. 5. Jung, U., and Ley, K. 1997. Regulation of E-selectin, P-selectin and intercellular adhesion molecule-1 expression in mouse cremaster muscle vasculature. Microcirculation 4: 311–319. 6. McEver, R. P., Beckstead, J. H., Moore, K. L., Marshall-Carlson, L., and Bainton, D. F. 1989. GMP-140, a platelet a-granule membrane protein, is also synthesized by vascular endothelial cells and is localized in Weibel-Palade bodies. J Clin Invest 84: 92–99. 7. Steinhausen, M., Snoei, H., Parekh, N., Baker, R., and Johnson, P. 1983. Hydronephrosis: a new method to visualise vas afferens, efferens, and glomerular network. Kidney Int 23: 794–806. 8. Buhrle, C. P., Hackenthal, E., Helmchen, U., Lackner, K., Nobiling, R., Steinhausen, M., and Taugner R. 1986. The hydronephrotic kidney of the mouse as a tool for intravital microscopy and in vitro electrophysiological studies of renin-containing cells. Lab Invest 54: 462–472. 9. Kuligowski, M. P., Kitching, A. R., and Hickey, M. J. 2006. Leukocyte recruitment to the inflamed glomerulus: a critical role for platelet-derived P-selectin in the absence of rolling. J Immunol 176: 6991–6999. 10. Salant, D. J. and Cybulsky, A. V. 1988. Experimental glomerulonephritis. Methods Enzymol 162: 421–461. 11. Kitching, A. R., Holdsworth, S. R., and Tipping, P. G. 1999. IFN-gamma mediates crescent formation and cell-mediated immune injury in murine glomerulonephritis. J Am Soc Nephrol 10:752–759. 12. Tipping, P. G., Huang, X. R., Berndt, M. C., and Holdsworth, S. R. 1994. A role for P-selectin in complement-independent neutrophil-mediated glomerular injury. Kidney Int 46: 79–88. 13. Ruth, A. J., Kitching, A. R., Kwan, R., Odobasic, D., Ooi, J., Timoshanko, J. R., Lucis, A. J., Hickey, M. J., and Holdsworth, S. R. 2006. Anti-neutrophil cytoplasmic antibodies and effector CD4+ cells play non-redundant roles in myeloperoxidase directed crescentic. glomerulonephritis. J Am Soc Nephrol 17: 1940–1949. 14. Li, M., O’Sullivan, K. M., Jones, L. K., Semple, T., Kumanogoh, A., Kikutani, H., Holdsworth, S. R., and Kitching, A. R. 2006. CD100 enhances dendritic cell and CD4+ cell activation leading to pathogenetic humoral responses and immune complex glomerulonephritis. J Immunol 177:3406–3412. 15. Hoi, A. Y., Hickey, M. J., Hall, P., Yamana, J., O’Sullivan, K. M., Santos, L. L., James, W. G., Kitching, A. R., and Morand, E. F. 2006. Macrophage migration inhibitory factor deficiency attenuates macrophage recruitment, glomerulonephritis, and lethality in MRL/lpr mice. J Immunol 177: 5687–5696. 16. Molitoris, B. A. and Sandoval, R. M. 2005. Intravital multiphoton microscopy of dynamic renal processes. Am J Physiol Renal Physiol 288: F1084–1089. 17. Soos, T. J., Sims, T. N., Barisoni, L., Lin, K., Littman, D. R., Dustin, M. L., and Nelson, P. J. 2006. CX3CR1+ interstitial dendritic cells form a contiguous network throughout the entire kidney. Kidney Int 70: 591–596. 18. Rosenblum, W. I. 1978. Fluorescence induced in platelet aggregates as a guide to luminal contours in the presence of platelet aggregation. Microvasc Res 15: 103–106. 19. Herrmann, K. S. 1983. Platelet aggregation induced in the hamster cheek pouch by a photochemical process with excited fluorescein isothiocyanate-dextran. Microvasc Res 26: 238–249. 20. Khan, A. I., Kerfoot, S. M., Heit, B., Liu, L., Andonegui, G., Ruffell, B., Johnson, P., and Kubes P. 2004. Role of CD44 and hyaluronan in neutrophil recruitment. J Immunol 173: 7594–7601. 21. Sasmono, R. T., Oceandy, D., Pollard, J. W., Tong, W., Pavli, P., Wainwright, B. J., Ostrowski, M. C., Himes, S. R., and Hume, D. A. 2003. A macrophage colony-stimulating factor receptor-green fluorescent protein transgene is expressed throughout the mononuclear phagocyte system of the mouse. Blood 101: 1155–1163.

Chapter 9

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

Abstract The basics of in situ hybridization have been widely applied to a diverse range of situations where we need to localize the distribution of nucleic acids. Advances in other molecular techniques such as the advent of gene microarrays has not diminished the significance of in situ hybridization, but rather highlight the importance of being able to identify the topology of gene expression. In situ hybridization offers a degree of precision that is unavailable with other molecular techniques. This chapter outlines techniques used to examine the spatial distribution of gene expression in the kidney using complementary RNA (cRNA) probes with both radioactive and non-radioactive labels.

Keywords In situ hybridization, Digoxigenin, Biotin, Tyramide, Autoradiography

1

Introduction

In situ hybridization is widely applied to a diverse range of situations where we need to localize the specific distribution of genes in tissue sections. The technique has proven to be particularly useful in the study of renal pathology, where the heterogeneous nature of kidney structure has limited the usefulness of whole organ techniques. Since the first description of in situ hybridization, a variety of protocols have been employed. Treatment of tissue ranges from the use of frozen sections, which show good preservation of messenger RNA (mRNA) but poor morphology, to standard histological methods for tissue preservation, such as paraformaldehyde or formaldehyde fixation followed by paraffin embedding. This latter method may show reduced preservation of tissue mRNA levels, but gives better structural information as the tissue morphology is better preserved (see Note 1). Similarly, a variety of probe types have been used, beginning with double-stranded cDNA probes and peptide nucleic acid probes and moving on to single-stranded RNA probes (riboprobes). In addition, short single-stranded oligonucleotide probes have also been used, allowing the user to synthesize specific probes from published sequences. From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_9, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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Lastly, various methods of probe labelling and detection have been employed, including different radioactive isotopes and, more recently, non-radioactive methods such as biotin, fluorescein, and digoxigenin (DIG). For examples of the use of these methods in various tissues see refs. (1–11). In this chapter we describe the standard methods used in our laboratory for detecting target mRNA in tissue sections. In general, single-stranded RNA probes are used on formalin or paraformaldehyde-fixed paraffin or frozen sections. There are a number of advantages of RNA probes over cDNA or oligonucleotide probes that make them preferable for in situ hybridization. These include the formation of tighter (RNA: RNA) hybrids, the possibility of higher stringency post-hybridization washing (using RNase to remove unbound probe), and the lack of a competing reaction, which occurs in the case of double-stranded DNA probes, which re-anneal in addition to binding to target mRNA. The choice of label and detection system depends to some extent on the abundance of the target mRNA; in the case of low-abundance mRNA species, we have continued to use radioactively labelled probes, which give better sensitivity in our hands, with 33P now being the isotope of choice, due to its relative safety in the laboratory, and its compromise between reasonably short exposure times and good resolution. However, when mRNA species of higher abundance are being detected, probe labelling with non-radioactive methods becomes more feasible.

2 2.1

Materials Tissue Preparation and Embedding

1. Paraformaldehyde (Merck, Darmstadt, Germany). 2. Phosphate-buffered saline (PBS), pH 7.4: 0.14 M NaCl, 0.003 M KCl, 0.008 M Na2HPO4, and 0.0015 M KH2PO4. 3. Ethanol, laboratory grade. 4. Chloroform. 5. Paraffin wax e.g. Paraplast™(McCormick Scientific, St. Louis, MO, USA) with a melting point of 56°C. 6. Stainless steel embedding molds (Tissue-Tek™, Sakura Finetek, Torrance, CA, USA). 7. Glass vials for tissue processing. 8. Freeze embedding compound such as OCT™ (Tissue-Tek™, Sakura Finetek).

2.2

Slide Preparation

1. Glass microscope slides. 2. Plastic or metal slide racks. 3. Laboratory-grade detergent.

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4. 3-Aminopropyltriethoxysilane (APES; Sigma–Aldrich, St. Louis, MO, USA). 5. Acetone.

2.3

Pretreatment of Tissue Sections

All buffers are treated with 0.05% diethylpyrocarbonate (DEPC) (Sigma–Aldrich), with the exception of Tris-based buffers. 1. Histolene™ (Histolabs/Fronine, Riverstone, Australia) or xylene. 2. Ethanol. 3. Phosphate-buffered saline (PBS): 0.14 M NaCl, 0.003 M KCl, 0.008 M Na2HPO4, and 0.0015 M KH2PO4. 4. Pronase buffer (P buffer): 50 mM Tris-HCl pH 7.5, 5 mM EDTA pH 8.0. 5. Pronase E (Sigma–Aldrich). 6. Antigen retrieval solution e.g. Citra™ (BioGenex, San Ramon, CA, USA) or 0.01 M citrate buffer. 7. 0.1 M Sodium phosphate buffer, pH 7.2. 8. 4% Paraformaldehyde in PBS. 9. Double-distilled water, DEPC treated. 10. 70% Ethanol.

2.4

Labelling of the Probe

2.4.1

Isotopic Labelling

1. cDNA in the appropriate in vitro transcription vector providing polymerase sites for cRNA production (T7, T3, SP6). 2. 5× Transcription buffer: 200 mM Tris-HCl pH 7.5, 30 mM MgCl2, 10 mM spermidine (Sigma–Aldrich), and 50 mM NaCl. 3. 100 mM Dithiothreitol (DTT; Roche Diagnostics, Mannheim, Germany). 4. RNasin™, ribonuclease inhibitor (Promega Corporation, Madison, WI, USA). 5. 10 mM Adenosine 5'-triphosphate (ATP), 10 mM cytidine 5'-triphosphate (CTP), 10 mM guanosine 5'-triphosphate (GTP), 12 µM uridine 5'-triphosphate (UTP), (Promega). 6. RNA polymerases T7, T3, and SP6 (Promega). 7. Radionucleotide: 5 [α-33P]UTP. 8. DNase I (Promega). 9. Transfer RNA (tRNA) 20 mg/mL stock (Roche). 10. 7.5 M Ammonium acetate. 11. 3 M Sodium acetate, pH 5.2. 12. Ethanol.

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13. Hydrolysis buffer: 80 mM NaHCO3, 120 mM Na2CO3, and 20 mM β-mercaptoethanol. 14. Stop buffer: 200 mM sodium acetate pH 6.0, 1% glacial acetic acid, and 10 mM DTT. 15. DEPC-treated double-distilled H2O. 16. Dry heat block or waterbath accurately set at 37°C. 17. Microcentrifuge.

2.4.2

Nonradioactive Probe Labelling Using DIG

All tissue is prepared in the same manner as for radioactive probes, i.e., Sects. 2.1–2.3. 1. cDNA in appropriate in vitro transcription vector, with polymerase sites for cRNA production (T7, T3, SP6). 2. 5× Transcription buffer (see Sect. 2.4.1., item 2). 3. DIG 10× labelling mix (Roche). 4. RNasin™ (Promega). 5. RNA polymerase (T7, T3, SP6). 6. 0.2 M EDTA, pH 8.0. 7. 4 M LiCl. 8. 100% and 75% ethanol. 9. Hydrolysis buffer: 0.06 M Na2CO3, 0.04 M NaHCO3. 10. Neutralization buffer: 0.2 M NaC2H3O2, 1% acetic acid. 11. DEPC-treated double-distilled H2O.

2.4.3

Nonradioactive Probe Labelling with Biotin

Tissue and slides are prepared the same as for DIG labelling, except that biotin RNA labelling mix is substituted at Sect. 2.4.2., item 3.

2.5

Dot Blot Analysis of Non-Radioactive Probes

2.5.1

Dot Blot Analysis of DIG-Labelled Probe

To determine the concentration of the DIG-labelled riboprobe, a dot blot is necessary. 1. Nylon membrane (Amersham-GE Healthcare, Piscataway, NJ, USA). 2. DIG-labelled RNA control (Roche). 3. Buffer 1: 0.1 M (maleic acid) C4H4O4 and 0.15 M NaCl, pH 7.5.

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Buffer 2: 1% skim milk powder in Buffer 1. Anti-DIG alkaline phosphatase antibody (Roche). Buffer 3: 0.1 M Tris-HCl, 0.1 M NaCl, and 0.05 M MgCl2. Alkaline phosphatase substrate (NBT/BCIP).

2.5.2

Dot Blot Analysis of Biotin-Labelled RNA Probes

Same procedure as for DIG-labelled probes except at step 5, substitute avidin– biotin–peroxidase complex (e.g. ABC from Vectastain™kit; Vector, Burlingame, CA, USA) and at step 7, color reaction is performed with diaminobenzidine–4HCl (DAB; Sigma–Aldrich).

2.6

Hybridization

1. 10 × salt solution: 3 M NaCl, 100 mM Na2HPO4, 100 mM Tris-HCl pH 7.5, 50 mM EDTA, 0.2% bovine serum albumin, 0.2% Ficoll, and 0.2% polyvinylpyrolidone. 2. Formamide (BDH, Poole, UK). 3. Dextran sulphate (Pharmacia Biotech, Uppsala, Sweden). 4. tRNA, 20 mg/mL. 5. DEPC-treated distilled H2O.

2.7

Post-Hybridization Washes

1. 20 × Standard saline citrate (SSC) solution: 3 M NaCl and 0.3 M sodium citrate. 2. Wash buffer: 2× SSC, 50% formamide. 3. RNase A (Sigma–Aldrich). 4. RNase buffer: 10 mM Tris-HCl pH 7.5, 1 mM EDTA, and 0.5 M NaCl.

2.8

Autoradiography and Emulsion for Radioactive Riboprobes

1. X-ray film cassette. 2. Film (XAR-% Kodak, Rochester, USA; or Hyperfilm™, Amersham). 3. Liquid nuclear research emulsion (gel form) (e.g. K5 nuclear emulsion, Ilford, Cheshire, UK; or NTB-2, Kodak). 4. Developer, Phenisol™ (Ilford), diluted 1 in 4 with distilled H2O.

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Hypam™fixer (Ilford), diluted 1 in 4 with distilled H2O. Harris hematoxylin stain. Eosin stain. Scott’s tap water: 82 mM MgSO4, 42 mM NaHCO3. Mounting medium, non-aqueous.

2.9 1. 2. 3. 4. 5. 6. 7.

DIG-Labelled Probe Detection

Tris-buffered saline: 0.9% NaCl and 50 mM Tris-HCl, pH 7.5. Antibody diluent: 1× TBS, 0.1% gelatine, and 0.2% BSA. NT buffer: 0.1 M NaCl and 0.1 M Tris-HCl, pH 9.5. NBT/BCIP (Roche). 10% polyvinyl alcohol (PVA). Harris’ hematoxylin stain. Aquamount® (BDH) or other aqueous mounting media.

2.10

Biotin-Labelled Probe Detection (with Signal Amplification)

1. Tris-buffered saline (1 × TBS): 0.15 M NaCl, 5 mM KCl, and 0.25 M Tris-HCl pH 7.5. 2. TBS-Tween: TBS with 5% Tween-20 (BDH). 3. 0.3% H2O2 in 100% methanol (v/v). 4. Blocking solution: 1.5% horse serum in TBS, 0.1% gelatin, and 0.2% BSA. 5. Amplification kit e.g. DAKO GenPoint™kit for catalyzed signal amplification containing primary streptavidin–horseradish peroxidase, biotinyl tyramide, secondary streptavidin–horseradish peroxidase (Dako, Carpinteria, CA, USA). 6. Diaminobenzidine-4HCl (DAB).

3 3.1

Methods Tissue Fixation, Processing, and Embedding

1. Place tissue biopsy in 4% paraformaldehyde/PBS, overnight at room temperature. 2. Wash the tissue in 7% sucrose/0.1 M sodium phosphate buffer overnight at 4°C. 3. Dehydrate tissue through graded alcohols 50%, 70%, 90%, 100%, and then two changes of 100% chloroform.

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4. Place tissue in molten paraffin wax (approximately 58°C) and leave tissue in wax for a minimum of 4 h. 5. Discard primary wax and replace with fresh wax and leave for 4 hours. 6. Ensuring correct orientation of the tissue, embed the tissue in wax using the stainless steel molds. Place the molds at -20°C for 1 h and then remove the wax blocks from the molds.

3.2 1. 2. 3. 4. 5. 6.

Coating Slides with APES

Place glass slides in racks and wash in an alkaline detergent overnight. Rinse the slides thoroughly with running water and then allow them to dry. Wrap the slides in aluminium foil and bake at 180°C for 3 h. Place slides in racks and immerse in a 2% solution of APES in acetone for 20 s. Rinse slides in acetone for 20 s and then in distilled H2O, twice. Dry the slides at 37°C overnight and store in an air-tight container.

3.3

Tissue Sectioning

1. Fill a small container with distilled H2O and prepare a waterbath at 42°C. 2. Cut 4- to 5-µm sections of the paraffin-embedded tissue on a microtome. 3. Place the sections into the H2O and then with an uncoated glass slide transfer the section into the waterbath. The section should flatten. 4. Mount the section with a coated slide and allow the section to dry overnight at 42°C.

3.4

Pretreatment of Tissue Sections

3.4.1

Pretreatment of tissue sections for radioactive probes

1. Dewax the sections in histolene or xylene and rehydrate through graded alcohols and finally DEPC-treated distilled H2O. 2. Microwave the sections in 1× Citra™ solution, to boiling point, add extra buffer if the level falls significantly and slides are not fully covered. Allow to cool to 37°C. 3. Rinse the sections in prewarmed (37°C) P buffer. 4. Digest tissue with pronase E in P buffer (125 µg/mL) at 37°C for 10 min. 5. Rinse twice in 0.1 M sodium phosphate buffer. 6. Post-fix the sections in 4% paraformaldehyde/PBS at room temperature for 10 min. 7. Rinse twice in 0.1 M sodium phosphate buffer. 8. Wash the sections in distilled H2O and dehydrate in 70% ethanol twice. 9. Air dry the sections and store at room temperature in a closed container until required.

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Pretreatment of Tissue for DIG or Biotin-Labelled Probes

The steps are identical to pre-treatment of tissue described above except at step 3, pronase E is incubated for 20 min.

3.5

Labelling the Riboprobe

3.5.1

Radioactive Probe Preparation

Template concentration is important in the labelling procedure. For riboprobe synthesis, 500–1,000 ng of template is recommended. 1. For one transcription reaction, the following final concentrations of reagents are required: 1 × transcription buffer, 16 mM DTT, 20 U RNasin, 400 µM ATP, 400 µM CTP, 400 µM GTP, 12 µM UTP, template (500–1,000 ng), 20 U appropriate RNA polymerase, 50 µCi 5'[α-33P]UTP, and distilled H2O to a final volume of 20 µL. 2. Incubate the reaction mixture at 37°C for 1 h in a dry heat block or waterbath. 3. Digest the template DNA with 1 U of DNase I and incubate the reaction at 37°C for a further 15 min. 4. Add 40 µg of tRNA and adjust the reaction volume to 100 µL with DEPCtreated distilled H2O. 5. Set aside 1 µL for scintillation counting. 6. Precipitate the riboprobe by adding 50 µL of 7.5 M ammonium acetate and 300 µL of 100% ethanol and place at -70°C for 20 min. 7. Pellet the riboprobe by centrifugation at 10,000×g, for 20 min at room temperature. 8. Remove the supernatant and wash the pellet with 70% ethanol. 9. Resuspend the riboprobe in 100 µL DEPC-treated distilled H2O and remove 1 µL for scintillation counting. 10. In the case of long probes, access to the target mRNA in the tissue may be limited. To improve penetration of the probe hydrolysis may be necessary. We have chosen a probe length of approximately 0.15 kb. For hydrolysis, add 100 µL of hydrolysis buffer to the riboprobe and incubate at 65°C for the appropriate length of time (see Note 2). 11. Terminate the hydrolysis reaction by adding stop buffer and then precipitate the hydrolyzed probe by adding 40 µL of 3 M sodium acetate, 40 µg tRNA, and 800 µL of 100% ethanol. 12. Precipitate as described in steps 7 and 8. 13. Resuspend in 100 µL of DEPC-treated distilled H2O, and take 1 µL for scintillation counting.

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Labelling the Probe—DIG

1. To a microcentrifuge tube add the following final concentration of reagents: 2 µg of template cDNA, 1× transcription buffer, 1× DIG labelling mix (from labelling kit), 20 U of RNasin™, (Promega), 20 U of the appropriate RNA polymerase, and DEPC-treated double-distilled H2O to a final volume of 20 µL. 2. Incubate the reaction mixture at 37°C for 2 h in a dry heat block or water bath. 3. Remove template DNA by digestion with DNase 1; see Sect. 3.5.1., step 3. 4. Precipitate DIG-labelled riboprobe by stopping the reaction with addition of 2 µL of 0.2 M EDTA, then add 2.5 µL of LiCl, 75 µL of 100% ethanol, and place at –70°C for 2 h. 5. Pellet DIG-riboprobe by centrifugation at 10,000×g for 20 min at room temperature. 6. Remove supernatant and wash pellet with 70% ethanol. 7. Resuspend pellet in 50 µL of DEPC-treated double-distilled H2O. 8. The riboprobe may require hydrolysis to ensure the correct size (see Sect. 3.5.1, step 10, also see Note 2). Add 100 µL of hydrolysis buffer and incubate at 60°C for the appropriate length of time; see also Note 2 for calculation of hydrolysis times. 9. Stop hydrolysis by adding 150 µL of neutralization buffer and 900 µL of 100% ethanol. 10. Place at –70°C for 2 h and pellet as described earlier. 3.5.3

Labelling the Probe—Biotin

Proceed as for labelling with DIG except substitute 1× biotin labelling mix for 1× DIG labelling mix (Sect. 3.5.2, step 1).

3.6

Dot Blot of Nonradioactive Probes

3.6.1

Dot Blot of DIG-Labelled Probe

Prepare serial dilutions of a DIG-labelled RNA control, available from Roche. 1. Spot 1 µL of the DIG-labelled controls and experimental probe onto a nylon filter. Make a number of dilutions of the experimental probe to gauge labelling efficiency. Probes to be spotted are diluted in a mixture of formaldehyde and SSC. 2. Fix the RNA onto the membrane by baking at 120°C for 30 min. 3. Rinse in Buffer 1. 4. Incubate the membrane in Buffer 2 for 30 min at room temperature.

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5. Incubate the membrane in anti-DIG peroxidase or anti-DIG alkaline phosphatase diluted 1:1,000 in Buffer 2 for 45 min. 6. Wash membrane in Buffer 1, twice. 7. Equilibrate membrane for 2 min in Buffer 3. 8. Wash membrane in 1× PBS, three times. 9. Using DAB according to manufacturer’s instructions, perform the color reaction by immersing the membrane in DAB/H2O2. Watch for brown color to develop. Alternatively, if using alkaline phosphatase detection, dissolve 1 tablet of fast red in Tris buffer and immerse the membrane. 10. Membranes can be washed in PBS and then kept as a record by sealing them in polythene film. 3.6.2

Dot Blot of Biotin-Labelled Probe

The procedure is the same as in Sect. 3.6.1, i.e. from steps 1–4 and steps 6–9; only step 5 differs. Instead of step 5, biotinylated probes can be easily detected on blots using avidin–biotin peroxidase complex followed by DAB detection.

3.7

Hybridization

3.7.1

Hybridization of Radioactive Probes

1. Make hybridization buffer consisting of: 1 × salt solution, 50% formamide, 10% dextran sulphate, and 360 µg/mL tRNA in a total volume of 500 µL. Five hundred microliters is sufficient for approximately 10 sections. 2. Add labelled riboprobe to the hybridization buffer at a concentration of 20 × 106 dpm per 500 µL of hybridization buffer. 3. Heat the probe/hybridization buffer mix to 85°C for 5 min before placing it on the sections. 4. Coverslip the sections and place in a humidified airtight chamber. 5. Hybridize overnight at 60°C. 3.7.2

DIG-Labelled Probes

Follow Sect. 3.7.1, steps 1–4. Hybridization with DIG-labelled probes is performed at 42°C, overnight. 3.7.3

Biotin-Labelled Probes

Follow Sect. 3.7.1., steps 1–4. Hybridization with biotin-labelled probes is generally performed at 37°C, overnight.

9 Using In Situ Hybridization to Localize Renal Gene Expression

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Post-Hybridization Washes

3.8.1

Radioactive Probes

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1. Heat wash buffer to 55°C and soak slides to remove cover slips. 2. Wash slides at 55°C for 30 min. Replace wash buffer and wash slides for a further 30 min. 3. Wash slides in three changes of RNase buffer and then incubate the sections with 150 µg/mL RNase A in RNase buffer at 37°C for 1 h, with agitation (shaking waterbath). 4. Wash the sections in 2× SSC for 45 mins at 55°C and then dehydrate through 70, 80, 90, and 100% ethanol, followed by air drying. 3.8.2

Post-Hybridization Washes—DIG

1. Heat wash buffer to 42°C and soak slides to remove coverslips. 2. Wash slides for a total of 60 min in wash solution, replacing after 30 min. 3. Rinse slides in three changes of RNase buffer and then incubate slides with RNase A (150 µg/mL) at 37°C for 1 h. 4. Wash slides in 2× SSC, 45 min at 37°C. 3.8.3

Post-Hybridization Washes—Biotin

Proceed as previously described for DIG probes except that washes are performed at 37°C instead of 42°C.

3.9

Autoradiography

1. Sections that have been hybridized with 33P-labelled probes can be placed on X-ray film (e.g. XAR-5 Kodak, Hyperfilm™), to provide an idea of the success or otherwise of the hybridization reaction. This preliminary autoradiography can also serve as a guide for exposure times required in the liquid emulsion autoradiography step. However, for small pieces of tissue or where only a few cells are labelled in the tissue section, this step may be omitted. Similarly, if a phosphorimager is available, this can be used to obtain a rapid autoradiographic result. 2. In a darkroom under safelight illumination (Ilford safelight filter number 904 or Kodak safelight filter number 2), weigh out 10 g of emulsion, add 6 mL of distilled H2O, and incubate at 42°C for 2 h to allow the emulsion to melt. 3. Pour the liquefied emulsion into a glass dipping chamber (available from Amersham) and “dip” the experimental slides ensuring all slides are coated evenly and that there are no air bubbles.

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4. Remove excess emulsion by allowing slides to drain vertically on absorbent paper in the dark. 5. Place slides into a plastic slide rack and store in a lightproof box containing desiccant. 6. Expose in the lightproof container for 10–20 days, depending on the strength of the hybridization signal.

3.10

Signal Development of Radioactive Probes

1. In a darkroom under safelight illumination, place slides in the diluted developer for 2 min with mild agitation. 2. Stop development by immersion in 0.5% acetic acid for 30 s. 3. Immerse the slides for 2 min in rapid fixer. 4. Rinse slides in running tap water for 5 min. 5. Stain slides with Harris’ hematoxylin, rinse in tap water and place in Scott’s tap water for 30 s or until hematoxylin appears blue, rinse in water and then stain with eosin. 6. Dehydrate sections through graded alcohols, rinse in two changes of histolene, and mount in non-aqueous mountant (Fig. 9.1).

Fig. 9.1 Darkfield micrograph of renal tissue probed with a 32P-labelled riboprobe for procollagen α1(I). Individual silver grains are seen as bright spots against a dark background. The use of darkfield microscopy improves the sensitivity of the technique, making it easier to distinguish specific and non-specific labelling. Aberrant expression for procollagen alpha1(I) is confined to peri-tubular interstitial cells in an experimental model of renal infection

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131

Detection of DIG-Labelled cRNA

1. Rinse slides in 1× TBS, three times. 2. Incubate the sections in 1.5% fetal calf serum made in antibody diluent, for 30 min. 3. Incubate sections in anti-DIG-alkaline phosphatase, diluted 1:250, for 1 h at room temperature. 4. Wash the slides in 1× TBS, three times. 5. Add alkaline phosphatase substrate NBT/BCIP diluted in NT buffer containing 1% PVA. 6. Color reaction should then be carried out in the dark. Sections may continue to develop color over several hours or overnight. 7. Wash sections in TBS, counterstain if desired, and mount in aqueous mountant.

3.12

Detection of Biotin-Labelled Probe (Amplification Method)

1. Rinse slides in 1 × TBS. 2. Quench slides in 0.3% H2O2 in methanol, 30 min at room temperature to remove endogenous peroxidase. 3. Wash slides in 1 × TBS, three times for 5 min each time. 4. Block sections using 1.5% serum in 1 × TBS, for 30 min at room temperature. Using reagents from a biotinyl tyramide amplification kit (e.g. GenPoint™, DakoCytomation) perform the following steps: 5. Add primary streptavidin (1:100–1:1,000) and incubate sections for 15 min at room temperature. 6. Wash slides in 1 × TBS + 0.5% Tween-20, for 5 min, three times. 7. Add biotinyl tyramide (undiluted) to sections and incubate for 15 min at room temperature. 8. Repeat step 6. 9. Add secondary streptavidin (undiluted) and incubate for 15 min. 10. Repeat step 6 (see Note 3). 11. Apply activated DAB (containing H2O2) and monitor color development. 12. Counterstain using hematoxylin, dehydrate, and mount using nonaqueous mountant.

4

Notes

1. Paraffin embedding is preferable for preservation of morphology, however, it does result in loss of mRNA and consequently reduced sensitivity of mRNA detection by in situ hybridization. In some cases frozen sectioning of unfixed tissue, post-fixation in 4% paraformaldehyde, and then in situ hybridization carried out as described in this chapter can result in markedly increased sensitivity. In other cases, frozen sectioning may allow detection of expression of

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genes that proved very difficult to detect in paraffin-embedded tissue. An example of this is interleukin (IL)-6 in muscle biopsies (11). 2. Hydrolysis formula: hydrolysis time T(min) =

length of probe (kb) - length of desired end product (kb) . 0.11 x length of probe (kb) x lengthh of end product (kb)

For example, starting with a probe that is 1.5 kb and requiring an end product of 0.15 kb, the hydrolysis time T (min): T=

1.5 - 0.15

= 54.5 min.

0.11 x1.5 x 0.15

3. The tyramide amplification system can be used to increase the sensitivity of in situ hybridization when the target mRNA is in low abundance. However, the conditions, including dilutions of the various components need to be optimized for each probe. Increasing the number of cycles of amplification may also lead to an increase in background. Even a single amplification step results in considerably stronger signal when compared with the same probe using biotin and a single streptavidin–peroxidase detection step.

References 1. Komminoth, P. (1992) Digoxigenin as an alternative probe labeling for in situ hybridization. Diagn. Mol. Pathol. 1, 142–150. 2. Komminoth, P., Merk, F.B., Leav, I., Wolfe, H.J., and Roth, J. (1992) Comparison of 35S- and digoxigenin-labeled RNA and oligonucleotide probes for in situ hybridization. Expression of mRNA of the seminal vesicle secretion protein II and androgen receptor genes in the rat prostate. Histochemistry. 98, 217–228. 3. Crabb, I.D., Hughes, S.S., Hicks, D.G., Puzas, J.E., Tsao, G.J., and Rosier, R.N. (1992) Nonradioactive in situ hybridization using digoxigenin-labeled oligonucleotides. Applications to musculoskeletal tissues. Am. J. Pathol. 141, 579–589. 4. Kerstens, H.M., Poddighe, P.J., and Hanselaar, A.G. (1995) A novel in situ hybridization signal amplification method based on the deposition of biotinylated tyramine. J. Histochem. Cytochem. 43, 347–352. 5. Darby, I.A., Evans, B.A., Fu, P., Lim, G.B., Moritz, K.M., and Wintour, E.M. (1995) Erythropoietin gene expression in fetal and adult sheep kidney. Br. J. Haematol. 89, 266–270. 6. Herbst, H., Wege, T., Milani, S., Pellegrini, G., Orzechowski, H.D., Bechstein, W.O., Neuhaus, P., Gressner, A.M., and Schuppan, D. (1997) Tissue inhibitor of metalloproteinase-1 and -2 RNA expression in rat and human liver fibrosis. Am. J. Pathol. 150, 1647–1659. 7. Darby, I.A., Bisucci, T., Hewitson, T.D., and MacLellan, D.G. (1997) Apoptosis is increased in a model of diabetes-impaired wound healing in genetically diabetic mice. Int. J. Biochem. Cell Biol. 29, 191–200. 8. Hewitson, T.D., Darby, I.A., Bisucci, T., Jones, C.L., and Becker, G.J. (1998) Evolution of tubulointerstitial fibrosis in experimental renal infection and scarring. J. Am. Soc. Nephrol. 9, 632–642. 9. Wookey, P.J., Tikellis, C., Nobes, M., Casley, D., Cooper, M.E., and Darby, I.A. (1998) Amylin as a growth factor during fetal and postnatal development of the rat kidney. Kidney Int. 53, 25–30. 10. Lindahl, P., Hellstrom, M., Kalen, M., Karlsson, L., Pekny, M., Pekna, M., Soriano, P., and Betsholtz, C. (1998) Paracrine PDGF-B/PDGF-Rbeta signaling controls mesangial cell development in kidney glomeruli. Development. 125, 3313–3322. 11. Hiscock, N., Chan, M.H., Bisucci, T., Darby, I.A., and Febbraio, M.A. (2004) Skeletal myocytes are a source of interleukin-6 mRNA expression and protein release during contraction: evidence of fiber type specificity. FASEB. J.18, 992–994

Chapter 10

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

Abstract Histochemistry is a basic technique in the analysis of cell and tissue biology. Advances in staining techniques have provided the resolving power necessary to study complex organ structures such as the kidney. In this chapter we detail standard histochemical techniques used in our laboratory to localise antigens and carbohydrates with immuno and lectin binding, respectively. Our focus is on immunohistochemistry with fixed paraffin-embedded tissue sections and its application in the research laboratory. Keywords Histology, Immunohistochemistry, Lectin, Kidney

1

Introduction

Histochemistry refers to a group of techniques that chemically label histological material. Histochemical techniques can be used to identify two main groups of molecules: proteins and carbohydrates. Proteins are readily identified with immunohistochemistry, carbohydrates are identified using the ability of lectins to bind carbohydrate groups. In this chapter we describe protocols used in our laboratory to examine both proteins and carbohydrates. The principle of immunohistochemistry is to use antibodies to histologically localise specific antigens in tissue sections. Such antibodies may be polyclonal (antisera) or monoclonal antibodies raised in culture. Polyclonal antibodies are produced by the immunisation of animals, most commonly the rabbit, and are obtained as stabilised antisera or purified immunoglobulin fractions. As the name implies, polyclonal antibodies are produced by multiple cells and are as a consequence immunologically dissimilar. Conversely, monoclonal antibodies are the product of an individual clone of plasma cells. After achieving an immune response, B lymphocytes from either the spleen or lymph node are collected and fused with non-secreting mouse myeloma cells. These hybrid cells (hybridomas) have both the capacity to produce specific antibody and longevity. Cells are propagated either in culture or in the peritoneal cavity of mice, with antibodies collected from cell culture supernatant and ascites fluid, respectively. From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_10, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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Antibody binding to tissue sections can be demonstrated with a number of different detection techniques (reviewed in ref. (1)), using both direct and indirect methods (Fig. 10.1). In direct methods, the antibody molecule or the Fab fragment of the antibody molecule is conjugated with a marker (reporter) such as peroxidase, biotin, or a fluorochrome (Fig. 10.1a). Antibody binding is then visualised by reaction with a chromogenic substrate, or in the case of fluorescent markers, by excitation with light of a specific wavelength. The usefulness of the technique is however often limited by the sensitivity and the availability of conjugated primary antibodies. In indirect methods, the primary antibody is not directly labelled, but rather detected with a second antibody to the relevant immunoglobulin. If the primary antibody is made in a rabbit for instance, the secondary antibody must be directed against rabbit immunoglobulin (Fig. 10.1b). Once again this binding is visualised by substrate reaction or fluorescence. The indirect method is more time consuming but offers important advantages, the major one being that a single secondary antibody can be used to detect a variety of primary antibodies raised in the same species. A variety of techniques have been developed to visualise antibody binding. In our laboratory, the avidin–biotin complex (ABC) method (2) is routinely used to detect primary antibody binding at the light microscope level. The procedure is based on the use of a biotinylated secondary antibody, and takes advantage of the high affinity that avidin (and streptavidin) has for biotin. After addition of a biotinylated secondary antibody, a preformed peroxidase–avidin–biotin complex is added. This binds to the conjugated antisera, and in doing so increases the number of biotin binding sites. Multiple binding sites result in amplification of the tissue signal,

a

b

c

Fig. 10.1 Schematic diagram of staining strategies. a Antigen detection with directly labelled primary antibody. b Indirect detection with labelled secondary antibody. c Principle of the avidin–biotin complex method (ABC) with a biotinylated secondary antibody. The preformed avidin–biotin complex binds to a biotinylated antibody with high affinity. The binding of these complexes to each other amplifies the signal

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Table 10.1 Lectin specificity for nephron segments Lectin

Botanical origin

Nephron specificity

Phaseolus vulgaris leukoagglutinin (PHA-L) Phaseolus vulgaris erythroagglutinin (PHA-E) Bandeiraea Simplicifolia I (BSL-I) Arachis Hypogaea agglutinin (PNA) Dolichos biflorus agglutinin (DBA)

Red kidney bean Red kidney bean

Brush border of proximal tubules, thick loop of Henle Brush border of proximal tubules

Griffonia

Some collecting ducts and vasa recta

Peanut

Distal convoluted tubules and collecting ducts Collecting ducts

Horse gram

which is useful when trying to detect low copy number antigens (Fig. 10.1c). The procedure described here is based on the Vector Laboratories Vectastain™ ABC kit. When combined with a second antibody and a different substrate, it also offers the opportunity to identify two antigens through multiple labelling. In the second part of this chapter we therefore detail a protocol for double labelling using the ABC method and alkaline phosphatase. Finally, a parallel, but distinct technique, is the use of histochemistry to detect lectin binding. Lectins are proteins or glycoproteins from plants that bind to carbohydrate moieties. Their value in renal pathology is that they can be utilised to localise and distinguish different nephron segments (Table 10.1) (3). This has proven particularly valuable in understanding the mechanisms and processes involved in nephrotoxicity. Histochemical techniques routinely use frozen and paraffin-embedded tissue. Freezing tissue results in less alteration to epitopes and therefore may offer improved staining characteristics compared with techniques based on paraffin embedding. However, in tissue with structures as complex as the glomerulus, paraffin embedding offers far more detailed morphology. In this chapter we confine our discussion to techniques that use fixed, paraffin-embedded tissue.

2 2.1

Materials Fixatives

The following is a list of commonly used fixatives. Selection of the appropriate fixative may need to be determined by trial and error, as all fixatives affect antigenicity in the tissue (4).

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Neutral-Buffered Formalin (NBF)

10% v/v Buffered formalin: 100 mL formalin (~40% aqueous solution of formaldehyde) (BDH, Poole, UK), 4 g NaH2PO4 (monohydrate), 6.5 g Na2HPO4 (anhydrous), and deionised water (dH2O) to 1 L (see Note 1). 2.1.2

Methyl Carnoys

Methyl Carnoys fixative: 60% methanol, 30% chloroform, and 10% glacial acetic acid v/v/v. 2.1.3

Mercuric Formalin (see Note 2)

1. Mercuric formalin fixative: combine 100 mL formalin (~40% aqueous solution of formaldehyde) (BDH), 9 g NaCl, 900 mL dH2O, and add HgCl with stirring until saturated. Store with residual undissolved HgCl to maintain saturation. 2. Lugol’s iodine: 2% w/v iodine in 70% ethanol. 3. 5% w/v Sodium thiosulphate in dH2O. 2.1.4

Paraformaldehyde

Four percent paraformaldehyde (PFA): Dissolve 4g of paraformaldehyde (BDH) in 100ml of 0.01 M phosphate-buffered saline (PBS) pH 7.4 by heating to a maximum of 60C with stirring. Add 5–10 drops of 1 M NaOH to clear.

2.2

Processing and Embedding

1. Microscope slides with frosted edges. 2. 3-Aminopropyltriethoxysilane (APES) (Sigma–Aldrich, St. Louis, MO, USA) (see Note 3). 3. Acetone. 4. Ethanol (95%). 5. dH2O. 6. Laboratory grade detergent e.g. Pyroneg™ (JohnsonDiversey, Sturtevant, WI, USA). 7. Slide racks. 8. Staining jars.

2.3

Immunohistochemical Staining

1. Xylene. 2. HistoClear™ (National Diagnostics, Atlanta, GA, USA) or equivalent low-toxicity hydrocarbon miscible with embedding wax and mounting media.

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3. 4. 5. 6. 7.

Ethanol (100%, 75%, 50% v/v). 30% v/v H2O2. Methanol. Antibody diluent (Dako, Glostrup, Denmark). Phosphate-buffered saline (PBS): prepare 10× stock with 1.37 M NaCl, 27 mM KCL, and 100 mM Na2HPO4 and adjust to pH 7.4 with HCl if necessary. Prepare a 1× working solution by mixing one part with nine parts water. 8. Quench solution: 10 µL 30% v/v H2O2 and 990 µL methanol. 9. Normal serum/biotinylated secondary antibody—Vectastain™ kit (Vector Laboratories, Burlingame, CA, USA). (a) Blocking serum: 150 µL stock normal serum in 10 mL of 1× PBS. (b) Biotinylated secondary antibody: 50 µL species-specific biotinylated antiIgG diluted in 10 mL of 1× PBS (see Note 4). 10. Primary antibody of interest. 11. ABC Elite™ Reagent (Vectastain™): combine 100 µL Reagent A and 100 µL B in 5 mL, prepare 30 min before use (see Note 4). 12. 2-mL Syringe and 0.2-µm syringe filter (Millipore, Billerica, MA, USA). 13. 3,3'- diaminobenzidine (DAB) chromogen substrate: dissolve 1 DAB tablet (Dako) in 10 mL of 1× PBS. Pass through a 0.2-µm syringe filter to remove undissolved material. Transfer 2 mL DAB chromogen solution to a clean test tube and add 1.5 µL 30% H2O2. Cover with foil until use, as DAB is light sensitive (see Note 3). 14. Harris Haematoxylin (Fronine, Melbourne, Australia). 15. Scott’s tap water: dissolve 20 g MgSO4.7H2O and 3.5 g NaHCO3 in dH2O and make up to 1 L (final concentration 0.08 M MgSO4, 0.041 M NaHCO3). 16. DePex™ (BDH) or other mounting media. 17. Wax pen (Dako). 18. Aluminium foil. 19. Microscope cover slips (22×40 mm). 20. Humidified chamber (e.g. plastic container lined with paper towel and kept moist with 0.01 M PBS).

2.4

Additional Reagents for Double Labelling (Alkaline Phosphatase Staining)

1. Tris-buffered saline (TBS): prepare 10 × stock with 1.37 M NaCl, 27 mM KCl, and 250 mM Tris-HCl. Prepare a working solution by dilution of one part with nine parts water. 2. 3,3'- diaminobenzidine (DAB) chromogen substrate: dissolve 1 DAB tablet (Dako) in 10 mL of 1 × TBS (see Note 5). Pass through a 0.2-µm syringe filter to remove undissolved material. Transfer 2 mL DAB chromogen solution to a clean test tube and add 1.5 µL 30% H2O2. Cover with foil until use, as DAB is light sensitive (see Note 3).

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3. Alkaline phosphatase anti-IgG, of appropriate class (Dako). 4. Alkaline phosphatase anti-alkaline phosphatase complex (APAAP) (Dakocytomation). 5. Fast Red substrate: dissolve 1× Sigma Fast Red Tablet (Sigma–Aldrich) in 1 mL of dH2O and filter through a 0.2-µm syringe filter (final composition 1.0 mg/mL Fast Red TR, 0.4 mg/mL naphthol, 0.15 mg/mL levamisole, and 0.1 M Tris-HCl buffer). 6. Aquamount™ (BDH) or other aqueous mounting media.

2.5

Antigen Retrieval (Microwave Treatment and Enzyme Digestion)

2.5.1

Microwave Pre-Treatment with Citrate Buffer (see Note 6)

1. Citrate buffer: 0.01 M citric acid monohydrate in dH2O, adjusted to pH 6.0 with 2 M NaOH, and make up to 1 L. 2. Plastic film (domestic food wrap). 3. Plastic Coplin jar (microwavable staining jar with internal ridges to hold slides apart).

2.5.2

Protease Digestion

1. Protease digest: dissolve 0.0125 g protease VIII (bacteria from Bacillus licheniformis; Sigma–Aldrich) in 50 mL of 1× PBS. 2. 0.01 M PBS. 3. Coplin jar. 2.5.3

Trypsin Digestion

1. Trypsin digest: add 100 mg trypsin (type II porcine pancreas trypsin; Sigma– Aldrich) to 100 mL of dH2O pre-heated to 37°C and stir gently. When dissolved, add 100 mg CaCl2 and adjust to pH 7.8 with 0.1 M NaOH. Keep at 37°C and use within 30 min. 2. Coplin jar.

3

Methods

The procedures below describe techniques to chemically label paraffin-embedded histological material. The methods outline preparation of sections on APES coated slides, the use of immuno histochemistry to localise proteins in situ, and finally, the

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use of lectin histochemistry to localise and detect specific nephron segments in kidney tissue.

3.1

Tissue Fixation

Fixation confers chemical stability on tissue, hardens tissue for sectioning and most importantly, halts autolysis and degradation. Fixatives preserve tissue by denaturing proteins through coagulation (e.g. methyl carnoys), cross-linking amino acids (e.g. paraformaldehyde) or both (mercuric formalin). These changes to the molecular form mean that fixation is often therefore a compromise between retention and preservation. It also alters tissue penetration and antigen exposure, which may be advantageous or disadvantageous. The appropriate fixative depends on a number of factors including the nature of the antigen and antibody, and any other anticipated usage of tissue (e.g. simultaneous study with in situ hybridisation). In each case tissue should be fixed for 4–24 h depending on the size of tissue. As a guide renal biopsy-sized tissue is fixed for 4 h, 1-cm2 portions for 18 h.

3.2

Tissue Processing and Embedding

1. Dehydrate tissue through graded alcohols (50%, 70%, 95%, and 100%) and clear in two changes of chloroform (45 min–1 h each). 2. Infiltrate tissue with two changes of paraffin wax (~56°C) for a total of 4–6 h. Excess heat should be avoided as it may potentially affect antigenicity. 3. Orientate and embed tissue in fresh wax using molds and embedding cassettes. 4. Place tissue and molds in a –20°C freezer for a minimum of 2 h before separating the block from the mold.

3.3

Preparation of APES-Coated Slides

Proper preparation of microscope slides is required to prevent the loss of tissue. We routinely coat microscope slides with APES to ensure maximum section adhesion and minimum section loss during incubation and washing (see Note 7). 1. Wash slides in laboratory detergent overnight (e.g. 20 g Pyroneg per 1 L tap water). 2. Collect slides in histology staining racks and wash in running tap water for 3 h (slides must be fully immersed). 3. Wash twice in dH2O, 5 min each. 4. Wash in freshly prepared 95% alcohol 2× for 5 min each. 5. Dry with hair drier.

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6. 7. 8. 9. 10. 11.

3.4

T.D. Hewitson, L. Grimwood

Prepare a 2% solution of APES in dry acetone (w/v) (see Note 3). Immerse slides for 10 s in a staining jar filled with APES/acetone. Wash twice in dry acetone for 5 min each. Rinse twice in dH2O. Air-dry in hot oven at 37°C for 12 h. Store in a covered dust-free container at room temperature.

Sectioning of Paraffin-Embedded Tissue

1. Heat water bath to 56°C. 2. Prepare a flotation bath by filling a small bowl with 20% v/v ethanol in water. 3. Cut 2- to 5-µm sections of paraffin embedded tissue with a microtome, in accordance with the manufacturer’s instructions. 4. Float sections on 20% ethanol to flatten and transfer to a water bath using a glass slide. 5. Collect sections on APES coated slides and stand upright to drain. Transfer to a slide rack and allow to dry overnight in an oven at 40°C.

3.5

Standard Immunohistochemistry Protocol

A flow sheet summarising the basic immunohistochemical technique is shown in Fig. 10.2. 1. De-wax in one change of xylene and two changes of HistoClear™ (National Diagnostics) for a minimum of 30 min in total. Re-hydrate sections through graded ethanol (100%, 100%, 70%, and 50%), allowing 1 min for each step. If sections are frozen or cell culture slides, immerse for 5 min in 1× PBS and start at step 4. 2. Rinse in tap water for 5 min. 3. Enzyme digest or microwave pre-treat if necessary (see Sect. 3.7). 4. If tissue has been fixed in mercuric formalin, wash sections in Lugol’s iodine for 5 min followed by 5% sodium thiosulphate for 5 min to remove mercuric chloride crystals. 5. Isolate tissue with a wax pen (Dako) (see Note 8). 6. Quench endogenous peroxidase activity by covering sections with quench solution and incubating for 20 min in humidified chamber. 7. Wash twice in 1× PBS (0.01 M PBS pH 7.4) for 5 min each. 8. Incubate sections with diluted normal serum (blocking serum; Vectastain™ ABC kit; Vector) for 10 min to block non-specific antibody binding sites. Normal serum is matched for the species in which the primary antibody was raised. 9. Blot with a tissue paper to remove serum.

10 Immuno and Lectin Histochemistry for Renal Light Microscopy Fig. 10.2 Flow sheet detailing the sequence of events in immunohistochemistry of light microscopy sections using the ABC method for localization of antibody binding. Optional steps include 1) treatment to remove mercuric chloride pigment in mercuric formalin fixed tissue, 2) enzymatic digestion, and 3) microwave treatment, as appropriate

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Fixation De-wax and re-hydrate paraffin embedded tissue sections

Digestion

Quench

Microwave

Normal serum block

Primary Antibody

Biotinylated Secondary Antibody

ABC Elite Complex

DAB

Counterstain

10. Incubate with primary antibody (diluted in antibody diluent) for 1–16 h (refer to Sect. 3.8 for a list of appropriate controls) (see Note 9). 11. Wash twice in 1× PBS for 5 min each. 12. Incubate with applicable biotinylated secondary antibody for 10 min (Vectastain™ ABC kit; Vector) (see Note 10). 13. Wash twice in 1× PBS for 5 min each. 14. Incubate with ABC Elite reagent (Vectastain™ ABC kit; Vector) for 15 min. 15. Wash twice in 1× PBS for 5 min each.

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16. Incubate with chromogen substrate for 2–10 min. Monitor the substrate reaction with a microscope, terminating the reaction by washing in dH2O for 5 min (see Note 11). 17. Wash in tap water for 5 min. 18. Counterstain in Harris Haematoxylin for 2 min. 19. Rinse in water for 5 min. 20. Dip 3 × in Scott’s tap water. 21. Dehydrate tissue by dipping sections sequentially in increasing grades of ethanol, two changes of HistoClear™ (National Diagnostics), and leave in a final change of xylene. 22. Mount with a cover slip and DePex™ (BDH) mounting media. 23. Examine under a light microscope. Antibody binding is visualised as a dark brown reaction product.

3.6

Double Labelling

Double-labelling refers to the use of two different coloured enzyme substrates that enable the differential or co-localisation of antibodies of different antigenic specificities. In this section, we detail the use of DAB and Fast Red to produce brown and red reaction products by the ABC and alkaline–phosphatase methods, respectively (see Note 12). 1. De-wax and re-hydrate sections through graded alcohols. 2. Quench endogenous peroxidase activity by covering sections with quench solution and incubating for 30 min in humidified chamber. 3. Block non-specific sites by covering with diluted normal serum for 30 min. 4. Incubate with primary antibody for 2 h at room temperature. 5. Wash twice in 1× TBS for 5 min each (see Note 5). 6. Incubate with appropriate biotinylated secondary antibody (Vector) for 10 min. 7. Wash twice in 1× TBS for 5 min each. 8. Incubate with avidin DH-biotinylated horseradish-peroxidase complex (Vectastain™ ABC kit; Vector) (5) for 15 min. 9. Wash twice in 1× TBS, 5 min each. 10. Cover sections with DAB chromogen substrate for 2–10 min. Monitor colour development by light microscopy (see Note 11). 11. Wash twice in 1× TBS for 5 min each. 12. Incubate with second primary antibody at room temperature (see Note 13) for 2 h. 13. Wash twice in 1× TBS, 5 min each. 14. Incubate for 10 min with a 1:50 dilution of alkaline–phosphatase-conjugated anti-IgG antiserum (Dako, species specific for primary antibody). 15. Wash twice in 1× TBS, 5 min each. 16. Incubate with alkaline–phosphatase anti-alkaline phosphatase (APAAP) (1:50 dilution) for 20 min.

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17. Repeat steps 14, 15, and 16 to amplify. 18. Incubate sections for 10–20 min with Fast Red™ substrate (Sigma–Aldrich). Monitor colour development on wet slides by light microscopy. 19. Rinse in running tap water. 20. Counterstain in Harris Haematoxylin for 1 min. 21. Dip in Scott’s tap water for 5 s. 22. Wash in tap water. 23. Mount with aqueous mounting media (e.g. Aquamount™; BDH) (see Note 14).

3.7

Pretreatments

In some cases antigen retrieval (or antigen recovery) may be necessary to expose or retrieve antigens masked by the tissue fixation process. There are various protocols available; some apply enzymes while others make use of heat-induced epitope retrieval. 3.7.1

Enzyme Digestion

Enzyme digestion improves accessibility of antigenic determinants. Enzymes that can be used include pepsin, pronase E, protease VIII, and trypsin. The requirement for digestion and the optimal concentration and time need to be determined by trial and error for individual antibodies. 3.7.1.1 1. 2. 3. 4.

Protease VIII Digest

Fill a Coplin jar with protease digest and pre-heat to 37°C in a water bath. Incubate de-waxed and re-hydrated sections for 3 min at 37°C. Wash in 1× PBS for 5 min. Resume at Sect. 3.5, step 4.

3.7.1.2

Trypsin Digest

1. Incubate de-waxed and re-hydrated sections in trypsin digest for 10–30 min at 37°C (see Note 15). 2. Wash in dH2O. 3. Resume standard protocol at Sect. 3.5, step 4.

3.7.2

Microwave Treatment

Pre-treatment of sections by boiling in citrate buffer using a microwave oven (6) is employed as an alternative to enzymatic digestion with some antibodies as follows:

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1. De-wax and re-hydrate paraffin-embedded sections. 2. Wash sections in dH2O for 5 minutes. 3. Place sections in a Coplin jar filled with citrate buffer and cover with plastic film. 4. Boil for 3 min. Refill the Coplin jar with buffer or dH2O at 1-min intervals (see Note 16). 5. Leave sections in the Coplin jar at room temperature for 15–20 min. 6. Wash in 1× TBS or 1× PBS (depending on intended protocol). 7. Resume protocol at step 4 (Sect. 3.5).

3.8

Controls

Inclusion of appropriate control sections is an essential part of histochemistry. 3.8.1

Specificity of Staining

Immunisation with an antigen may result in the production of contaminating antibodies, or monoclonal antibodies may have cross-reacting or non-specific binding. Control sections from other organs (e.g. spleen, lymph node) and normal renal tissue are useful for confirming specificity of staining. Substitution of the antibody with non-immune serum or an irrelevant antibody from the same species at the same protein concentration is used to confirm antibody specificity. Wherever possible, serial sections are labelled using two different antibodies to the same immunogen. Specificity is established by confirming that staining patterns are similar. If available, antibodies can also be pre-absorbed with an excess of purified immunogen overnight, with the supernatant being used for immunoperoxidase staining at the same dilution as primary antibody. Some matrix proteins, collagen in particular, are not highly immunogenic and give rise to low titer antisera. This complicates the problem of non-specific binding (7). 3.8.2

Technical Controls

Specificity of the enzymatic reaction is established by confirming that staining is absent when the primary antiserum is omitted.

3.9

Lectin Histochemistry

Although there are no specific markers for renal tubules, the distribution of lectinbinding sites can be used to characterise nephron segments (Table 10.1). Binding is easily visualised directly through the use of lectins labelled with biotin or horseradish peroxidase (HRP) (Fig. 10.3). It is however often necessary to use a

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Fig. 10.3 a 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 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™. 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 (see Color Plate 6)

combination of lectins to identify specific nephron segments (8). For example, since Arachis Hypogaea agglutinin (PNA) stains both distal tubules and collecting ducts, and Dolichos biflorus agglutinin (DBA) stains collecting ducts, those tubules only staining for PNA are distal tubules.

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1. De-wax and re-hydrate neutral buffered formalin (NBF)-fixed tissue by immersing sequentially in xylene (15 min), HistoClear™ (10 min twice) and graded ethanol (100%, 100%, 75%, and 50%; 1 min each). 2. Wash twice in 1× PBS for 5 min each. 3. Quench endogenous peroxidase activity for 20 min with 0.3% H2O2 in methanol v/v in humidified chamber. 4. Wash twice in 1× PBS for 5 min each. 5. Incubate with a 1:50 dilution of HRP-conjugated lectin diluted in antibody diluent (Table 10.1) for 1 h. 6. Wash twice in 1× PBS, 5 min each. 7. Incubate for 15 min with ABC Elite solution (Vectastain™; Vector). 8. Wash twice in 1× PBS, 5 min each. 9. Develop with DAB chromogen for 2–7 min (see Sect. 3.5, steps 16). 10. Wash in tap water for 2 min. 11. Counterstain with Harris Haematoxylin for 30 s. 12. Wash in tap water for 5 min. 13. Dehydrate in increasing grades of alcohol, Histoclear™ (National Diagnostics), and xylene. 14. Mount with DePex™ (BDH), and cover slip.

4

Notes

1. NBF is readily available commercially. 2. Mouse kidneys fixed in mercuric formalin are often too brittle to be easily sectioned. Other fixatives are recommended. 3. Consult the Material Safety Data Sheet (MSDS) before use. 4. In our experience, diluted normal serum and biotinylated secondary antibody solutions are stable for 7 days when stored at 4°C. Diluted ABC Elite kit is stable for 5 days if stored at 4°C. 5. PBS is known to inhibit the alkaline–phosphatase substrate reaction, therefore TBS is used for all reagent preparation and washes. 6. A number of excellent proprietary antigen retrieval solutions are available commercially. These vary in composition and pH. 7. Other possible coatings to use include gelatin and poly-l-lysine. In general these coatings are less satisfactory as they tend to leave smears on the slide surface and are subject to bacterial growth when stored for an extended period. 8. Providing a hydrophobic barrier around the tissue section helps to conserve reagents. It also means that with care, serial sections on the same slide can be stained with different primary antibodies. 9. Antibody dilutions may need to be determined empirically. As a guide, dilute monoclonal and polyclonal antisera to a protein concentration of about 5–20 µg/mL. Extending the incubation time can improve binding but may also increase background. 10. If primary antibody is directly conjugated with biotin or HRP, omit steps 12 and 13 (Sect. 3.5). 11. The intensity of the reaction should be monitored with a microscope to ensure that the staining is stopped before the sections turn uniformly brown. 12. Most investigators believe that the ABC technique is more sensitive than alkaline phosphatase. Therefore, wherever possible, investigators should use the higher sensitivity ABC

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13.

14. 15. 16.

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method for lower antigen density in double-labelling studies. Conversely, the alkaline– phosphatase technique is better suited to higher density antigens. Double labelling using two antibodies raised in the same species (e.g. two mouse monoclonal antibodies) can be problematic as the common secondary antibody will bind to both primary antisera. In such circumstances, it is not possible to distinguish between the two antibodies. A common solution is to use one directly conjugated antibody (e.g. biotin conjugated antiBrdU; Fig. 10.3b), in which case a secondary antibody is not required. The Fast Red reaction products are alcohol soluble therefore slides must be cover slipped with aqueous mounting media. As a guide, the optimum time for trypsin treatment is generally about 15 min for glutaraldehyde/formalin fixed tissues, and 30 min for tissue fixed in formaldehyde. It is important that sections are covered with buffer throughout microwave treatment to prevent the tissue from drying out.

References 1. Mackenzie, C.D. (1992) Immunocytochemistry. In: Roitt, I. M., Delves, P. J. (eds): Encyclopedia of immunology, London, Academic Press, pp 783–790. 2. Hsu, S.M., Raine, L., Fanger, H. (1981) Use of avidin-biotin -peroxidase complex (ABC) in immunoperoxidase technique: a comparison between ABC and unlabeled antibody (PAP) procedures. J. Histochem. Cytochem. 29:577–580. 3. Truong, L.D., Phung, V.T., Yoshikawa, Y., Mattioli, C.A. (1988) Glycoconjugates in normal human kidney. A histochemical study using 13 biotinylated lectins. Histochemistry 90:51–60. 4. Hancock, W.W., Becker, G.J., Atkins, R.C. (1982) A comparison of fixatives and immunohistochemical technics for use with monoclonal antibodies to cell surface antigens. Am. J. Clin. Pathol. 78:825–831. 5. Ellis, J., Halliday, G. (1992) A comparative study of avidin-biotin-peroxidase complexes for the immunohistochemical detection of antigens in neural tissue. Biotech. Histochem. 67:367–371. 6. Shi, S-R., Chaiwun, B., Yong, L., Cote, R.J., Taylor, C.R. (1993) Antigen retrieval technique utilizing citrate buffer or urea solution for immunohistochemical demonstration of androgen receptor in formalin fixed paraffin sections. J. Histochem. Cytochem. 41:1599–1604. 7. Bissell, D.M. (1990) Cell-matrix interaction and hepatic fibrosis. Progr. Liver Dis. 9:143–155. 8. Cachat, F., Lange-Sperandio, B., Chang, A.Y., Kiley, S.C., Thornhill, B.A., Forbes, M.S., Chevalier, R.L. (2003) Ureteral obstruction in neonatal mice elicits segment-specific tubular cell responses leading to nephron loss. Kidney Int. 63:564–575. 9. Hewitson, T.D., Wu, H., Becker, G.J. (1995) Interstitial myofibroblasts in experimental renal infection and scarring. Amer. J. Nephrol. 15:411–417.

Chapter 11

Immuno and Lectin Histochemistry for Renal Electron Microscopy Mitsuru Nakajima

Abstract The combination of histochemical techniques and electron microscopy is a powerful tool to study the mechanisms and pathology of renal disease. Through the use of electron-dense markers such as colloidal gold, biologists are able to localize immune deposits, cellular receptors, and extracellular proteins, amongst others. In this chapter, the protocols for making colloidal gold, conjugating colloidal gold to protein A, and post-embedding labeling with a protein A–gold complex are described. Finally, a parallel technique for histochemical labeling with lectin–gold complexes is provided. Keywords Histochemistry, Kidney, Electron microscopy, Colloidal gold, Protein A, Lectin

1

Introduction

For biologists, transmission electron microscopy (EM) offers unparalleled precision in examining ultrastructure. The technique has always been widely used in renal medicine where the localization and distribution of immune deposits has diagnostic significance. Not surprisingly therefore, the use of histochemical staining techniques in EM offer an important tool in the study of renal pathology. Numerous reports of immunohistochemical labeling in human renal biopsy specimens or animal kidneys have been published. The fundamental basis of all histochemistry in EM is the choice of an electrondense marker to visualize specific binding. As in light microscopy, this can be used to localize both immunological (antibodies) and chemical binding (e.g., lectins). In general, two methods have been used in renal immunoelectron microscopy: a pre-embedding method and a post-embedding method. In the former, thick sections of tissue are immunostained with peroxidase before being embedded and cut for EM. This immunoenzyme labeling is sensitive in immunoreaction but has several disadvantages: procedures are complicated, a high degree of skill in cutting sections is necessary, the penetration of antibodies is poor, and localization of signals might not be distinguishable from electron-dense deposits. From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_11, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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Post-embedding labeling with immunogold, a more recent technique, solves a number of these problems. Procedures are simple, techniques are similar to those for conventional EM, and the signal is easy to visualize, even if on the electron-dense deposits. Furthermore, simultaneous double labeling is possible. Immunoreactivities might be near equal to the pre-embedding method. Thus, immunoelectron microscopy using the immunogold post-embedding method has established itself as the method of choice in renal research (Fig. 11.1) (1–4 ). The technique is based on the appropriate combination of primary antibodies and gold particles conjugated with protein A or immunoglobulin. Protein A is a cell wall constituent produced by Staphylococcus aureus that has the ability to bind to the Fc region of mammalian IgG (5). Its binding reaction depends upon IgG subclasses and species, namely, that to IgGs from rabbit or guinea pig is strong and that from goat, sheep or mice much weaker less strong (Table 11.1). Protein A–colloidal gold is therefore particularly suitable for immunohistochemical labeling with polyclonal antibodies from rabbit or guinea pig. In the case of other mammalian species, especially mouse monoclonal antibodies, it is usually necessary to use immunoglobulin–colloidal gold conjugates as a secondary antibody. In our experience, although the preparation of high-quality, high-affinity gold–protein A complexes is straightforward, the preparation of immunoglobulin–gold is not. We therefore routinely

Fig. 11.1 Immunoelectron micrograph of protein A–gold labeling using an antibody against the complement component C9. Labeling is localized to mesangial deposits and fibrillar structures in the capillary lumen (reproduced from ref. (16 ) with the permission of S. Karger AG, Basel, Switzerland)

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Table 11.1 Relative affinity of protein A for antibodies (derived from ref. (15)) Species Human Rabbit Guinea pig Mouse Rat Goat

Relative affinity ++++ ++++ ++++ ++ +/– –

prepare protein A–gold in our laboratory and obtain immunoglobulin–gold complexes commercially. What follows below is a detailed description of protocols for preparing protein A–gold conjugates, immunogold EM, and guidelines for double labeling. Finally, a parallel technique for histochemical labeling with lectin–gold complexes is provided.

2

Materials

2.1 1. 2. 3. 4.

Preparation of Colloidal Gold

Gold chloride crystals (BDH, Poole, UK). 1% (w/v) trisodium citrate dehydrate in double-distilled H2O. 1% (w/v) tannic acid in double-distilled H2O. 25 mM potassium carbonate.

2.2

Conjugation of Colloidal Gold to Protein A

1. Protein A (Amersham, GE Healthcare Life Sciences, Little Chalfont, Buckinghamshire, UK). 2. 0.005 M NaCl. 3. 5% Polyethylene glycol (MW 20,000). 4. 0.45-µm Millipore filter. 5. 5% Glycerol in distilled H2O (v/v) containing 0.05% polyethylene glycol (w/v) and 0.01% sodium azide (w/v).

2.3

Conjugation of Colloidal Gold to Lectins

1. 0.1 M Potassium carbonate. 2. 0.1 M HCl.

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pH test paper. Lectin of interest (E.Y. Laboratories, San Mateo, CA, USA). 0.005 M NaCl. 5% Polyethylene glycol (MW 20,000). 0.45-µm Millipore filter. 5% glycerol in distilled H2O (v/v) containing 0.05% polyethylene glycol (w/v) and 0.01% sodium azide (w/v).

2.4

Tissue Fixation

2.4.1

Preparation of Paraformaldehyde–Lysine Periodate (PLP) Fixative for Protein A–Gold Labeling

1. Stock A: dissolve 1.827 g l-lysine HCl in 50 mL distilled water. After adjusting the pH to 7.4 with 0.1 M Na2HPO4 , the solution is made up to 100 mL with 0.1 M phosphate buffer pH 7.4. This solution can be stored at 4°C for up to 10 days. 2. Stock B: mix 8.0 g of paraformaldehyde in 100 mL of distilled water and heat to 60°C with stirring. One to five drops of 1 M NaOH are added slowly until the mixture turns clear. After filtration through filter paper, this solution can be stored at 4°C for up to 1 day. Paraformaldehyde is highly toxic, therefore this process should be performed in a fume hood. 3. Finally, when required, PLP is prepared by combining three parts of stock A solution with one part of stock B solution and adding sodium m-periodate to a final concentration of 0.01 M. Use within 1 day.

2.4.2

Preparation of Paraformaldehyde/Glutaraldehyde Fixative for Lectin–Gold Labeling

Prepare a solution of 2% paraformaldehyde (w/v) and 2.5% glutaraldehyde (v/v) in 0.1 M phosphate buffer, pH 7.4.

2.5 1. 2. 3. 4. 5. 6. 7.

Embedding and Section Cutting

0.1 M phosphate buffer, pH 7.4. Sucrose. Ethanol (EtOH). LR Gold resin® (London Resin Company, Reading, Berkshire, UK). Ultra violet (UV) light box for resin polymerization. Gelatin embedding capsules. 300-mesh nickel grids.

11 Immuno and Lectin Histochemistry for Renal Electron Microscopy

2.6 1. 2. 3. 4. 5. 6. 7.

Post-Embedding Labeling with Protein–Gold Complexes

Bovine serum albumin (Sigma–Aldrich, St. Louis, MO, USA). 0.01 M phosphate-buffered saline (PBS) pH 7.4. Primary antibody. Protein A–gold complex (or gold-conjugated secondary antibody). Filter paper. Laboratory wash bottle filled with distilled H2O. Saturated aqueous solution of uranyl acetate (BDH).

2.7 1. 2. 3. 4. 5. 6.

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Post-Embedding Labeling with Lectin–Gold Complex

Bovine serum albumin (Sigma–Aldrich). 0.01 M PBS pH 7.4. Lectin–gold complex. Filter paper. Laboratory squeegee bottle filled with distilled H2O. Saturated aqueous solution of uranyl acetate (BDH).

3 3.1

Methods Preparation of Colloidal Gold

Colloidal gold particles are made by chemical reduction of chloroauric acid (6 ). In our experience, 15-nm-diameter gold particles are suitable for immunocytochemical labeling of kidney sections visualized with transmission EM. Simultaneous double labeling is possible by using 5-nm gold particles in conjunction with 15-nm particles, since a threefold difference in diameter is necessary for discrimination between both sizes. In either case, the key step is the production of monodispersed gold particles without aggregates. The combined tannic acid–citrate reduction method described by Slot and Geuze (7 ) is generally accepted to be the most straightforward and reliable method for producing 3- to 17-nm colloidal gold particles. 1. An ampoule containing 1.0 g of gold chloride crystals is washed in 100% EtOH, dried, and then broken. The broken ampoule, still containing the gold chloride crystals, is immersed in 50 mL of double-distilled water and stored in the dark as 2.0% gold chloride solution. 2. Freshly prepare a gold solution by adding 0.5 ml of 2.0% gold chloride solution to 79.5 mL of double-distilled water in a well-cleaned Erlenmeyer glass flask (see Note 1). 3. Heat the solution to 60°C in a hot-water bath with stirring.

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4. Freshly prepare the reducing mixture. Combine 4 mL of 1% trisodium citrate dehydrate and 16 mL of double-distilled water for 15-nm colloidal gold particles, or 4 mL of 1% trisodium citrate dehydrate, 1 mL of 1% tannic acid, 1 mL of 25 mM potassium carbonate, and 14 mL of double-distilled water for 5-nm colloidal gold particles. 5. Quickly add the reducing mixture to gold solution while stirring (see Note 2). 6. Wait until colloid formation is complete. The color of the solution will change progressively, with the color of the solution indicating particle size. In the case of 15-nm particles, wait for solution to turn bright red (~1 h). For the 5-nm solution, wait for a bright red color (appears within minutes). 7. Heat the solution to boiling point when the appropriate color is reached.

3.2

Conjugating Colloidal Gold to Protein A (6, 8, 9)

Although colloidal gold particles in isolation are unstable, they can form stable complexes with proteins. The binding between the gold and protein molecules is due to electrostatic absorption, where the negative charge of the gold particles interacts with positive charge of the protein. Thus, conjugating colloidal gold to protein A by electrostatic absorption can be used to produce an immunohistochemical marker. The complex is stable for several months. 1. 100 to 200 µg of protein A—i.e., enough to stabilize 5 mL of colloidal gold solution in our experience—is transferred to a plastic container and dissolved in 0.1 mL 0.005 M sodium chloride. To this, 5 mL of colloidal gold solution is added. After 1 to 2 min, stabilize the gold probe by adding 0.15 mL of 5% aqueous polyethylene glycol solution drop-wise through a 0.45-µm Millipore filter. 2. Purify and concentrate the crude protein A–gold complex by ultracentrifugation. Spin 15-nm colloidal gold at 55,000×g for 40 min at 4°C on a 2.5-mL cushion of 5% glycerol containing 0.05% polyethylene glycol and 0.01% sodium azide, filtered through a 0.45-µm Millipore filter. For 5-nm colloidal gold, spin at 125,000×g for 60 min on the same cushion. 3. After centrifugation, about 6.5 mL of water-clear supernatant is carefully aspirated and discarded, leaving about 1 mL of condensed solution to be stored at 4°C until use.

3.3

Post-Embedding Labeling with Protein A–Gold Complex (1, 3, 4 )

For the purpose of post-embedding immunohistochemical labeling, the choice of resin (see Note 3) and fixative (see Note 4) is very important, since preservation of both antigenicity and ultrastructure are often mutually exclusive. In our studies of both human biopsy tissue and animal renal tissue, experience suggests that

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periodate–lysine–paraformaldehyde (PLP) fixative (10) and LR Gold® resin are the most suitable and straightforward. 3.3.1

Tissue Collection, Processing, and Embedding

1. Small pieces of human or animal kidney tissue are fixed in PLP solution for 2 h at 4°C. 2. Wash with 0.1 M phosphate buffer pH 7.4 containing 7% sucrose three times for 10 min each. 3. Dehydrate in graded EtOH (50%, 75%, 95%, and 100%) for 10 min each. 4. Immerse tissue in equal parts of 100% EtOH and LR Gold® resin (London Resin) for 30 min, followed by pure LR Gold® resin for 30 min. 5. Embed tissue in fresh LR Gold® (London Resin) and polymerize at 4°C for 24 h in sealed gelatin capsules using UV irradiation. 6. Use standard EM techniques to cut survey sections and select an area of interest. 7. Cut ultrathin sections (silver to pale gold in color) and collect on uncoated 300mesh nickel grids (see Note 5). 3.3.2

Immunogold Labeling

1. Incubate sections by inverting grid (section face down) on a drop of 0.1% bovine serum albumin (BSA) in 0.01 M PBS pH 7.4 (hereafter referred to as BSA/PBS) for 5 min at room temperature. Transfer grids consecutively to a drop of the reagents in steps 2–5 (see Note 6). 2. Appropriately diluted primary antibody (for 30 min at room temperature or for 24 h at 4°C). 3. Three drops of BSA/PBS (wash) for 5 min each. 4. Appropriately diluted protein A–colloidal gold complex (or appropriate immunoglobulin–colloidal gold complex) for 30 min at room temperature (see Notes 7 and 8). 5. Three drops of BSA/PBS (wash) for 5 min each. 6. Rinse grids with running distilled water from a squeegee bottle, and dry. 7. Stain sections for 5 min with a saturated aqueous solution of uranyl acetate, using routine techniques. 8. Examine by transmission EM (see Note 9).

3.4

Double Cytochemical Labeling with Combination of Post-Embedding Method and Protein A–Gold Complex (11)

For simultaneous identification of two different antigens in the same section, double labeling is performed using 15-nm and 5-nm colloidal gold particles. Again the choice of embedding media is important. LR Gold® (London Resin) is stable in an

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electron beam and requires no supporting film on the grid. Immunostaining can therefore take place through both sides of the section. 1. Ultrathin sections are first labeled with the primary antibody followed by protein A–15-nm gold complex as described above. In this step, it is important to take care that the section is stained by placing an uncoated nickel grid face down onto the top of a drop of the reagent. Do not let the grid sink into the reagent. 2. Repeat the process by labeling from the opposite side of the grid by placing the grid face down onto the second primary antibody followed by protein A–5-nm gold, and complete as above (see Sect. 3.3.2, steps 5–8).

3.5

Post-Embedding Labeling with Lectin–Gold Complex (12)

As described in the previous chapter, lectins are glycoproteins with the ability to bind to carbohydrates. Lectin–gold complexes can be easily prepared in the laboratory, and are useful tools for cytochemical tools for localization of specific carbohydrates in human or animal kidney tissue.

3.5.1

Preparation of Gold–Lectin Conjugates

1. The pH of the gold solution is adjusted to the isoelectric point for each lectin (Table 11.2) with 0.1 M potassium carbonate or 0.1 M hydrochloric acid. The pH should be measured by pH test paper such as litmus paper, since nonstabilized colloidal gold will plug the pore of a pH meter electrode. 2. 100 to 200 µg of lectin—i.e., enough to stabilize 5 mL of colloidal gold solution in our experience—is weighed into a plastic container and dissolved in 0.1 mL of 0.005 M sodium chloride. To this, 5 mL of pH-adjusted colloidal gold solution is added. After 1–2 min, 0.15 mL of 5% polyethylene glycol solution is added to stabilize the gold probe. 3. Crude lectin–gold complexes are purified and concentrated by ultracentrifugation. Spin 15-nm colloidal gold at 55,000×g for 40 min at 4°C on a 2.5-mL cushion

Table 11.2 Isoelectric point of various lectins Lectin Dolichos biflorus agglutinin Griffonia simplicifolia IB(4) Griffonia simplicifolia lectin II Helix pomatia agglutinin Arachis Hypogaea (peanut lectin agglutinin) Glycine max (soybean) Ulex europaeus

Abbreviation DBA GSI-B4 GS-II HPA PNA

Isoelectric point (pH) 5.0 5.0 5.0 7.4 6.3

SBA UEA-I

6.1 6.3

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of 5% glycerol containing 0.05% polyethylene glycol and 0.01% sodium azide, filtered through a 0.45-µm Millipore filter. Five-nanometer gold complexes are spun at 125,000×g for 60 min, on the same cushion. 4. After centrifugation, about 6.5 mL of water-clear supernatant is carefully aspirated and discarded, leaving about 1 mL of condensed solution to be stored at 4°C until use.

3.5.2

Preparation of Tissue

1. Small pieces of human or animal kidney tissue are fixed in paraformaldehyde/ glutaraldehyde for 2 h at 4°C. PLP fixation is not suitable for lectin labeling because it modifies the carbohydrate. 2. Wash in PBS, dehydrate, embed, and cut tissue as above (Sect. 3.3.1).

3.5.3

Lectin–Gold Labeling

1. Pretreat sections by placing grids, section down, on a drop of BSA/PBS for 5 min at room temperature. 2. Transfer to a drop of lectin–gold complex solution for 30 min at room temperature (see Note 7). 3. Wash three times by transferring consecutively to three drops of BSA/PBS (5 min each). 4. Use a wash bottle to rinse sections in running distilled water. 5. Use filter paper to blot excess water, and leave to dry. 6. Stain sections with uranyl acetate for 5 min. 7. Examine using transmission EM.

4

Notes

1. Many investigators suggest that only siliconized glassware should be used in making the gold solution, because unstable colloidal gold will adhere to the uncoated glass. In our experience however, thoroughly cleaned nonsiliconized glassware is adequate. 2. The formation of protein–colloidal gold complex is pH dependent, with optimal protein A–gold complex formation occurring at pH 5.9–6.2 (8). Reduction of chloroauric acid occurs at pH 6.0, therefore pH adjustment of the gold colloid is not necessary for protein A–gold complex formation. 3. Although the epoxy resins have been used extensively as embedding medium for conventional EM, they are not suitable for immunoelectron microscopy since they exhibit low water adsorption and many hydrophobic structures, resulting in poor immunoreaction with antibodies. On the other hand, acrylic resins such as Lowicryl® (Polysciences Inc., Warrington, PA, USA) and LR Gold® (London Resin) have highly hydrophilic characteristics with good immunohistochemical labeling properties. Ultrathin sections permit full penetration of aqueous solutions such as antibodies. Furthermore, these acrylic resins are polymerized by exposure to ultraviolet

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light at low temperature, therefore preservation of antigenicity is much better than that in heat-cured epoxy resins (13). Having said this, in our experience, the heat-cured resin LR White® (London Resin) provides for good immunohistochemical labeling of IgA, IgG, IgM deposits, complement components, and fibrinogen in human renal biopsy tissue, provided the cure temperature is kept below 60°C. Choice of fixative is very important for immunoelectron microscopy, since preservation of both antigenicity and ultrastructure is required. PLP has been proposed as a fixative for glycoprotein antigens that should stabilize periodate-oxidized polysaccharide chains through lysinemediated crosslinks, either directly or by the intermediation of formaldehyde. Thus, PLP is recommended for immunoelectron microscopy of kidney tissue. Although postfixation in osmium is generally avoided because of its adverse effect on antigenicity, we have successfully preserved antigenicity while improving ultrastructure by limiting en bloc osmium staining to 10 min (14 ). Note that PLP fixative is not suitable for histochemical labeling with lectin–gold complex. The copper grids used in conventional EM can corrode in the labeling process. Nickel grids are therefore routinely used in post-embedding labeling. In each step, nickel grids should be mounted section face down onto a drop of reagent. Before transferring to the next reagent, use a piece of filter paper to gently blot the reagent from the edge of the grid. Drying the grid adequately is essential to reduce background and prevent the grid sinking into the reagent. Appropriate dilution of protein A–gold or lectin–gold complex depends on the stock concentration. As a guide, red-colored protein A–gold and lectin–gold complexes should be diluted to a pale pink color for use. Gold-conjugated secondary antisera are used when the primary antiserum binds poorly to protein A (e.g., mouse monoclonal primary antisera). We have achieved good results with gold conjugates from Amersham. Protein A binds to the Fc region of mammalian IgG. Although it seems theoretically possible that protein A will bind to the native IgG immune complexes in electron dense deposits, this has not been reported.

References 1. Nakajima, M., Hirota, T., Kusumoto, K., Taira, K., Kamitsuji, H. (1987) Immunoelectron microscopic study of glomerular lesions using a postembedding method with a protein A-gold complex. Nephron. 46, 182–187. 2. Smith, P.S. (1988) Ultrastructural immuno-gold localization of immune deposits in human renal biopsies. Pathology. 20, 32–37. 3. Nakajima, M., Hewitson, T.D., Mathews, D.C., Kincaid-Smith, P. (1991) Platelet-derived growth factor mesangial deposits in mesangial IgA glomerulonephritis. Nephrol Dial Transplant. 6, 11–16. 4. Akazawa, H., Nakajima, M., Nishiguchi, M., Yamoto, Y., Sado, Y., Naito, I., Yoshioka, A. (2005) Quantitative immunoelectron-microscopic analysis of the type IV collagen alpha1–6 chains in the glomerular basement membrane in childhood thin basement membrane disease. Clin Nephrol. 64, 329–336. 5. Goudswaard, J., van der Donk, J.A., Noordzij, A., van Dam, R.H., Vaerman, J.P. (1978) Protein A reactivity of various mammalian immunoglobulins. Scand J Immunol. 8, 21–28. 6. Hughes, D. (2005) Preparation of colloidal gold probes. In: Methods Mol Biol. Vol. 295: Immunochemical protocols, 3rd edn. (Burns R. ed.) Humana Press, Totowa, NJ, pp. 155–172. 7. Slot, J.W., Geuze, H.J. (1985) A new method of preparing gold probes for multiple-labeling cytochemistry. Eur J Cell Biol. 38, 87–93.

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8. Roth, J. (1984) The protein A-gold technique for antigen localization in tissue sections by light and electron microscopy. In: Immunolabelling for Electron Microscopy (Polak, J.M. and Varndell, I.M., eds.) Elsevier Science Publishers, Amsterdam, The Netherlands, pp. 113–121. 9. Tanaka, H., Haga, S., Takatsuki, K., Yamaguchi, K. (1984) Localization of adult T-cell leukemiaassociated antigens by the immunocolloidal gold method. Cancer Res. 44, 3493–3504. 10. McLean, I.W., Nakane, P.K. (1974) Periodate-lysine-paraformaldehyde fixative. A new fixation for immunoelectron microscopy. J Histochem Cytochem. 22, 1077–1083. 11. Bendayan, M., Stephens, H. (1984) Double labeling cytochemistry applying the protein A-gold technique. In: Immunolabelling for Electron Microscopy (Polak, J.M. andVarndell, I.M. eds.) Elsevier Science Publishers, Amsterdam, The Netherlands, pp. 143–154. 12. Nakajima, M., Ito, N., Nishi, K., Okamura, Y., Hirota, T. (1988) Cytochemical localization of blood group substances in human salivary glands using lectin-gold complexes. J Histochem Cytochem. 36, 337–348. 13. Causton, B.E. (1984) The choice of resins for electron immunocytochemistry, in Immunolabelling for Electron Microscopy. (Polak, J.M. and Varndell, I.M. eds.) Elsevier Science Publishers, Amsterdam, The Netherlands, pp. 29–36. 14. Nakajima, M., Mathews, D.C., Hewitson, T., Kincaid-Smith, P. (1989) Modified immunogold labelling applied to the study of protein droplets in glomerular disease. Virchows Arch A Pathol Anat Histopathol. 415, 489–499. 15. Harlow, E., Lane, D. (1988) Antibodies: A laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, USA. 16. Nakajima, M., Hewitson, T.D., Mathews, D.C., Forbes, I., Kincaid-Smith, P. (1991) Atypical structured glomerular deposits: An immunohistochemical study. Am J Nephrol. 11, 151–156.

Chapter 12

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

Abstract Immunohistochemistry for pimonidazole adducts serves to define hypoxia within tissues. For this purpose, pimonidazole is delivered in vivo, binds to thiol groups at oxygen tensions below 10 mmHg, and is visualized with help of commercially available anti-pimonidazole antibodies. Renal parenchymal oxygen distribution is highly variable under normal conditions and during acute renal failure and chronic renal disorders. Pimonidazole immunostaining clearly helps in delineating hypoxic regions within the kidneys, but technical pitfalls should be taken into account. In particular, tissue fixation by in vivo perfusion is strongly recommended in order to eliminate artificial staining, because immersion fixation per se can promote a hypoxic environment within kidney tissue.

1

Introduction

Pimonidazole (PIM; Fig. 12.1) belongs to a group of 2-nitro-imidazole compounds (also named bioreductive agents) that were originally designed as radiosensitizers. At oxygen tensions below 10 mmHg, pimonidazole is reduced by 2-nitro-reductase (1, 2) and binds to thiol groups. Such pimonidazole–protein adducts are immunogenic, and specific antibodies raised against them allow us to visualize tissue hypoxia with the help of immunohistochemistry (3, 4). However, it is important to recognize that hypoxia may already occur at tissue oxygen tensions higher than 10 mmHg (5), and therefore a negative stain can not rule out hypoxia. In the kidney, low or very low tissue oxygen tensions exist under physiologic conditions (6–12) as a consequence of mechanisms/structural relationships that allow urine to be concentrated. Oxygen electrode measurements enable us to roughly draw an “oxygen map” of the normal kidney (Fig. 12.2) with a corticomedullary gradient. However, it is noteworthy, due to the limited spatial resolution of Clark-type oxygen electrodes, that such measurements may underestimate the degree of hypoxia at the cellular level. There is considerable variation of oxygen electrode recordings in the cortex (6–12), and, indeed, anatomic studies indicate that the medullary rays may receive far less oxygen than the labyrinth, since the former are supplied by venous blood originating from ascending vasa recta (13). From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_12, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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N

NO2 N

OH

N+ H Fig. 12.1 Structure of pimonidazole hydrochloride

Fig. 12.2 Probable oxygen gradients in the normal kidney (according to oxygen electrode measurements, anatomic studies, and pimonidazole staining). Note that pimonidazole staining only visualizes hypoxia <10 mmHg, and therefore suspected oxygen gradients are not fully represented by pimonidazole immunohistochemistry. The depth of grey scale corresponds with the degree of hypoxia

Cl−

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Moreover, several lines of evidence suggest that even in the physiologically hypoxic renal outer medulla, oxygenation is unevenly distributed, and gradients exist between vascular bundles and the interbundle zone, in which there is high oxygen consumption for tubular salt reabsorption (14–16). Pimonidazole immunohistochemistry has confirmed renal oxygen gradients (15–27) as they had been predicted by oxygen electrode recordings (6–12), anatomic studies (13), and renal injury models (in which medullary tubular damage increases in the mid interbundle zone, in parallel with increasing distance from the oxygen supplying vasa recta, the prime oxygen source in the medulla (14–16)). In general, the different techniques available for hypoxia imaging (e.g., immunohistochemistry for pimonidazole (15–27) and hypoxia-inducible factors (HIFs) (28, 29), oxygen electrode measurements (6–12), blood oxygen level-derived (BOLD) magnetic resonance imaging (MRI) (30, 31)) should be regarded as complementary, since they are likely have different sensitivities and specificities.

2 2.1

Materials (See Note 1) Delivery of Pimonidazole In Vivo

1. Pimonidazole hydrochloride is purchased from Natural Pharmacia International, Belmont, MA. The dry substance is dissolved in 0.9% NaCl. We recommend a 100 mg/mL stock solution. 2. Anaesthetic: ether

2.2

Retrograde Perfusion Fixation of Kidneys Via the Infrarenal Aorta

1. Fixation solution: freshly depolymerized paraformaldehyde (3%) is prepared by combining 30 g paraformaldehyde powder, 100 mL of 10× concentrated phosphatebuffered saline (PBS), 500 to 600 mL water, and 1 to 2 mL of 1 M NaOH. Total volume should be around 700–800 mL. Heat the solution to 50–60°C under a laminar flow hood with continuous magnetic stirring. Use a thermometer to control the temperature. Boiling must be avoided. When the paraformaldehyde solution is clear, cool the solution on ice until it reaches room temperature. Add water up to a volume of 900 mL. Adjust the pH to 7.4 with help of 1 M and 0.1 M HCl. Make up to 1,000 mL with water (see Note 2). 2. Rinsing solution: PBS at pH 7.4 (see Note 3). 3. Anesthetic: sodium pentobarbital in 0.9% NaCl (6% w/v) (see Note 4). 4. Perfusion cannula: the largest possible cannula should be chosen, since perfusion quality is improved by large-diameter cannulas. For animals heavier than 180 g, a 16-gauge commercially available venous cannula can be used (e.g., from Becton

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Fig. 12.3 Perfusion cannula: 1) obturator, 2) tape wrapped around the cannula, 3) hole in the tape (see Sect. 2.2, step 4 and Sect. 3.2, steps 3 and 4)

and Dickinson, Heidelberg, Germany; or B. Braun, Melsungen, Germany). The diameter of the cannula should be slightly larger than the diameter of the infrar-enal aorta, since the vessel is elastic. Large cannulas have the additional advantage that they are tightly enclosed by the aorta and no suture is needed to keep it in place. Wrap a strong tape around the cannula, at about 1 cm from the can-nula tip (Fig. 12.3). The tape will assure that the cannula does not slip too deeply into the aorta (thus possibly obstructing the renal arteries), and will not slip out of the aorta during high-pressure perfusion. For this latter purpose, make a small hole into the tape and fix it with a needle (see Sect. 3.2, step 4). To assure that no air bubbles enter the perfusion cannula and that coagulation does not occur, fill the cannula with heparin. 5. Surgical instruments and additional materials: cork platform (~20 × 20 × 1 cm), scissors, two to three microsurgery hooks, gauze dressings (10 × 10 cm), cotton swabs, blunt clamp (~10 cm), microsurgical blunt tweezers, microsurgical clamp, and microsurgery scissors.

2.3

Pimonidazole Immunohistochemistry

1. Water-repelling immunostaining pen (PAP PenTM; G.Kisker, Steinfurt, Germany). 2. Catalyzed signal amplification (CSATM) kit (DAKO, Hamburg, Germany) (see Note 5).

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3. Pronase E 0.05% (Merck, Darmstadt, Germany) in PBS with 0.2% BrijTM 35 (Merck). 4. Humidified incubation chamber (see Note 6). 5. Tris-buffered saline wash buffer (TBST): 0.05 M Tris-HCl pH 7.6, 0.3 M NaCl, and 0.1% Tween 20. 6. Anti-pimonidazole antibody (Natural Pharmacia International, Belmont, MA, USA).

3 3.1

Methods Delivery of Pimonidazole In Vivo

1. One hour prior to sacrifice, pimonidazole is injected at 6 mg/100 g either intravenously (tail vein) or intraperitoneally under brief ether anesthesia. We recommend that the injection volume be 0.5–0.7 mL (diluted in 0.9% NaCl) (see Note 7).

3.2

Retrograde Perfusion Fixation of Kidneys Via the Infrarenal Aorta

1. Regulation of perfusion pressure: fill both the rinsing and the fixation solution into separate bottles of at least 1,000 mL. Connect the two bottles with a Y-shaped line containing a three-way stopcock to rapidly switch between the solutions (Fig. 12.4). Lift the bottles 2m above the bench where the perfusion is about to be done. Fill the fixation line, then fill the rinsing line. Make sure that no air bubbles are in the lines (see Note 8). 2. Anesthesia: induce anesthesia with an intraperitoneal injection of pentobarbital at 1 to 2 µL/g body weight (250 to 500 µL per 250-g animal) (see Note 9). 3. Cannulation of the infrarenal aorta: position the rat on the cork platform. Perform a mid-abdominal incision to open the peritoneal cavity. Make additional transverse incisions at both sides of the lower abdomen to improve access to the infrarenal aorta. Gently move aside the intestinum with the help of gauze dressings moistened with saline (37°C), and fix them with the help of hooks. Expose the infrarenal aorta and caval vein with the help of cotton swabs. For optimal results stay caudal of the renal pedicles, since traction of the renal pedicles may cause vasoconstriction and impair perfusion. Fasten a blunt clamp at the left iliac artery about 1 to 2 mm below the aortic bifurcation, such that the weight of the clamp lifts up the aortic bifurcation. Hold the right iliac artery with blunt microsurgical tweezers and gently drag it ventrally, such that the aortic bifurcation is optimally exposed. Insert the cannula into the aortic bifurcation and push it about 1 cm into the aorta. Clamp the aorta above the end of the inserted cannula with the help of

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Fig. 12.4 Perfusion fixation. Phosphate-buffered saline (PBS; rinsing solution); paraformaldehyde (PFA; fixation solution). Using a three-way stopcock enables the operator to quickly switch from rinsing to fixation solution, thus avoiding a drop of perfusion pressure

a microsurgical clamp. Remove the metal obturator of the cannula and attach the cannula to the perfusion line. Be sure that the cannula is free of air bubbles (see Note 10). 4. Fix the cannula by inserting a long (at least 5 cm) 20-gauge needle/cannula sagittally through the whole in the tape wrapped around the perfusion cannula (see Fig. 12.3), directed beside the animal’s spine and anchored in the cork platform (see Note 11). 5. Perfusion fixation: open the inferior caval vein with a 3- to 5-mm incision using microsurgical scissors (see Note 12). Open the three-way stopcock for the rinsing solution immediately thereafter. After 20 to 30 s of rinsing, quickly switch the stopcock to the fixation solution. Perfusion fix for 5 min. Muscle fibrillations may occur for a few seconds between 30 and 90 s after the start of perfusion fixation. This phenomenon seems to be triggered either by direct action of the fixation solution on the striate muscles or by convulsions and should not be misinterpreted as insufficient anesthesia. Be prepared to secure the perfusion cannula in case muscle fibrillation occurs. 6. Harvest the kidneys: gently remove the kidneys (see Note 13). They should be blood free and of rubber-like consistency. Make transverse cuts. Try to obtain “long papillas,” since pimonidazole staining is strongest in the lower papilla. Kidney slices should be kept for an additional 2 to 4 h in the fixation solution at 4°C, after which they should be transferred into ice-cooled PBS and processed for paraffin embedding.

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The protocol may be adapted to the available technical resources, surgical skills, and personal preferences. However, take care with the following: be as quick as possible (to avoid anaesthesia-induced hypoxia); the animal should be deeply anesthetized; perfusion pressure must be constant; quickly switch between rinsing solution and fixation solution; perfusion quality should be evaluated on routine staining; tubules should be expanded and the brush border of proximal tubules intact (Fig. 12.5). With suboptimal perfusion, the tubules of the inner stripe and medullary rays are usually collapsed (Fig. 12.5). Note that perfusion may be more difficult in diseased kidneys than in healthy kidneys.

Fig. 12.5 Perfusion quality. Good perfusion quality is demonstrated by expanded tubules and intact brush border of proximal tubules (asterisks). Arrows point to collapsed tubules (mostly proximal tubules). The latter demonstrate poor/inhomogeneous perfusion. The outer stripe of the outer medulla (c, d), medullary rays (a), and parts of the labyrinth (b) are particularly prone to insufficient perfusion. Any immunohistochemical staining that predominantly occurs in such collapsed tubules is likely to be an artifact

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Pimonidazole Immunohistochemistry

1. Use 3-µm paraffin sections (see Note 14). 2. Deparaffinize paraffin sections by immersing for 5 min each in the following solutions: three times in xylene, twice in 100% ethanol, twice in 96% ethanol, once in 90% ethanol, once in 70% ethanol, once in 50% ethanol, and once in water (see Note 15). 3. Pronase digestion: preheat (~30 min) a Coplin jar with 0.05% pronase E in PBS with 0.2% BrijTM 35 at 40°C in an incubator. Transfer the freshly deparaffinized slides into the pronase solution, and digest for 15 min. Transfer the slides into TBST at room temperature (see Note 16). 4. Encircle the sections with a water-repelling immunostaining pen (Pap PenTM) while holding the slides horizontally. Wipe away the fluid around the sections with the help of blotting paper. Leave a 2- to 4-mm space around the sections. Take care that the sections stay moist at all times. Avoid dipping the pen into the moist area surrounding the sections. 5. Perform the following steps according to the CSATM protocol (with incubation steps of 15 min and two washing steps of 5 min in TBST, unless stated otherwise). 1) biotin block; 2) peroxidase block; 3) protein block (do not rinse); and 4) anti-pimonidazole antibody at a dilution of 1:1,000 in DAKO antibody diluent (see Note 17), keep for 1 h at room temperature, then overnight at 4°C in a humidified chamber. Apply 5) secondary biotinylated anti-mouse antibody; 6) streptavidin–biotin complex; 7) amplification reagent; 8) streptavidin–peroxidase; and 9) chromogen (diaminobenzidine). Stop chromogen formation after 0.5 to 2 min by washing the slides twice in water (see Note 18). Mount sections with cover slips.

3.4

Analysis and Interpretation

Most of the pimonidazole staining protocols published thus far (19–27) use neither perfusion fixation (which improves morphologic assessment and reduces artificial staining) nor the amplification technique (which allows use of much higher antibody dilutions, thus increasing specificity) presented herein. Interestingly, immersionfixed kidneys stained with the nonamplified technique show far more pimonidazole signal than perfusion-fixed kidneys stained with the amplification technique (Table 12.1). Preliminary results from our laboratory suggest that the disparity relates to hypoxia generation during immersion fixation. When using the amplification technique in perfusion-fixed control rat kidneys, signals were detectable in some, but not all, papillas and deep portions of the inner stripe of the outer medulla (Fig. 12.6). Using the amplification technique in immersion-fixed control rat kidneys led to a pattern very similar to the one obtained by the nonamplified technique in immersion-fixed kidneys: signals were stronger and much more abundant in the papilla and in the inner stripe of the outer medulla. Additional signals appeared in

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Table 12.1 Pimonidazole staining in control rat kidneys Pimonidazole staining in control rat kidneys Perfusion fixation

Immersion fixation

Renal zone

Amplified

Non-amplified

Amplified

Non-amplified*

Cortex

–

n.d.

Outer stripe

–

n.d.

Inner stripe

+/– n.d. (deep mTALs, ascending vasa recta) +/– n.d. (low papilla, mostly IC/EC, some CDs)

+ (medullary rays) ++ (S3) +++ (mTALs, CDs)

++ (medullary rays) +++ (S3) +++ (mTALs, CDs)

Papilla

+++ +++ (almost entire (almost entire papilla, all cells) papilla, all cells)

CDs collecting ducts; IC/EC interstitial cells/endothelial cells; mTALs medullary thick ascending limbs of the loop of Henle; n.d. not done; S3 S3 segment of the proximal tubule. Staining intensity: − no staining; +/− weak and inconsistent staining (in less than 50% of sections); + weak but consistent staining; ++ moderate staining; +++ strong staining *As presented in the literature (19–27)

Fig. 12.6 (continued)

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Fig. 12.6 (continued) Pimonidazole staining in normal kidneys. Immunohistochemical staining for pimonidazole adducts (PIM) using the amplified method. a arcuate artery; med. ray medullary ray; v arcuate vein; vb vascular bundle; dotted line in (c) marks the border between the low cortex (upper part) and outer stripe; dotted lines in (e) and (f) mark the border between the deep inner stripe (upper part) and the inner medulla/papilla. Perfusion-fixed (left-hand side) opposed to immersion-fixed (right-hand side) control rat kidneys. Immersion-fixed kidneys show stronger and more abundant staining in all renal zones: S3 portions of proximal tubules in the medullary rays (b) and outer stripe (d), virtually all cells, but predominantly thick ascending limbs of the loop of Henle (mTALs) in the inner stripe (f), and virtually all cells of the papilla (h). By contrast, with perfusion fixation, signals inconsistently appear in the papilla (mostly interstitial cells, g) and in the deep inner stripe (mTALs vascular bundles—probably ascending vasa recta, e). Compare with Table 12.1

the outer stripe of the outer medulla and in medullary rays (which both contain S3 segments of the proximal tubule). Possibly, collapsed S3 segments nonspecifically bind the primary anti-pimonidazole antibody. Thus, immersion fixation emphasizes hypoxic patterns recognized in a properly fixed kidney as well as showing new sites of staining. Such a staining pattern is likely the result of the additional hypoxia that occurs in kidneys fixed by this method. In contrast, perfusion fixation likely occurs

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almost instantaneously and homogeneously. We recommend the amplified technique in perfusion-fixed kidneys, since it seems to have the highest specificity so far. In each experimental series, sufficient control kidneys should be run in order to determine baseline pimonidazole staining.

4

Notes

1. All solutions should be prepared in water that has a resistance of 18.2 M-Ω and a total organic content of less than five parts per billion. This standard is referred to as “water” in this chapter. 2. If the paraformaldehyde does not dissolve completely, add 1 M NaOH. The fixative solution is stable for 48 h at 4°C (protect from light when storing). 3. Alternatively, 4% hydroxy–ethyl–starch in PBS (pH 7.4) can be used. 4. Pentobarbital solution is stable for 48 h at 4°C. 5. Do not use the CSATM kit beyond the expiration date. Make aliquots of 500–750 µL of each kit reagent. Keep all reagents at 4°C. The aliquoted reagents should equilibrate at room temperature for at least 1 h before starting the staining procedure. The kit may loose some activity starting 1 to 2 months before the expiration date, which can lead to weaker signals. In our experience, this is only a problem when the amount of antigen is very low (e.g., some transcription factors). This is unlikely to be a problem with pimonidazole immunohistochemistry, given the large amount of antigen. 6. Humidified incubation chamber: make sure that the slides stay horizontal. Humidification is obtained by placing blotting paper on the bottom of the chamber. Moisten the paper with water. To avoid bacterial contamination, clean the box every week with detergent and 70% ethanol. To avoid drying during incubation overnight, seal the incubation chamber with tape. 7. Pimonidazole should be injected between 2 h and 30 min before sacrifice at a dose between 6 and 10 mg/100 g. The timing and dosage should be consistent throughout the experiment. 8. Both the height of the bottles containing the perfusate solutions and the diameter of the perfusion cannula influence the perfusion flow. Optimal results are obtained when 300–400 mL of paraformaldehyde is consumed during the 5 min of perfusion fixation (per 200- to 300-g rat). 9. Anesthesia: the animal should be deeply anesthetized, since pain-induced stress may cause vasoconstriction and impair renal perfusion. Therefore, no reaction should be seen when pricking the tail. If necessary, add additional pentobarbital intraperitoneally. To avoid perforating the dorsal aortic wall, lift the tip of the cannula a little. Usually, the tip of the cannula is still visible through the aortic wall. 10. Ligation of the suprarenal aorta is an alternative method, leading to almost selective renal perfusion and consuming much lower perfusate volumes. However, care should be taken not to put traction on the renal pedicles, since this may cause vasoconstriction. Moreover, suprarenal aortic ligation should be performed immediately after caval vein incision and the start of perfusion. 11. Alternatively, you may fasten the perfusion line to the cork platform. However, this technique is less reliable when muscular fibrillations occur. 12. A large venous outflow is mandatory for high perfusate flow. You may have to enlarge the caval vein incision early during perfusion. If the venous outflow is too narrow, kidney swelling and interstitial edema will occur. 13. Artifacts may occur if too much pressure is put on the kidneys while harvesting. Note that the papilla is not completely in the transversal axis, but slightly oriented caudally. In order to achieve “long papillas,” section the kidney slightly oblique to the transverse axis. This perfusion technique gives good results not only for kidneys, but also for the liver, gut, and spleen. You may harvest these organs as well.

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14. Two to 4-µm sections may also be used, but section thickness should be consistent throughout the experiments. Uniform section thickness is assured by using parallel sections. If you are unable to perform at least 10 parallel sections (which may be the case with diseased kidneys), increase the section thickness stepwise by 1 µm. The quality of microtome sections can be considerably increased by the following “tricks”: trim the paraffin block prior to sectioning. For this purpose, remove the paraffin by oblique cuts starting about 2- to 4-mm from the edge of the specimen. Precut the block until the whole specimen is on the section (you may use an old blade for this, keeping the portion of the blade in mind). Keep the precut blocks at -20° for at least 4 h. For cutting, use a new blade but at the same portion used in precutting. 15. Renew the xylene baths after every 40 to 50 sections. 16. Digestion time may need adjustment, depending on the thickness of sections and fixation. Some protocols use microwave heating in citrate buffer (pH 6.0) instead of pronase digestion. According to the experience in our laboratory, heating may cause intracellular redistribution of signals from the cytoplasm into the nucleus. 17. Since the primary anti-pimonidazole antibody used in this protocol is raised in the mouse, this protocol is not recommended for the use in mouse kidneys. Using a fluorescein-tagged antipimonidazole antibody and anti-fluorescein secondary antibody (32) allows detection of pimonidazole adducts in both rats and mice. However, this alternative protocol needs verification in larger experimental series. Preliminary data from our laboratory indicate that using fluorescein-tagged anti-pimonidazole antibodies enhances the sensitivity but may reduce the specificity of the method. 18. The chromogen 3,3'-diaminobenzidine tetrahydrochloride (DAB) is toxic. Use gloves and store DAB waste in appropriate containers. DAB may be aliquoted and stored at −20°C for several years. Acknowledgment We thank Professor Sebastian Bachmann for advice in renal perfusion fixation and for reviewing the manuscript.

References 1. Franko, A. J., Chapman, J.D. (1982) Binding of 14C-misonidazole to hypoxic cells in V79 spheroids. Br. J. Cancer 45, 694–699. 2. Raleigh, J.A., Franko, A.J., Koch, C.J., Born, J.L. (1985) Binding of misonidazole to hypoxic cells in monolayer and spheroid culture. Br. J. Cancer 51, 229–235. 3. Arteel, G.E., Thurman, R.G., Yates, J.M., Raleigh, J.A. (1995) Evidence that hypoxia markers detect oxygen gradients in liver: pimonidazole and retrograde perfusion of rat liver. Br. J. Cancer 72, 889–895. 4. Kennedy, A.S., Raleigh, J.A., Perez, G.M., Calkins, D.P., Thrall, D.E., Novotny, D.B., Varia, M.A. (1997) Proliferation and hypoxia in human squamous cell carcinoma of the cervix: first report of combined immunohistochemical assays. Int. J. Radiat. Oncol. Biol. Phy. 37, 897–905. 5. Jankovic, B., Aquino-Parsons, C., Raleigh, J.A., Stanbridge, E.J., Durand, R.E., Banath, J.P., MacPhail, S.H., Olive, P.L. (2006) Comparison between pimonidazole binding, oxygen electrode measurements, and expression of endogenous hypoxia markers in cancer of the uterine cervix. Cytometry Part B (Clinical Cytometry)70B,45–55. 6. Leichtweiss, H.P., Lübbers, D.W., Weiß, C., Baumgärtl, H., Reschke, W. (1969) The oxygen supply of the rat kidney: measurements of intrarenal pO2. Pflügers Arch. 309(4),328–349. 7. Baumgärtl, H., Leichtweiss, H.P., Lübbers, D.W., Weiß, C., Hurland, H. (1972) The oxygen supply of the dog kidney: Measurements of intrarenal pO2. Microvasc. Res. 4, 247–257.

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8. Günther, H., Aumüller, G., Kunke, S., Vaupel, P., Thews, G. (1974) The oxygen supply of the kidney. I. Distribution of O2 partial pressures in the rat kidney under normal conditions (author’s transl)]. Res. Exp. Med. (Berl). 163(3),251–264. 9. Schurek, H.J., Jost, U., Baumgärtl, H., Bertram, H., Heckmann, U. (1990) Evidence for a preglomerular oxygen diffusion shunt in rat renal cortex. Am. J. Physiol. 259(6 Pt 2),F910–915. 10. Lübbers, D.W., Baumgärtl, H. (1997) Heterogeneities and profiles of oxygen pressure in brain and kidney as examples of the pO2 distribution in the living tissue. Kidney Int. 51(2),372–380. 11. Liss, P., Nygren, A., Erikson, U., Ulfendahl, H.R. (1997) Intrarenal oxygen tension measured by a modified clark electrode at normal and low blood pressure and after injection of x-ray contrast media. Pflügers Arch. 34(6),705–711. 12. Welch, W.J., Baumgärtl, H., Lübbers, D., Wilcox, C.S. (2001) Nephron pO2 and renal oxygen usage in the hypertensive rat kidney. Kidney Int. 59, 230–237. 13. Kriz, W. (1981) Structural organization of the renal medulla: comparative and functional aspects. Am. J. Physiol. 241, R3–R16. 14. Brezis, M., Rosen, S. (1995) Hypoxia of the renal medulla—its implications for disease. N. Engl. J. Med. 332(10),647–655. 15. Rosenberger, C., Heyman, S.N., Rosen, S., Shina, A., Goldfarb, M., Griethe, W., Frei, U., Reinke, P., Bachmann, S., Eckardt, K.U. (2005) Up-regulation of HIF in experimental acute renal failure: evidence for a protective transcriptional response to hypoxia. Kidney Int. 67(2),531–542. 16. Rosenberger, C., Shina, A., Rosen, S., Goldfarb, M., Eckardt, K.U., Heyman, S.N. (2006) Hypoxia inducible factors and tubular cell survival in isolated perfused kidneys. Kidney Int. 70, 60–70. 17. Rosenberger, C., Griethe, W., Gruber, G., Wiesener, M.S., Frei, U., Bachmann, S., Eckardt, K.U. (2003) Cellular responses to hypoxia after renal segmental infarction. Kidney Int. 64(3),874–886. 18. Goldfarb, M., Rosenberger, C., Abassi, Z., Shina, A., Zilbersat, F., Eckardt, K.U., Rosen, S., Heyman, S.N. (2006) Acute-on-chronic renal failure in the rat: functional compensation and hypoxia tolerance. Am. J. Nephrol. 26, 22–33. 19. Zhong, Z., Arteel, G.E., Connor, H.D., Yin, M., Frankenberg, M.V., Stachlewitz R.F., Raleigh, J.A., Mason, R.P., Thurman, R.G. (1998) Cyclosporin A increases hypoxia and free radical production in rat kidneys: prevention by dietary glycine. Am. J. Physiol. Renal Physiol. 275, F595–F604. 20. Suga, S.I., Phillips, M.I., Ray, P.E., Raleigh, J.A., Vio, C.P., Kim, Y.G., Mazzali, M., Gordon, K.L., Hughes, J., Johnson, R.J. (2001) Hypokalemia induces renal injury and alterations in vasoactive mediators that favor salt sensitivity. Am. J. Physiol. Renal. Physiol. 281, F620–F629. 21. Yin, M., Zhong, Z., Connor, H.D., Bunzendahl, H., Finn, W.F., Rusyn, I., Li, X., Raleigh, J.A., Mason, R.P., Thurmann, R.G. (2003) Protective effect of glycine on renal injury induced by ischemia-reperfusion in vivo. Am. J. Physiol. Renal Physiol. 282, F417–F423. 22. Basile, D.P., Donohoe, D.L., Roethke, K., Mattson, D.L. (2003) Chronic renal hypoxia after acute ischemic injury: effects of L-arginine on hypoxia and secondary damage. Am. J. Physiol. Renal Physiol. 284, F338–F348. 23. Matsumoto, M., Tanaka, T., Yamamoto, T., Noiri, E., Miyata, T., Inagi, R., Fujita, T., Nangaku, M. (2004) Hypoperfusion of peritubular capillaries induces chronic hypoxia before progression of tubulointerstitial injury in a progressive model of rat glomerulonephritis. J. Am. Soc. Nephrol. 15, 1574–1581. 24. Manotham, K., Tanaka, T., Matsumoto, M., Ohse, T., Miyata, T., Inagi, R., Kurokawa, K., Fujita, T., Nangaku, M. (2004) Evidence of tubular hypoxia in the early phase in the remnant kidney model. J. Am. Soc. Nephrol. 15, 1277–1288. 25. Tanaka, T., Miyata, T., Inagi, R., Fujita, T., Nangaku, M. (2004) Hypoxia in renal disease with proteinuria and/or glomerular hypertension. Am. J. Pathol. 165(6),165:1979–1992. 26. Tanaka, T., Kojima, I., Ohse, T., Inagi, R., Miyata, T., Ingelfinger, J.R., Fujita, T., Nangaku, M. (2005) Hypoxia-inducible factor modulates tubular cell survival in cisplatin nephrotoxicity. Am. J. Physiol. Renal Physiol. 289, 1123–1133. 27. Wang, P.X., Sanders, P.W. (2005) Mechanism of hypertensive nephropathy in the Dahl/Rapp rat: a primary disorder of vascular smooth muscle. Am. J. Physiol. Renal Physiol. 288, F236–F242.

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28. Rosenberger, C., Mandriota, S., Jürgensen, J., Wiesener, M.S., Hoerstrup, J.H., Frei, U., Ratcliffe, P.J., Maxwell, P.H., Bachmann, S., Eckardt, K.U. (2002) Expression of hypoxia inducible factor-1alpha and -2alpha in hypoxic and ischemic rat kidneys. J. Am. Soc. Nephrol. 13(7),1721–32. 29. Rosenberger, C., Rosen, S., Heyman, S. (2005) Current understanding of HIF in renal disease. Kidney Blood Pres. Res. 28, 32–350. 30. Prasad, P.V., Epstein, F.H. (1999) Changes in renal medullary pO2 during water diuresis as evaluated by blood oxygenation level-dependent magnetic resonance imaging: effects of aging and cyclooxygenase inhibition. Kidney Int. 55, (1)294–8. 31. Prasad, P.V., Priatna, A., Spokes, K., Epstein, F.H. (2001) Changes in intrarenal oxygenation as evaluated by BOLD MRI in a rat kidney model for radiocontrast nephropathy. J. Magn. Reson. Imaging. 13, 744–7. 32. Samoszuk, M.K., Walter, J., Mechetner, E. (2004) Improved immunohistochemical method for detecting hypoxia gradients in mouse tissues and tumors. J. Histochem. Cytochem. 52, (6)837–839.

Chapter 13

Identification of Apoptosis in Kidney Tissue Sections Glenda Gobe

Abstract The need for identification of specific modes of cell death, like apoptosis and necrosis, is driven by their detrimental or beneficial effect in different forms of disease, and the need in many instances of disease to modulate their levels. Apoptosis, an organized, gene-driven, and often energy-dependent mode of cell death, may be identified in tissue sections by its distinct morphological features, DNA degradation that is executed by endonucleases, and by presence of certain proteins, like the activated caspases. In the kidney, apoptosis is central to the development of a normal healthy kidney and it has been noted in glomeruli, the tubulo-interstitium, and renal vasculature in renal diseases or syndromes as diverse as acute kidney injury and chronic kidney disease of various causes, renal complications of diabetes and hypertension, sepsis, immune disorders and inflammation, nephrotoxicity, and in the development, progression, and treatment of renal cancers. Many research articles analyze apoptosis in tissue sections using the TUNEL assay that detects DNA strand breaks in situ in tissue sections. This method has been criticized because of false-positive or false-negative findings, and in situ analysis of activated caspase-3, thought to be the “executioner” caspase in the apoptotic pathway, may be a good alternative for quantifying apoptosis by light microscopy. The morphology of apoptosis, however, remains a standard that should not be ignored. This chapter reviews current methods of identifying apoptosis in tissue sections, with an emphasis on identification and quantification in the kidney using molecular methods. Keywords Apoptosis, Necrosis, TUNEL, Caspase, Cell death

1

Introduction

The morphologic hallmarks of apoptosis, that include nuclear chromatin margination and fragmentation, cellular condensation and blebbing, and the production of membrane-bound apoptotic bodies with preserved organelles and occasional nuclear fragments, are still considered by many as the gold standard for identification From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_13, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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of apoptosis in tissue sections (1–3). The term “apoptosis”, derived from the Latin for “apo” = away and “ptosis” = falling to signify loss of cells via apoptosis as similar to a tree losing its leaves in autumn (4), gives a visual picture of single cell deletion by this process in tissues. In direct contrast with apoptosis, the other key mode of cell death is necrosis. Necrotic cells lose energy and fail to maintain homeostasis; and in necrotic cells there is general cellular swelling and degradation, release of lysosomal enzymes, and an initiation of an acute inflammatory response, usually absent in apoptosis. Contiguous sheets of cells die, in comparison with the single cell deletion often seen in tissue sections with apoptosis. There has now been an attempt in the literature, in renal as well as other tissue and organ disease research, to present other modes of cell death, for example, oncosis or autophagic cell death (5, 6 ). Whilst it is interesting, in specific situations, to consider these less well-known types of cell death, there are in fact many structural and molecular similarities between them and the better-recognized cell death types of apoptosis and necrosis.

1.1 Light and Electron Microscopy for Morphological Characteristics of Apoptosis The most common and by far the cheapest way of identifying apoptosis is by its distinct morphology in hematoxylin and eosin (H&E)-stained or periodic acid– Schiff (PAS)-stained histological tissue sections. Although this chapter concentrates on molecular methods for identification of apoptosis, in particular in nephrology research, identification of apoptosis by light and electron microscopy must be discussed. In H&E-stained sections, apoptotic cells and bodies can be seen as dense (dark blue) nuclear fragments and a condensed, eosinophilic (affinity for eosin, pink or orange) cytoplasm (Fig. 13.1a and Fig. 13.2a). The transmission electron microscopy (TEM) view of apoptosis is demonstrated in Fig. 13.1b. Although apoptotic cells often die singly, several apoptotic cells may be found adjacent to each other, indicating the process may sometimes be synchronous. There is generally little acute inflammatory infiltrate, such as a neutrophil infiltrate, around these cells. In epithelial or endothelial-lined structures, apoptotic cells often bleb into the lumens rather than use energy in phagocytic deletion by neighbouring cells or activated macrophages (see Note 1). In contrast, necrosis often occurs in contiguous sheets of cells and does initiate an inflammatory infiltrate (Fig. 13.2b). The exceptions to the simple morphological descriptors of apoptosis and necrosis (3) and the need to develop some histological expertise with normal versus pathological characteristics of any tissue under study, can make assessment of apoptosis using H&E-stained sections difficult. Figure 13.3 summarizes some of the causes, characteristic changes in a timeline, and pathways of apoptosis and necrosis. The specific morphological characteristics of apoptosis were first described from TEM analysis (4 ). Thus, the condensed and often crescentic look of the nuclear chromatin, the fact that the apoptotic bodies maintained membrane integrity,

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Fig. 13.1 a Light microscopy using an H&E-stained paraffin-embedded section. Apoptotic cells are indicated by arrows in the distal tubular epithelium of rat kidney 2 days after ischaemia– reperfusion injury. Note the compacted nuclear chromatin and the “halo” around the shrunken apoptotic cells. In (b), electron microscopy of the same tissue as (a) demonstrates how apoptotic bodies that were phagocytosed by a macrophage (M) in the tubular epithelium are starting to undergo lysosomal degradation

Fig. 13.2 In this figure, apoptosis (arrows) in the renal tubular epithelium and the interstitium (a) 4 days after cisplatin injury contrasts markedly with tubular necrosis (b; examples of necrotic tubular profiles arrows) and inflammation (pockets of neutrophils indicated with asterisks) in human acute transplant rejection. The arrowhead in (b) indicates “anoikis” and apoptosis in what is likely a tubular epithelial cell that has lost adhesion and sloughed into the lumen of a tubule that is showing less extensive injury than the others indicated by arrows

and the phagocytosis and degradation of the apoptotic bodies in lysosomes of adjacent tissue cells or macrophages (Fig. 13.1b) were all originally described as ultrastructural characteristics. Electron microscopy is still a specific and sensitive way of analyzing for apoptosis, but the need for ultramicrotomes, the transmission

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Fig. 13.3 Examples are given of some causes of apoptosis and necrosis in kidney disease, and a summary of morphological characteristics and markers that may be used, as seen in paraffinembedded sections in a progressive timeline from initiation to cell death

electron microscopes themselves, and expertise for their use possibly hinders their more common application. An added bonus of using TEM is the application of biochemical and molecular markers of apoptosis for very specific ultrastructural localization of apoptosis and its associated proteins in tissues.

1.2

Biochemical and Molecular Characteristics of Apoptosis

Histology and TEM may be supplemented by biochemical or molecular characteristics of apoptosis. One of the widely accepted biochemical characteristics of apoptosis is internucleosomal DNA fragmentation by endonucleases. This feature was originally used for apoptosis identification because of its creation of a ladder-like appearance in DNA separated using agarose gel electrophoresis (7, 8), but by the early 1990s there was a swing to the in situ end labelling of the fragmented DNA, first using a Klenow DNA polymerase method, but then using the enzymatic method of terminal deoxyribonucleotidyl transferase (TdT)-mediated deoxyuridine triphosphate (dUTP) nick end labelling (9, 10). This method is now popular and is best known as TUNEL. Using TUNEL in tissue sections, dUTP attaches enzymatically to the 3′ end of the DNA fragments. If dUTP is labelled with a compound (like biotin) that reacts

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with a chromogenic substrate using common immunohistochemical (IHC) techniques, the enzymatic tag can be visualized under the microscope (Fig. 13.4a). The method is now used widely in studies of apoptosis in kidney disease (11). A limited population of intensely TUNEL-positive cells, that lack apoptotic morphology, may be seen. The opposite may also occur—apoptotic morphology without TUNEL. The labelled cells without apoptotic morphology may indicate a pre-apoptotic state and can be counted as apoptosis. There are, however, several caveats to analyzing the presence and contribution of apoptosis using TUNEL. While DNA fragmentation has proved to be a useful marker of apoptosis, some studies show that DNA fragmentation is not always required for apoptosis (12). In addition, the molecular or biochemical methods of quantifying apoptosis have to be made with the knowledge that, in some instances, they may not be specific for apoptosis. For example, in the early stages of necrosis, fragmentation of the DNA may label positively. During straightening of the DNA during mitosis, DNA is nicked to allow the process to occur and so again a positive enzymatic TUNEL label may occur. Both of these cases should be negated by identification of typical necrotic or mitotic nuclei using their specific morphology. Finally, there have been several reports, in renal research, of abnormally high levels of TUNEL identified early (0–2 days) after ischaemia–reperfusion injury (Fig. 13.4b), in comparison with the moderate increase in apoptosis identified at the same time using morphological means (13). Free radical damage to the DNA, as occurs after ischaemia–reperfusion, may manifest as single- and double-strand breaks, which may or may not be repaired. The cells in which DNA repair is not completed should continue to undergo

Fig. 13.4 A. TUNEL (ApopTag™) labelling of nuclei (examples indicated by arrows) in the renal tubular epithelium and interstitium 14 days after ischaemia–reperfusion injury in the rat. b In contrast with (a), where single cell labelling is obvious, here TUNEL is shown to label non-specifically in almost all nuclei 24 h after ischaemia–reperfusion. There is no morphological evidence, at this time or at later times when such labelling is not seen, that all cells die by apoptosis

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apoptosis, the presence of which is verifiable by morphological means. Unfortunately, the repairable DNA strand breaks may also label. If these are counted as “apoptotic”, estimates of apoptosis using enzymatic labelling will be incorrectly high. Thus, some caution should be applied in interpretation of molecular and biochemical labelling for apoptosis. Statistical comparison of biochemical labelling with morphology should be made routinely. In more recent times, in situ localization of activated caspases has found some favour for histological labelling of cells in, or fast approaching, apoptosis (Fig. 13.5). The caspases are a family of cysteine proteases that usually cleave their substrates after aspartate residues. There are multiple mammalian caspases that exist in cells as inactive precursor proteins (14). Once caspases are activated, they cleave a large number of proteins within the cell. This then allows processing of the morphological changes of apoptosis. For example, caspase-mediated cleavage of a putative endonuclease, caspase-activated DNase (CAD), from its inhibitor (ICAD) leads to activation of CAD and fragmentation of the DNA. Activation of caspase precursors is achieved by adaptor proteins that bind to them via shared motifs. For example, caspase-9 is activated after association of the caspase recruitment domain (CARD) in its pro-domain with the CARD in another adaptor protein, Apaf1, in the apoptosome. CAD or activated caspase-9 may be localized in tissue sections. In situ localization of activated (cleaved) caspase-3, thought to

Fig. 13.5 Cleaved (activated) caspase-3 labelling in renal tubular epithelium after oxidative stress. Note that several of the labelled cells show the morphology of apoptosis (the asterisk demonstrates one example)

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be the “executioner” caspase acting prior to the structural disaggregation of the cell during apoptosis, has found particular favour for histological labelling of apoptosis. In the kidney, activation of caspase-3 correlated well with TUNEL in the damaged tubular epithelium during ischaemic acute renal failure (15). A recent caveat, however, is that caspase-independent cell death can occur, and caspase activation can occur without any indication of apoptosis when morphological characteristics are sought (6, 16, 17). Several other molecular markers may be used to assist in situ detection of apoptosis in tissue sections, but many are tissue specific. For example, apoptotic changes in plasma membrane composition may be indicated by annexin V or tissue-type transglutaminase; other molecular markers include the deathinducing signaling complex (DISC), inhibitor of apoptosis protein (IAP), ICAD and CAD, poly(ADP-ribose) polymerase (PARP) (18), death receptors like Fas and tumour necrosis factor receptor (TNFR), and the balance of localized pro-apoptotic Bcl-2 proteins (like Bax, Bak, Bid, and Bad) against anti-apoptotic ones (like Bcl-2 and Bcl-XL) (19). These markers may be analysed in tissue sections using IHC, but would, almost invariably, need a second biochemical or molecular marker such as TUNEL or activated caspase-3, along with morphology, to be sure of identification of the complete apoptotic process in tissue sections.

1.3

Is it Apoptosis and What Is the Significance of its Presence?

Apoptosis is a rapid process, taking approximately 4 to 6 hours from initiation to the structural disassembly of the apoptotic cell (7 ). In paraffin sections, we are looking at a 4-µm thin fraction of a segment of the whole kidney at a moment in time. A single cell has a thickness in the vicinity of 8–10 µm. Thus, even low numbers of apoptotic cells found in the histological section of the kidney can account for significant loss of tissue. To be sure it is apoptosis that is being identified, a general “rule of three hits” may be applied for identification and quantification of apoptosis: 1) is apoptotic cell death expected in a certain situation?; 2) can it be identified morphologically in comparison, say, with reversible cell injury or necrosis?; and 3) is there at least one biochemical or molecular test for apoptosis that is positive? In most instances, if these run true, the conclusions made about any analyses of apoptosis should be sound. Manuscript reviewers seem reassured by presentation of a linear regression analysis that compares and finds a significantly positive correlation between apoptosis, identified using morphology, with any biochemical or molecular marker, particularly where most of the data are to be presented as the biochemical or molecular marker. The methods for analyses of apoptosis in tissue sections are now described, with particular references to morphology, TUNEL, and activated caspase-3 labelling, and special notes are made for relevance in kidney sections. Where applicable, some notes are also given for analysis of apoptosis in cell culture experiments.

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Materials

All general chemicals and most histological stains are found from suppliers such as Sigma–Aldrich Pty. Ltd. (Castle Hill, Australia) or Promega Corporation (Madison, WI, USA).

2.1

Tissue Fixation

Tissue may be fixed in either neutral buffered formalin or 4% paraformaldehyde in phosphate buffer. 1. 10% Neutral buffered formalin: mix 4.05 g monobasic sodium phosphate (NaH2PO4.2H2O), 5.85 g dibasic sodium phosphate (Na2HPO4) and 100 mL of 40% formaldehyde solution in 900 mL distilled water (dH2O) until dissolved. 2. 4% Paraformaldehyde in 0.1 M phosphate buffer: (a) Solution A (8% paraformaldehyde): in a fume hood dissolve 80 g paraformaldehyde in 1 L of dH2O at 50°C by adding 2.0 M NaOH until clear, allow to cool and adjust pH to neutral with HCl. (b) Solution B (0.2 M dibasic sodium phosphate): dissolve 2.84 g dibasic sodium phosphate (Na2HPO4) in 100 mL dH2O. (c) Solution C (0.2 M monobasic sodium phosphate): dissolve 3.12 g monobasic sodium phosphate (NaH2PO4.2H2O) in 100 mL dH2O. To use, mix 36 mL of solution B with 14 mL of solution C and 50 mL of solution A.

2.2

Staining Materials

2.2.1

H&E Histostain

1. Menzel Superfrost Plus™ glass histology slides (Menzel-Gläser, Braunschweig, Germany). 2. Phosphate-buffered saline (PBS), pH 7.4. 3. Xylene. 4. Graded ethanols at absolute, 95%, and 70% (v/v). 5. Mayer’s hematoxylin. 6. Blueing solution: 1 g sodium bicarbonate in 1 L distilled water. Tap water or other blueing solutions like Scott’s Tap Water may also be used. 7. Eosin. 8. Xylene-based mounting medium (see Note 2).

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Periodic Acid–Schiff (PAS) Histostain

1. 1% Periodic acid in dH2O (w/v). 2. Schiff reagent: boil 200 mL of dH2O, remove flask from heat and add 1 g of basic fuchsin. Allow to cool to 50°C. Add 2 g of potassium metabisulphate (K2S2O5) whilst mixing. Cool to room temperature and add 2 mL of concentrated HCl, mix, and let the solution stand in the dark overnight. Add a large amount of activated charcoal to the solution, shake it well, and filter. The solution should be clear/pale yellow. Store in a dark glass bottle at 4°C. 3. Mayer’s hematoxylin. 4. Xylene and graded ethanols. 5. Xylene-based mounting medium (see Note 2).

2.2.3

Toluidine Blue Histostain for Resin Sections

1. Toluidine blue stain: dissolve 1 g toluidine blue stain and 1 g sodium tetraborate in 100 mL boiling water, stir for 30 min, cool, and filter. 2. Ethanol. 3. Permanent mounting medium (see Note 2).

2.3

TUNEL (See Note 3)

1. ApopTag™ Apoptosis In Situ Detection Kit (Cat No S7100; Serologicals Corp., Chemicon International, Temecula, CA, USA). 2. DNAase. 3. Menzel Superfrost Plus™ adhesive slides Menzel-Gläser (Menzel-Gläser). 4. Graded ethanols. 5. PBS, pH 7.4. 6. Coplin jar. 7. Proteinase K (20 µg/mL in 0.05 M Tris-HCl, 0.01 M CaCl2 pH 8.0; Cat No. V3021; Promega). 8. 3% v/v Stock hydrogen peroxide (H2O2) in PBS. 9. TdT enzyme (made by mixing 9.5 µL of kit reaction buffer with 3 µL TdT). 10. Nescofilm™ (Bando Chemical, Kobe, Japan) or Parafilm™ (Sigma–Aldrich). 11. Anti-digoxygenin conjugate. 12. 3,3′-Diaminobenzidine hydrochloride (DAB) with H2O2 as substrate: 4 mL of DAB (DAKO, Botany, Australia) mixed with 3 µL of 30% H2O2 stock solution. Use within 5 min of adding H2O2 as the solution is unstable. Alternatives to DAB are Vector Laboratories (Burlingame, CA, USA) Nova Red™, VIP (purple), or SG (blue) chromogens. 13. Mayer’s hematoxylin.

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14. Graded ethanols. 15. Xylene. 16. Xylene-based mounting medium (see Note 2).

2.4 Activated or Cleaved Caspase-3 (or Other Protein) IHC from Paraffin Sections 1. 2. 3. 4. 5. 6. 7. 8. 9.

10. 11.

12. 13.

2.5

Menzel Superfrost Plus™ adhesive slides. Xylene, graded ethanols. PBS, pH 7.4. 0.001 M EDTA/0.01 M Tris-HCl, ph 8.0. Heat-induced epitope retrieval (HIER) solutions (see below). Decloaking chamber (Biocare Medical, Concord, CA, USA). 1% H2O2 v/v, 0.1% w/v sodium azide in PBS. Normal goat serum (10% in PBS) (Vector Laboratories). Primary anti-caspase-3 antibody (rabbit anti-cleaved caspase-3 ASP175; Cat No. 9661S; Cell Signaling, Danvers, MA, USA) at a suggested dilution of 1:100 in PBS. Dako Envision-plus™ anti-rabbit secondary antibody. DAB with H2O2 as substrate: 4 mL of DAB (DAKO) mixed with 3 µL of 30% H2O2 stock solution. Use within 5 min of adding H2O2 because the solution is unstable. Mayer’s hematoxylin. Xylene-based mounting medium (see Note 2).

Double Protein Staining Using Immunofluorescence (IF)

1. 2. 3. 4.

Tris-buffered saline/0.1% Tween 20 (TBST) (see Note 4). 3% Bovine serum albumin (BSA) in TBST. Primary antibodies derived from different species of origin. Fluorescent secondary antibodies to the species of origin of the primaries (Molecular Probes, Eugene, OR, USA). 5. Aqueous mounting medium (see Note 2) with anti-fade for fluorescence microscopy.

2.6

Heat-Induced Epitope Retrieval Solutions

A variety of HIER solutions may be employed. What follows is a selection of commonly used examples.

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1. Citrate (0.01 M) pH 6.0: dissolve 3.84 g of citric acid (anhydrous) in 1,800 mL of distilled water. Adjust to pH 6.0 using concentrated NaOH and make up to 2,000 mL with distilled water. 2. EDTA (1 mM) pH 8.0: dissolve 0.37 g of EDTA in 1,000 mL of distilled water. Adjust pH to 8.0 using 1 M NaOH. 3. Tris/EDTA (10/1 mM) pH 9: add 1.21 g of Tris and 0.37 g of EDTA in 1,000 mL of distilled water. Adjust to pH 9 if necessary. 4. 10× TRIS-HCl/EDTA (20 mM/0.65 mM)–Tween 20 (0.0005%) pH 9.0: dissolve 14.4 g Tris and 1.44 g EDTA in 550 mL of dH2O. Adjust to pH 9 using 1 M HCl, and add 0.3 mL Tween 20. Make up to 600 mL with distilled water. This is a 10× concentrate, which should be diluted with distilled water as required (e.g. 25 mL diluted with 225 mL distilled water). 5. Sodium Citrate Buffer (10 mM)–Tween 20 (0.05%) pH 6.0: dissolve 2.94 g tri-sodium citrate in 1,000 mL distilled water. Adjust to pH 6.0. Add 0.5 mL of Tween 20 and mix well. 6. Tris-NaOH Buffer (10 mM)–Tween 20 (0.05%) pH 10: dissolve 1.21 g Tris Base in 1,000 mL distilled water. Adjust to pH 10 using 1 M NaOH. Add 0.5 mL Tween 20 and mix well.

3 3.1

Methods Tissue Fixation for Histology (See Note 5)

Ten percent neutral buffered formalin or 4% paraformaldehyde in 0.1 M phosphate buffer pH 7.4 is routinely used for histological fixation of tissue prior to tissue staining (see Note 5). TUNEL and IHC for activated caspase-3 work well after these fixatives. A broad range of analyses can be performed if the removed kidney is cut into separate pieces for 1) immersion-fixation for microscopy and IHC, 2) embedding in OCT and storage at -80°C for preparation of frozen tissue sections needed for some IHC methods, and 3) freezing of tissue, usually in liquid nitrogen followed by storage at -80°C, suitable for extracting messenger RNA (mRNA) or protein for molecular analysis (polymerase chain reaction [PCR], Western blotting). If fine cellular structure or cellular ultrastructure is required, the latter sometimes necessary to verify morphology of apoptosis using TEM, a general-purpose ultrastructural fixative should be used such as 4% paraformaldehyde, 4% glutaraldehyde, and 0.1% calcium chloride in 0.1 M sodium cacodylate buffer, pH 7.4. This can be used for immersion fixation of small pieces of kidney or the tissue may be perfused. Glutaraldehyde makes the use of TEM fixative problematic for TUNEL and IHC, and in the case of a need for TEM and TUNEL or IHC, the immersion fixative method should be used for the TEM tissue. If perfusion is selected, the fixative may be applied via the aorta retrograde to blood flow, nicking the vena cava to allow the perfusate to flow easily. An initial application of 0.1% heparin in normal

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isotonic saline (50 to 100 mL for a whole rat) will remove blood from fixed tissue. Flow rates should be at approximately normal blood pressure. Tissues are then perfused with fixative (50 to 100 mL for a whole rat), again at a suitable rate and pressure. After perfusion, tissues are dissected out and cut into appropriately sized blocks for histology or electron microscopy. The tissue should be left in fixative for a further 2 to 4 h before washing at least three times in 0.1 M sodium cacodylate buffer, pH 7.4, or similar, over a period of several hours to remove all traces of free aldehydes.

3.2

H&E Protocol for Paraffin and Frozen Sections and Cultured Cells

3.2.1

Paraffin Sections

1. Cut 3- to 6-µm sections to standard or Menzel Superfrost Plus™ adhesive slides and air-dry overnight at 37°C. 2. Dewax and rehydrate through descending graded alcohols to PBS, pH 7.4, using a standard protocol: three changes of xylene, 3 min each; two changes of 100% ethanol, 2 min each; 95% ethanol, 1 min; 70% ethanol, 1 min; and PBS, 2 min. 3. Tissue sections are then stained with Mayer’s hematoxylin for 4 min, washed in tap water (2 × 3 min) and immersed in “bluing solution” for 2–3 min, then returned to water. 4. Tissue sections are then dehydrated in 70% ethanol (1 min), 95% ethanol (2 × 2 min), and absolute ethanol (1 min). 5. Eosin is applied for 2 min followed by washing in absolute ethanol (3 × 2 min), clearing in xylene (3 × 2 min), and mounting and coverslipping with DEPEX™ or Permount™ mounting medium (see Note 2). H&E stains nuclei blue and cytoplasm pink–orange. 3.2.2

Frozen Sections

Sections are cut onto slides, rehydrated in PBS, and then undergo the same staining protocol as paraffin-embedded sections. 3.2.3

Cultured Cells

In experiments involving cell culture (also see Note 6), cells are grown on sterile glass cover slips, treated, then fixed and stained to provide an assessment of apoptosis in a similar manner to histological sections. The level of apoptosis evident in such in vitro studies may also be assessed using flow cytometric analysis of trypsinized single cell suspensions following cell surface staining for annexin V expression or staining of nuclear DNA with DNA binding dyes such as propidium iodide.

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187

Periodic Acid–Schiff Stain

PAS is an alternative stain to H&E. 1. Bring sections on plain slides to water. 2. Treat with 1% periodic acid for 10 min and wash in water. 3. Treat with Schiff reagent for 10 min. Do not over stain in Schiff reagent—it is irreversible. 4. Wash for several minutes in tap water to bring out the color of Schiff. 5. Stain nuclei with Mayer’s hematoxylin for 2 min. 6. Blue the sections as for H&E staining. Dehydrate in graded ethanols, clear in xylene, and mount sections in DEPEX™. Glycogen and other periodate reactive carbohydrates stain magenta and nuclei stain blue.

3.4

Light Microscopy for Apoptosis Assessment

To view stained sections under the microscope, use a low power objective (×10) to scan the kidney sections to gauge the extent and localization of pathologic changes. H&E stains nuclei blue and cytoplasm pink/orange. PAS stains the nuclei blue and the cytoplasm pink/magenta. The pale diffuse nuclear chromatin and pale pink/ magenta cytoplasm of healthy cells contrasts markedly with the condensed, dark blue nuclear chromatin of apoptotic cells and apoptotic bodies, and the dense eosinophilic (taking the eosin colour preferentially) or deep magenta staining of the cytoplasm in apoptosis. A halo can often be seen around apoptotic cells, because they tend to shrink and convolute in the apoptotic process (see Fig. 13.1a). In addition, apoptotic cells within solid tissues are often located within the phagolysosomes of macrophages (see Fig. 13.1b) or neighbouring healthy cell (20). The number of apoptotic cells can be counted in defined renal cell types and zones, for example, glomeruli, tubular epithelium, interstitial space, cortex, or medullary zones. Quantification of renal cell apoptosis is usually carried out using a ×40 objective lens in a defined area of tissue. Multiple fields are assessed and the level of apoptosis may be recorded as a percent of total cells or the number of apoptotic cells per square millimeter of tissue.

3.5

Resin Sections and Toluidine Blue Stain

Morphology of apoptosis is very clear when sections are cut at 0.5–1 µm, as can be done with resin sections. Included here is a stain for toluidine blue on resin. 1. Heat resin sections on slides on the hot plate for 5 min, apply three drops of stain and wait 30 to 60 s. Do not let the slide dry.

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2. Rinse under running water to remove all excess stain including the dried edge. 3. Differentiate for 1 to 2 min in three changes of absolute ethanol. Wipe the back of the slide and dry on the hot plate. Gently blowing on the slide as it dries will help retard the formation of “tide” lines. 4. Cover slip using DEPEX™ or similar. This stain shows nuclei as deep blue and cytoplasm as pale pink/blue.

3.6

Protocol for TUNEL (Using Apotag™ In Situ Apoptosis Detection) on Paraffin Sections

The ApopTag™ in situ detection kit is used by our laboratory (see Note 3). Always use a positive and negative control slide with each batch of slides. Positive slides may be prepared by treating sections with DNAase or using sections with known high levels of apoptosis. Negative controls are prepared by omitting the TdT enzyme from the TUNEL reaction. 1. Paraffin sections (3–4 µm) are affixed to Menzel Superfrost Plus™ adhesive slides and air-dried overnight at 37°C, dewaxed, and rehydrated through descending graded ethanols to PBS, pH 7.4, using a standard protocol. 2. 150 µL of freshly diluted proteinase K is added to the section for 10 min at room temperature. 3. Slides are washed twice in dH2O in a Coplin jar (2 × 2 min). 4. Endogenous peroxidase activity is blocked by incubating the sections in 3% hydrogen peroxide (H2O2) in PBS for 5 min followed by two washes in PBS (2 × 5 min). 5. Apply 17.5 µL of the kit equilibration buffer (neat) directly onto the section and incubate for 10 min (note, incubation time varies greatly, from 10 s to 60 min). 6. Pipette off and apply 12.5 µL working strength TdT enzyme (made up by mixing 9.5 µL of reaction buffer with 3 µL TdT) for 60 min at 37°C. Use a preheated humidified chamber placed into a 37°C waterbath, and cover the sections with Nescofilm™ or Parafilm™ squares to prevent dehydration of the section. 7. Tip off the TdT enzyme and apply working strength stop wash buffer (made by mixing 100 µL of stop wash buffer with 3.4 mL of dH2O), agitate for 15 s, and incubate for 10 min. 8. Wash sections (3 × 2 min in PBS) and incubate with 17.5 µL anti-digoxygenin conjugate (neat) for 30 min at room temperature, again covering sections with Nescofilm™ or Parafilm™ squares. 9. Wash (4 × 2 min in PBS) and develop color in DAB with H2O2 as substrate for 5 min. Alternatives are Vector Laboratories Nova Red™, VIP (purple), or SG (blue) chromogens. 10. Wash sections in running water for 5 min and counterstain with a light hematoxylin (no eosin) for 2 min. 11. Dehydrate, clear in xylene, and mount using DEPEX™ and glass cover slips.

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189

Protocol for Activated or Cleaved Caspase-3 (or Other) Immunohistochemistry

Always run a negative and positive control slide with each batch of slides. The positive sections should be from tissue with known positivity for activated caspase-3 (from previous tests). Negative controls are prepared by omitting the primary caspase-3 antibody from the procedures. 1. Sections (3–4 µm) are cut onto Menzel Superfrost Plus™ adhesive slides, airdried overnight at 37°C, and dewaxed and rehydrated through descending graded alcohols to PBS pH 7.4 using a standard protocol. 2. Sections are transferred to 0.001 M EDTA/0.01 M Tris-HCl, pH 8.0, and subjected to a predetermined time in HIER solution (see Sect. 3.9), using a decloaking chamber (Biocare Medical) and the HIER of choice at 125°C. There are many HIER solutions, but the one most commonly used is the citrate buffer solution. 3. After completion of the cooling cycle, the containers of slides are removed and the slides are allowed to cool for a further 20 min before transferring back to PBS. 4. Endogenous peroxidase activity is blocked by incubating the sections in 1% H2O2 and 0.1% sodium azide in PBS for 10 min. 5. Sections are washed (3 × 5 min in PBS), and normal goat serum (10% in PBS) is applied for 20 min. 6. Excess normal serum is decanted from the sections and the primary anti-caspase 3 antibody is applied overnight at 4°C at a suggested dilution of 1:100 in PBS. 7. Sections are washed (3 × 5 min in PBS) and DAKO Envision-plus™ secondary antibody (suggested dilution, 1:2,000 in PBS) is applied for 30 min. 8. Sections are washed (3 × 5 min in PBS) and colour is developed in DAB with H2O2 as substrate for 5 min. 9. Sections are washed in gently running tap water for 5–10 min to remove excess chromogen, lightly counterstained in Mayer’s hematoxylin, then dehydrated through ascending graded alcohols, cleared in xylene, and mounted using DEPEX™ or similar.

3.8

Double Staining Using Immunofluorescence

In some instances, double-staining of TUNEL with other proteins may be needed in paraffin-embedded sections. 1. Sections are dewaxed in xylene before rehydrating in graded ethanols to TBST buffer. 2. Non-specific binding sites are blocked with 3% BSA in TBST for 1 h at room temperature.

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3. Sections are then incubated with primary antibody (prepared in a species other than sheep or goat) against the selected protein in 3% BSA in TBST overnight at 4°C and washed (3 × 10 min in TBST). 4. Incubations for ApopTag™ labelling are carried out using the protocol detailed previously. 5. Sections are then incubated with a fluorescent secondary antibody mixture containing, say, an Alexa-Fluor™-labelled donkey anti-mouse IgG or another secondary appropriate to the antibody to your protein of interest, at a dilution of 1:200 in 1% BSA in TBST for 1 hr at room temperature. 6. Sections are washed (3 × 30 min in TBS) and then mounted with glass cover slips using either DAKO fluorescent anti-fade mounting medium (DAKO) or Vectashield™ mounting medium for fluorescence with 4′-6-diamadino-2phenylindole (DAPI; Vector Laboratories) and allowed to set overnight in the dark prior to light and fluorescence microscopy, and photography (Note 7 refers to endogenous and background fluorescence).

3.9

HIER Microwave Method

1. Deparaffinize sections and rehydrate through graded alcohols to dH2O using a standard protocol. 2. Place slides in staining jar containing the HIER solution of choice. In a microwave oven, heat the slides until the solution is boiling (3 min on high/900 W for 24 slides in 250 mL HIER solution). Reduce power to medium/150 W and heat (simmer) for 7 to 17 min. Effect and time may vary depending on the microwave oven, the volume of HIER solution, and the number of trays. 3. Allow slides to reach room temperature and place slides in wash buffer.

4

Notes

1. In the kidney, apoptotic cells and membrane-bound apoptotic bodies are either taken up by adjacent renal tissue cells that have become phagocytic under the influence of changed surface molecules, like phosphatidyl serine, on apoptotic cells, or by more professional phagocytes such as macrophages, or they are lost to the renal nephron lumen and washed away with the filtrate. Energy conservation is undoubtedly a factor in the pathophysiology of renal disease, and so the energy-consuming process of engulfment and breakdown of apoptotic cells by adjacent renal tubular epithelial cells and macrophages may happen less often than the sloughing of viable epithelial cells that later become apoptotic (called anoikis), or epithelial cells that have become apoptotic whilst still in the epithelium, into the tubular lumina. The quantification of apoptosis in renal tissue sections is therefore sometimes difficult and the numbers determined often an underestimation of the contribution of apoptosis to tissue loss. 2. Xylene-based mounting media are DEPEX™ (BDH, Poole, UK) or Permount™ (Fisher, Pittsburgh, PA, USA). A general aqueous mounting medium is Gel Mount™ (Sigma–Aldrich Cat No. G0918). For fluorescence microscopy, use DAKO fluorescent anti-fade mounting medium (DAKO) or Vectashield™ mounting medium for fluorescence with DAPI (Vector Laboratories).

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3. There are many ways of preparing for the TUNEL reaction. These can be found in the literature or using web-based searches. We use the ApopTag™ Apoptosis In Situ Detection Kit from Millipore Corporation; Cat No. S7100. This works well in kidney sections, both experimental and human. 4. We routinely use PBS in all IHC and IF, but some laboratories insist that Tris-buffered saline should be used. 5. All fixation should be carried out in a fume hood with gloves worn at all times because paraformaldehyde, sodium cacodylate, and glutaraldehyde are hazardous. Perfusion of animal tissues with the perfusate should be carried out under supervision of someone familiar with perfusion techniques and with appropriate ethical clearances. 6. There are several important considerations when assessing apoptosis in vitro. The homogeneous cell culture of a single cell type, that might be adherent or in suspension, is usually being studied in the absence of phagocytic cells. Apoptotic cells in adherent cultures can be assessed in a similar manner to tissue sections by growing the cells on glass or plastic cover slips, fixing, and staining the cells on the cover slips for histological or molecular characteristics. Note that apoptotic cells may detach from the growth plate or cover slips and assessment of apoptosis only in the remaining adherent culture may result in a gross underestimation of the extent of cell death that has occurred. Cells that are induced to undergo apoptosis in suspension cultures, and those that bleb into the supernatant from dying adherent cultures, tend to undergo a secondary degradative process termed “secondary necrosis” and it is sometimes difficult in this context to identify cells as being apoptotic in origin. Viable cells may also detach with injury and then undergo apoptosis as a secondary phenomenon. This process is sometimes referred to as “anoikis” (demonstrated in Fig. 13.2b). Thus, the supernatant from such cultures also needs to be collected and assessed. 7. Endogenous fluorescence may be a problem. Causes include fixation artifacts (free aldehydes in the tissue), lipofuscin pigment, elastin, and collagen. Tissues that have been aldehyde fixed with formaldehyde, paraformaldehyde, or glutaraldehyde may be autofluorescent (this is almost certain with glutaraldehyde) and this activity can be quenched by incubating the sections in fresh 0.1% sodium borohydride in PBS for 20–30 min followed by washing (3× 5 min in PBS).

References 1. Kerr, J.F., Gobe, G.C., Winterford, C.M., and Harmon B.V. (1995) Anatomical methods in cell death. Methods Cell Biol. 46, 1–27. 2. Andrade, L., Vieira, J.M., and Safirstein, R., (2000) How cells die counts. Am J Kidney Dis. 36, 662–668. 3. Gobe, G.C. and Harmon, B.V. Apoptosis: morphological criteria and other assays. Encyclopedia Life Sciences 2006 [cited 2007 June 14]; Available from: http://www.mrw.interscience. wiley.com/emrw/9780470015902/els/article/a0002569/current/abstract?hd=All,apoptosis. 4. Kerr, J.F., Wyllie, A.H. and Currie, A.R. (1972) Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer. 26, 239–257. 5. Takeda, M., Shirato, I., Kobayashi, M., and Endou, H. (1999) Hydrogen peroxide induces necrosis, apoptosis, oncosis and apoptotic oncosis of mouse terminal proximal straight tubule cells. Nephron. 81, 234–238. 6. Colell, A., Ricci, J.E., Tait, S., Milasta, S., Maurer, U., Bouchier-Hayes, L., Fitzgerald, P., Guio-Carrion, A., Waterhouse, N.J., Li, C.W., Mari, B., Barbry, P., Newmeyer, D.D., Beere, H.M., and Green, D.R. (2007) GAPDH and autophagy preserve survival after apoptotic cytochrome c release in the absence of caspase activation. Cell. 129, 983–997. 7. Wyllie, A.H. (1980) Glucocorticoid-induced thymocyte apoptosis is associated with endogenous endonuclease activation. Nature. 284, 555–556.

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8. Arends, M.J., Morris, R.G., and Wyllie, A.H. (1990) Apoptosis. The role of the endonuclease. Am J Pathol. 136, 593–608. 9. Gavrieli, Y., Sherman, Y., and Ben-Sasson, S.A. (1992) Identification of programmed cell death in situ via specific labeling of nuclear DNA fragmentation. J Cell Biol. 119, 493–501. 10. Ansari, B., Coates, P.J., Greenstein, B.D., and Hall, P.A. (1993) In situ end-labelling detects DNA strand breaks in apoptosis and other physiological and pathological states. J Pathol. 170, 1–8. 11. Ueda, N. and Shah, S.V. (2000) Role of endonucleases in renal tubular epithelial cell injury. Exp Nephrol. 8, 8–13. 12. Danial, N.N. and Korsmeyer, S.J. (2004) Cell death: critical control points. Cell. 116, 205–219. 13. Gobe, G.C. and Endre, Z.H. (2003) Cell death in toxic nephropathies. Semin Nephrol. 23, 416–424. 14. Oliver, L. and Vallette, F.M. (2005) The role of caspases in cell death and differentiation. Drug Resistance Updates. 8, 163–170. 15. Villanueva, S., Cespedes, C., and Vio, C.P. (2006) Ischemic acute renal failure induces the expression of a wide range of nephrogenic proteins. Am J Physiol Regul Integr Comp Physiol. 290, R861–870. 16. Cummings, B.S., Kinsey, G.R., Bolchoz, L.J., and Schnellmann, R.G. (2004) Identification of caspase-independent apoptosis in epithelial and cancer cells. J Pharmacol Exp Ther. 310, 126–134. 17. Nakagawa, Y., Iinuma, M., Naoe, T., Nozawa, Y., and Akao, Y. (2007) Characterized mechanism of alpha-mangostin-induced cell death: Caspase-independent apoptosis with release of endonuclease-G from mitochondria and increased miR-143 expression in human colorectal cancer DLD-1 cells. Bioorg Med Chem. 15, 5620–5628. 18. Chang, H.Y. and Yang, X. (2000) Proteases for cell suicide: functions and regulation of caspases. Microbiol Mol Biol Rev. 64, 821–846. 19. Gobe, G., Zhang, X.J., Willgoss, D., Hogg, N., and Endre, Z. (2000) Relationship between Bcl-2 genes, growth factors and apoptosis in acute ischemic renal injury in the rat. J Am Soc Nephrol. 11, 454–467. 20. Strater, J., Gunthert, A.R., Bruderlein, S., and Moller, P. (1995) Microwave irradiation of paraffin-embedded tissue sensitizes the TUNEL method for in situ detection of apoptotic cells. Histochem Cell Biol. 103, 157–160.

Chapter 14

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

Abstract The pathology of progressive renal disease is characterized by glomerular and interstitial inflammation, glomerulosclerosis, and tubulointerstitial fibrosis. This is a consequence of excessive matrix synthesis, reduced matrix degradation, and contraction (reorganization) of extracellular matrix. Fibroblasts, and to a lesser degree, other mesenchymal cells, are known to contribute to renal scar formation through local proliferation, synthesis, and reorganization of matrix proteins. Although much work has focused on the balance between collagen synthesis and degradation, the mechanisms of parenchymal collapse and contraction are becoming increasingly important. Like their counterparts in the skin, the contractile properties of renal fibroblasts are now well recognized. This chapter details an in vitro method for studying the contraction of collagens by homogeneous populations of cultured cells. The method can be altered so that reagents influencing this process may also be studied. Keywords Fibroblast, Collagen, Contraction,

1

Introduction

Fibrosis and sclerosis, so-called pathological scarring, are the ultimate consequence of chronic injury in the kidney. The definition is a histological one; both processes being defined as a disproportionate increase in extracellular matrix. Largely composed of collagen, this process is not unique to renal disease, because matrix accumulation is found in all organ parenchyma following injury. Moreover, a growing number of studies suggest that parallels exist between renal scar formation and dermal wound healing (1). However we now increasingly recognize that pathological scarring is due not only to increased synthesis and decreased degradation of matrix, but also the reorganization of this matrix to increase its density. This chapter describes an in vitro model of this process that can be used to study the mechanisms and regulation of parenchymal contraction in chronic injury renal disease. From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_14, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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The presence of fibroblasts in wounded tissue is an established phenomenon. The role of renal fibroblasts, like that of the mesangial cell in the injured glomerulus, appears to be multi-functional. They form stress fibers and are thought to be responsible for the contraction process observed in tissue healing of parenchymatous organs. Both are of mesenchymal origin, possess contractile features and express 1 integrins, cell receptors necessary for cellular binding to collagen I fibers (2). It is also well documented that both cell types actively produce proteins implicated in matrix remodeling, including collagens I, III, and V and fibronectin. In general, mesangial cells display similar phenotypic and behavioural characteristics to fibroblasts in vitro. This is not altogether surprising and has resulted in mesangial cells being referred to as the myofibroblasts of the glomerulus (3). Originally described by Bell et al. (4), the cell-populated collagen lattice provides the opportunity to study the mechanisms of cell-mediated contraction and cell–matrix interactions. Cells are transferred into liquid collagen I, which is then polymerized, thereby entrapping the cells in a matrix of collagen I fibrils. In this way, cells can be cultured, maintained, and studied within a three-dimensional matrix, a situation perceived to be similar to an in vivo parenchymal environment. Interactions between cells themselves and their environment can be observed. When cultured within a lattice comprised of collagen, cells are known to reorganize individual collagen fibrils and increase matrix density (Fig. 14.1 and 14.2). Furthermore, by using the lattices as either tethered (anchored) or free floating, the influence of stress fiber formation on the repair process can be observed. These two types of collagen lattice, although similar, are thought to represent very different stages of wound healing. Within the free-floating lattice environment, contractile forces are generated by cells almost immediately after

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 (see color plate 7).

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Fig 14.2 Contraction of collagen lattices by rat mesangial cells (Rat Mes), human renal fibroblasts (HRF), and rat renal fibroblasts (Rat Fib). Lattices populated with Hela cells have been included as a negative control.

the collagen lattice has formed, but unlike cells in attached collagen lattices, they are not able to generate and maintain tension lines (5). Thus it is proposed that free-floating and attached collagen lattices represent the early and later stages of wound healing, respectively (6). Further to this, free-floating collagen lattices such as those described below provide a method to study in isolation the effects of given substances or physiological factors on cell-mediated contraction of extracellular matrix. For this reason, this system has been used in interventionist studies where the effects of different parameters on the rate of contraction were examined (7–9). The protocol described here uses fibroblast-populated collagen lattices. The technique can and has been adapted by others to examine mesangial cells and vascular smooth muscle cell contraction (Fig. 14.2).

2

Materials

This protocol assumes a general knowledge of aseptic cell culture techniques, which are required for the procedure. All solutions are sterile (see Note 1).

2.1

Cell Culture

1. Autoclave unit. 2. 100-, 200-, and 500-mL Schott™ bottles (Schott, Duran, Germany).

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3. Sterile deionized water (sterile dH2O). 4. 0.01 M Phosphate-buffered saline (PBS): reconstitute each PBS tablet (Oxoid, Hampshire, UK) in 100 mL of sterile dH2O. 5. Filter unit (300-mL capacity) with 0.2-µm cellulose acetate filters (Advantec MFS Inc., Dublin, CA, USA) and vacuum pump. 6. Pipet-Aid™ (BD Falcon, BD Biosciences, San Jose, USA) or equivalent for aspirating and changing media. 7. Medium: for each 400 mL of medium, combine 40 mL filter-sterilized fetal calf serum (FCS; SAFC Biosciences, Sigma–Aldrich, St. Louis, MO, USA), 10 mL of 1 M Hepes (SAFC Biosciences), 4 mL of 200 mM glutamine (SAFC Biosciences), and 8 mL penicillin–streptomycin (formulated to contain 5,000 U/mL penicillin and 5,000 µg/mL streptomycin; Sigma–Aldrich) and make up to 400 mL with 1× Dulbecco’s Modified Eagle Medium (DMEM; SAFC Biosciences). This solution is sterilized by passing through a sterile 0.2-µm cellulose acetate filter unit into a sterile 500-mL glass bottle by vacuum extraction. The bottle is capped and medium is stored at 4°C (see Note 2). 8. Trypsin–EDTA (1×) (SAFC Biosciences). 9. Trypan blue (Sigma–Aldrich). 10. Borosilicate 5-mL glass tubes (Chase-Scientific, Rockwood, TN, USA). 11. Haemocytometer. 12. Manual tally counter. 13. 50-mL Polypropylene tubes (TPP, Trasadingen, Switzerland). 14. 25-cm2 and 75-cm2 cell culture flasks with vent caps (TPP). 15. Disposable sterile syringes. 16. Disposable 0.2-µm pore syringe filters. 17. Disposable 5- and 10-mL sterile pipettes (Costar Corning Incorporated, Corning, NY, USA). 18. Tissue culture incubator set at 37°C with 95% O2/5% CO2.

2.2

Collagen Lattices

1. 0.3% acid-solubilized collagen I in acetic acid, pH 3.0 (Cellagen™; MP Biomedicals, Solon, OH, USA). 2. Sodium bicarbonate: 11.76 mg/mL made up in sterile deoionized H2O. 3. MEM (10× concentrate) (SAFC Biosciences). 4. Glass bowl filled with ice. 5. Magnetic stirrer. 6. Sterile scalpel blades (No. 22). 7. 12-well cluster plates (well diameter of 24 mm). 8. Perspex or transparent metric ruler. 9. Tissue culture incubator set at 37°C with 95% O2/ 5% CO2.

14 In Vitro Model of Parenchymal Contraction

2.3

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Histology

1. Fixative, e.g. neutral buffered formalin, 4% paraformaldehyde or mercuric formalin. 2. 100% Alcohol. 3. Chloroform. 4. Paraplast™ (McCormick Scientific, St. Louis, MO, USA). 5. dH2O. 6. 10-mL Borosilicate bottles. 7. Disposable pipettes. 8. Timer. 9. Incubator set at 60°C. 10. Microtome. 11. Silane-coated glass microscope slides. 12. Incubator set at 40°C. 13. 0.01 M phosphate-buffered saline. 14. Xylene. 15. 100%, 95%, and 75% alcohol (prepared in distilled water). 16. Tap water. 17. Staining jars. 18. Primary antibody of choice. 19. Fluorescein isothiocyanate (FITC)-conjugated anti-IgG appropriate for primary antibody (Dakocytomation, Glostrup, Denmark): diluted 1:50 in antibody diluent (Dakocytomation). 20. Humidified staining chamber (lunch box containing paper towels saturated with 0.01 M PBS). 21. Aluminum kitchen foil. 22. Aquamount™ (BDH, Poole, UK). 23. Cover slips. 24. Forceps. 25. Fluorescent microscope.

3

Methods

The contribution of renal mesenchymal cells (fibroblasts, vascular smooth muscle cells, and mesangial cells) to the scarring process through reorganization of matrix can be studied in an in vitro system utilizing a cell-populated collagen I lattice. Lattice experiments provide a method of studying rapid cell-mediated contraction of collagenous matrix in a controlled experimental environment. Within free-floating collagen lattices, such as those described here, contractile forces are generated by cells almost immediately after the collagen lattice has formed (5). Collagen I is used here because it is a principal extracellular matrix component of renal fibrosis (10).

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Preparation of Fibroblasts

All cell culture procedures are carried out in a biohazard hood that meets standard requirements for handling cell culture material in a sterile environment. 1. Cells are routinely grown in Hepes-buffered DMEM containing 10% FCS, penicillin, and streptomycin. The media is heated to approximately 37°C for 15 min in a water bath prior to treating cells. 2. At confluence, media is removed using a sterile pipette. The cell monolayer is washed twice using 2.5 mL of 0.01 M PBS solution. Cells are incubated for 5 min with 1× trypsin–EDTA solution at 37°C in a standard cell culture incubator. After this time, trypsin action is inhibited by adding 5 mL of growth media to the flask. 3. Dislodged cells present in the media are then collected by aspiration using a sterile pipette and placed into a sterile 50-mL polypropylene tube. A cell pellet is formed by centrifugation at 3,000 × g for 5 min. The supernatant is removed and the cell pellet is resuspended in 2 mL of fresh sterile growth media. 4. 100 µL of cell suspension solution is diluted 1:1 with 0.4% trypan blue in a 5-mL borosilicate tube. The stain is allowed to penetrate dead cells over a period of 3–5 min at room temperature. Trypan blue is one of several stains recommended for use in dye-exclusion procedures for viable cell counting. The method is based on the principle that living cells do not absorb trypan blue whereas dead cells do absorb the dye and appear blue upon visualization (see Notes 3 and 4). 5. Set up the haemocytometer with the cover slip. Draw up 50 µL of cell/trypan blue solution and slowly transfer into both chambers of the haemocytometer at the edge of cover slip with pipette tip and allow each chamber to fill by capillary action. 6. Count the viable cells in each chamber of the haemocytometer using a tally counter. Move from left to right, counting the four corner 1-mm squares and the 1-mm centre square (non-viable cells stain blue). 7. Repeat the cell count for chamber 2. 8. Determine the average cell count for both chambers i.e. the sum of total number of cells divided by two. 9. Determine the total number of cells with the formula: Total number cells = C/Sq × 104 × D × V where “C” is the number of cells counted; “Sq” is the number of squares counted (usually five); “104” is the depth of the haemocytometer; “D” is the dilution factor (usually two if equal volumes of cell suspension and trypan blue are used); and V is the remaining volume of cell suspension. For example, if the average number of cells is 438; then: 438/5 × 104 × 2 × 1.9 mL = 3.33 × 106. Thus, 3.33 × 106 is the total number of cells in 1.9 mL media. 10. 2×105 cells are required for each 2-mL collagen lattice (see Notes 5 and 6).

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11. The volume of cell suspension containing 2 × 105 cells is then placed in a sterile tube. Cells are pelleted by centrifugation at 3,000 × g for 5 min and resuspended in 200 µL medium (DMEM and 10% FCS) (see Note 7).

3.2 Preparation of Collagen Lattices Lattices are prepared on ice under sterile conditions and using sterile ingredients (see Note 8). 1. The basic lattice recipe consists of sodium bicarbonate (11.76 mg/mL), 10× concentrated MEM (prepared in sterile water) and liquid collagen I in a volume ratio of 1:1:7 (see Notes 9 and 10). 2. Calculate the total number of wells required (a minimum of n = 3 per group). 3. For each well (lattice) combine 200 µL sodium bicarbonate, 200 µL of 10× MEM and 1.4 mL liquid collagen I. 4. Mix ingredients together in a 50-mL sterile glass beaker containing a small magnetic stirring bead placed on top of a magnetic stirring device. 5. 1.8 mL cold liquid lattice mixture is then placed into the well of a 12-well cluster plate. To this, the 200 µL cell suspension is added and mixed by aspiration. 6. The collagen is then allowed to polymerize at 37°C in a standard cell culture incubator for 10–20 min (see Note 11). 7. Upon polymerization, the plate is removed from the incubator and placed in a sterile cell culture hood. 8. The solid lattice is gently detached from the rim of the cluster plate well using a sterile scalpel blade. This is done by running the blade around the edges of the well (rimming). 9. 2 mL Growth medium is then placed over each lattice, before the plate is returned to the incubator (see Note 12).

3.3

Measuring Collagen Lattice Contraction

1. Initial lattice diameter, prior to release, is equivalent to the diameter of the cluster plate used. 2. Reduction in lattice diameter is measured by placing the lattice over a transparent ruler on a light box, then recording the diameter size in millimeters (see Notes 13 and 14). 3. Measurements are taken at the time (T) of lattice release (T0) and at 24 h and 48 h post-release (T24 and T48, respectively) (Fig. 14.2). 4. Readings at each time point are then subtracted from the initial lattice diameter (T0) to obtain a percentage reduction over time (see Notes 13, 14, and 15). 5. Alternatively, reduction in lattice diameter can be expressed as a reduction in lattice area using the formula: area = r2.

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Processing Collagen Lattices for Histology

Upon completion of contraction experiments, lattices may be fixed and crosssectioned. Upon cross-sectioning, cells within the lattices may be examined using immunohistochemical techniques. Manual processing of collagen lattices is performed in a standard laboratory fume hood at room temperature. All waste solutions should be collected and disposed of using appropriate protocols. 1. Remove individual collagen lattices from culture dish and place into a 10-mL clean borosilicate bottle. Cover lattice with chosen fixative and incubate for 2 h (see Note 16). 2. Remove fixative using a pipette and replace solution with 50% alcohol (prepared in deionized distilled water) for 30 min. 3. Remove 50% alcohol and replace solution with 70% alcohol for 30 min. 4. Remove 70% alcohol and replace solution with 95% alcohol for 30 min. 5. Remove 95% alcohol and replace solution with 100% alcohol for 30 min, 2×. 6. Remove alcohol and add chloroform for 1 h, 2×. Agitate gently throughout (see Note 17). 7. Remove chloroform. Add approximately 8 mL of Paraplast™ and incubate for 1 h at 54–60°C in an incubator to allow wax infiltration. 8. Remove Paraplast™ and add a further 8 mL fresh molten Paraplast™. Incubate for 1 h at 54–60°C (see Note 18). 9. Lattices can then be embedded using standard histological procedure. 10. 2-µm Sections are cut using a microtome and collected onto clean silane coated slides. Sections are dried overnight at 40°C in an incubator prior to staining.

3.5

Immunofluorescent Staining of Lattice Cross-Sections

Immunohistochemistry is a technique that allows the localization of specific antigens in histological tissues using antibodies derived from the serum of immunized animals or immortal cell lines. Either monoclonal or polyclonal antisera may be used. The antigen–antibody complex is then detected using a secondary antibody conjugated to FITC (see Note 19). Immunofluorescence procedures are performed at room temperature in a humidified staining chamber unless specified otherwise. 1. Dewax sections as per standard histological protocol. 2. Sections are incubated in the normal serum of the species in which the primary antiserum is raised for 20 min. 3. The primary antiserum is diluted to an appropriate working concentration with an antibody diluent. Sections are covered in prepared antibody and incubated overnight at 4°C. 4. Sections are removed from the chamber and washed three times in 0.01 M PBS for 5 min each time.

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5. FITC-conjugated secondary antibody is diluted 1:50 in 0.01 M PBS. Sections are covered with prepared antibody and incubated for 30 min (see Notes 20 and 21). 6. Sections are removed from the chamber and washed three times in 0.01 M PBS for 5 min each time. 7. Sections are cover slipped using Aquamount™ and staining is viewed using a fluorescent microscope (see Note 22).

4

Notes

1. This technique is performed in a biohazard hood suitable for cell culture use. All equipment and solutions need to be sterilized using an autoclave. In the event that this is deemed unsuitable, equipment should be thoroughly wiped down using 70% alcohol. Solutions should be passed through a 0.2-µm syringe filter prior to use. 2. The individual components of cell culture media are prepared according to manufacturers’ instructions. DMEM and Hepes are aliquoted and stored in sterile bottles at 4°C. FCS, glutamine, and penicillin–streptomycin solution are stored at –20°C in 10-mL in aliquots. 3. An haemocytometer is used to quantitate the number of cells in suspension that do not absorb the blue dye, i.e. the number of viable cells. Each large square on the haemocytometer is 1 ×1 × 0.1 mm, thus giving a volume of 0.1 mm3 or 1 ×10–4 cm3 (mL). Also it is important to note that the volume used to resuspend pellet may be adjusted and must be incorporated into the calculations accordingly. 4. Using the cell number derived by haemocytometer, a total cell number in the remaining 1.90 mL of cell suspension can be calculated. 5. Cell number can be varied by using less collagen solution in wells of lesser diameter. The degree of contraction observed within lattices is directly proportional to the number of cells seeded into each lattice. 6. The volume of each lattice can be altered, allowing for the use of plates of different sizes or differing numbers of cells by following the ratio of 7:1:1:1, where seven times the volume of collagen is used to one times the volume of 10× αMEM, sodium bicarbonate, and cell suspension volume for each lattice (see also Note 7). 7. In the case of experiments in which reagents are being tested for a potential effect on cellmediated contraction, cells may be resuspended in base medium containing the test substance at the desired concentration. They will then be mixed with the liquefied lattice solution and polymerized accordingly. However, it may be useful to perform a toxicity assay before performing contraction studies to rule out cell death as a contributor to any decrease in contraction observed. 8. All work is performed in a biohazard hood suitable for cell culture unless stated otherwise. Equipment introduced into the sterile environment for the purpose of preparing collagen lattices is wiped thoroughly with 70% alcohol. 9. The basic lattice ratio of 1:1:7 (and 1× cell suspension volume) is the optimal recipe for the experimental parameters described here. When altered, the time required for lattices to polymerize may change. Likewise the time needed for cells to reorganize and contract lattices may differ from that detailed in this method. 10. Collagen I was chosen on the basis of being a principal extracellular matrix component of renal fibrosis. However, the addition of other forms of liquid collagens may be substituted for collagen I. This system may then be used to study the contractile effects of mesenchymal cells in different collagenous environments. 11. The coverplate is placed over the top of the culture dish before it is removed from the biohazard hood and placed in a standard 95% air/5% CO2 cell culture incubator. This is to ensure that the lattice remains sterile.

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12. In the case of experiments in which reagents are being tested for a potential effect on cell-mediated contraction, they should be prepared at the desired concentrations in base medium and added to the lattices following lattice polymerization. 13. Due to the semi-transparent nature of the collagen lattices, their diameter can easily be measured using a Perspex metric ruler when placed on top of a light box (or a similar light source). Alternatively, diameter measurements can be made by holding a Perspex ruler to the underside of the cell culture dish. 14. Measurements should be taken at desired time points. For most experiments, measurements should be taken at T0 (immediately after rimming of lattices has been performed), T24 (24 h after rimming of lattices) and T48 (48 h after lattice rimming). However, experiments in which cells are contracting lattices at a particularly fast rate may require measurements at 2-h intervals. 15. Other methods of measuring lattice diameter over time include scanning the lattices in situ at given time points using a standard computer scanner. The images can then be transferred to an image analysis program and diameter and area measurements obtained. Photocopied images may also be useful for calculating diameter and area of lattices. 16. Lattices may be fixed in mercuric formalin, 10% neutral buffered formalin, or 4% paraformaldehyde. 17. Agitation aids dehydration and clearing of the lattice material. This is best achieved by placing 10-mL bottles on a shaking device. However, it can also be achieved by gently shaking bottles by hand every 10 min. 18. After this 1 h incubation, bottles containing the lattices can be removed from the incubator and wax allowed to harden. The lattices can then be stored in this state and embedded at a later time if required. 19. Other methods of detection may be used such as avidin–biotin complex or fluorochromeconjugated secondary antibodies at appropriate concentrations. 20. FITC-conjugated antibodies are light sensitive so they should be shielded from light wherever possible. This can be achieved by wrapping the tube in which antibody is diluted in aluminium foil until it is added to the sections. 21. The humidified chamber should be wrapped in aluminum foil to prevent bleaching of FITCconjugated secondary antibody. 22. Cover slipping of slides should be carried out quickly. Slides should be stored in aluminum foil between cover slipping and viewing.

References 1. Wardle, E. (2000) Modulatory proteins and processes in alliance with immune cells, mediators and extracellular proteins in renal fibrosis. Renal Failure 21, 121–133. 2. Kelynack, K.J., Hewitson, T.D., Nicholls, K.M., Darby, I.A., and Becker, G.J. (2000) Human renal fibroblast contraction of collagen I lattices is an integrin mediated process. Nephrol. Dial. Transplant. 15, 1766–1772. 3. Johnson, R., Floege, J., Yoshimura, A., Iida, H., Couser, W., and Alpers, C. (1992) The activated mesangial cell: a glomerular myofibroblast? J. Am. Soc. Nephrol. 2, S190–S197. 4. Bell, E., Ivarsson, B., and Merrill, C. (1979) Production of a tissue like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential in vitro. Proc. Natl. Acad. Sci. USA 76, 1274–1278. 5. Tomasek, J., Haaksma, C., Eddy, R., and Vaughan, M. (1992) Fibroblast contraction occurs on release of tension in attached collagen lattices: dependency on an organized actin cytoskeleton and serum. Anat. Rec. 232, 359–368.

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6. Grinnell, F. (1994) Fibroblasts, myofibroblasts and wound contraction. J. Cell Biol. 124, 401–404. 7. Hewitson, T.D., Martic, M., Kelynack, K.J., Pagel, C., Mackie, E.J., and Becker, G.J. (2005) Thrombin is a pro-fibrotic factor for rat renal fibroblasts in vitro. Nephron-Exp. Nephrol. 101, e42–e49. 8. Hewitson, T.D., Martic, M., Kelynack, K.J., Pedagogos, E., and Becker, G.J. (2000) Pentoxyfilline reduces in vitro renal myofibroblast proliferation and collagen secretion. Am. J. Nephrol. 20, 82–88. 9. Kelynack, K.J., Hewitson, T.D., Martic, M., McTaggart, S., and Becker, G.J. (2002) Lovastatin downregulates renal myofibroblast function in vitro. Nephron 91, 701–707. 10. Sharma, A, Mauer, S, Kim, Y, and Michael, A. (1993) Interstitial fibrosis in obstructive nephropathy. Kidney Int. 44, 774–788.

Chapter 15

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

Abstract The investigation of the effects of mechanical stretch on mesangial cells can provide important insights into glomerular pathophysiology related to increased intraglomerular pressure (Pgc). Elevated Pgc, leading to the transmission of abnormal mechanical stress to resident renal glomerular cells, is a major pathogenic factor in progressive glomerular fibrosis and the ultimate loss of renal function. Mesangial cells, at the center of the glomerular tuft, produce extracellular matrix components when exposed to mechanical stretch, thus providing a model system simulating in vivo pathology. Stretch-induced activation of the mitogen-activated protein kinase Erk has been linked to matrix production. The integrity of the actin cytoskeleton, an important transmitter of mechanical signals, is required for activation of Erk and its upstream kinase Raf-1. The GTPase RhoA, a central regulator of the actin cytoskeleton, is also activated by stretch, and both Erk activation and subsequent matrix upregulation are dependent on its function. The investigation of signaling pathways activated by mechanical stretch and associated with abnormal matrix production or regulation will assist in the development of new therapies targeted at slowing the progression of renal disease.

Keywords Mechanical stress, Mesangial cell, Erk, Raf-1, Actin cytoskeleton, RhoA, Extracellular matrix, Immunofluorescence, Fibronectin, ELISA

1

Introduction

Increased glomerular capillary pressure (Pgc) is an important hemodynamic determinant of progression of glomerular sclerosis and hence chronic renal impairment of diverse etiology (1). Interventions that normalize Pgc, such as interruption of the renin–angiotensin system, reduce glomerular injury and matrix expansion (2). Increased Pgc transmits to mesangial cells (MC), which provide architectural support for the glomerular capillary tuft, as mechanical strain (3). MC that are subjected to cyclic strain alternating with relaxation increase extracellular matrix protein synthesis, recapitulating the increased matrix deposition characteristic of From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_15, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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glomerular sclerotic injury (4, 5). In this chapter, I outline techniques for the application of mechanical stress to the study of MC pathophysiology. Both in vivo and in cultured MC, activation of the mitogen-activated protein kinase Erk by mechanical strain induces activation of the transcription factor AP-1 (6, 7), with subsequent accumulation of extracellular matrix proteins such as fibronectin (8). We and others have shown that stretch induces the formation of filamentous actin (F-actin) stress fibers in MC (7, 9) and that disruption of the actin cytoskeleton prevents stretch-induced Erk and AP-1 activation and downstream fibronectin upregulation (7, 10). Membrane translocation and activation of Raf-1, the kinase upstream of Erk activation, also requires actin cytoskeletal integrity (11). Formation of F-actin stress fibers is predominantly regulated by the small GTPase RhoA (12). Indeed, stretch-induced activation of the Raf-1/Erk pathway and fibronectin upregulation is dependent on activation of RhoA and its downstream kinase Rho-kinase (10). Targeting of either Erk or RhoA signaling thus represents a potential treatment strategy in prevention of hemodynamically mediated glomerular injury. Numerous methods allow analysis of these two interrelated pathways, including Western blotting for phosphorylated forms of Raf-1 and Erk and specific activity assays, as well as imaging techniques.

2 2.1

Materials Cell Culture and Lysis

1. Dulbecco’s Modified Eagle’s Medium (DMEM) with 1,000 mg/L d-glucose, l-glutamine, and 110 mg/L sodium pyruvate (Invitrogen, Carlsbad, CA, USA): reconstitute powder with distilled water and 7.5% (w/v) sodium bicarbonate solution (Invitrogen) to a final concentration of 3.7 g/L sodium bicarbonate, and filter through a 0.22-µm bottle-top filter (Mississauga, ON, Canada ) (VWR, West Chester, PA, USA). Prior to use, medium is supplemented with 20% fetal bovine serum (FBS; Sigma–Aldrich, St. Louis, MO, USA; or Invitrogen) that has been heat inactivated at 55°C for 30 min, as well as 100 U/mL penicillin and 100 µg/mL streptomycin (Invitrogen). 2. Trypsin solution (0.5% with EDTA; Invitrogen): prepare working concentration at 1:3 dilution in phosphate-buffered saline (PBS), see below. 3. PBS, 10×: 1.36 M NaCl, 14.7 mM KH2PO4, 80 mM Na2HPO4, 26.8 mM KCl with pH adjusted to 7.4. Dilute 100 mL with 900 mL water for use. 4. Freezing medium: DMEM as in item 1 above, with 10% DMSO (Sigma– Aldrich). 5. Cell lysis buffer: 50 mM Tris pH 7.4, 150 mM NaCl, 1% Triton X-100, 10% glycerol, 5 mM EDTA, 100 µM sodium vanadate, 1 mM β-glycerophosphate, 1 mM

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sodium fluoride, 2 µg/mL leupeptin, 10 µg/mL aprotinin, and 1 mM PMSF. Buffer is prepared without the final three components and stored at 4°C (see Note 1). 6. Polypropylene cell lifters (Fisher, Pittsburgh, PA, USA).

2.2

Setting Up the Stretch Apparatus

Mechanical strain may be applied to cultured cells in a number of different ways. We routinely use a Flexcell Tension PlusTM system (Flexcell International, Hillsborough, NC, USA) in our laboratory. The following components are required: 1. Tension Plus System, FX-4000TTM (Flexcell International) including computer (controller), baseplate and gasket, FlexLinkTM, drying filter, and water trap (see Note 2). 2. Vacuum pump capable of generating a maximum vacuum of 0.02 torr and airflow rate of 6 cfm (Flexcell International or other supplier). Oil-free pumps generate less noise and require less maintenance. 3. Bioflex plates (Flexcell International), coated with bovine collagen I (see Note 3). 4. FlexstopTM reusable rubber stoppers (Flexcell International).

2.3

Assay of Active Erk and Raf-1 by Western Blotting

1. Bradford Protein Assay (BioRad Laboratories, Hercules, CA, USA). 2. Reducing sample buffer, 5×: 0.1 M Tris pH 6.8, 5% sodium dodecyl sulfate (SDS), 50% glycerol, 10% β-mercaptoethanol, and 0.03% (w/v) bromophenol blue. This is stored in aliquots at –20°C. 3. Tris-buffered saline with Tween-20 (TBST), 10×: 165 mM Tris-HCl, 1.37 M NaCl pH 7.6, and 1% Tween-20. Dilute 100 mL with 900 mL water for use. 4. Blocking buffer: 5% (w/v) nonfat milk in TBST. 5. Bovine serum albumin (BSA, biotechnology grade; Bioshop, Burlington, Ontario, Canada). 6. Primary antibody dilution buffer: 5% albumin in TBST. 7. Primary antibodies: rabbit or mouse anti-phospho Erk1/2 Thr202/Tyr 204 (Cell Signaling, Danvers, MA, USA); mouse anti-phospho Raf-1 Ser338 (Upstate Biotechnology, Charlottesville, VA, USA); rabbit anti-Erk1/2 (Cell Signaling); and rabbit anti-Raf-1(C-12; Santa Cruz Biotechnology, Santa Cruz, CA, USA). 8. Secondary antibodies: goat anti-rabbit or anti-mouse (Bio-Rad Laboratories). 9. ECL (Amersham Biosciences, Little Chalfont, Buckinghamshire, UK). 10. X-Omat Blue XB-1 Kodak film (Perkin Elmer, Waltham, MA, USA).

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Assessment of Membrane Translocation of Raf-1

1. Hypotonic lysis buffer for cytosol-membrane fractionation: 10 mM Tris pH 7.5, 100 µM sodium vanadate, 1 mM β-glycerophosphate, 1 mM sodium fluoride, 2 µg/mL leupeptin, 10 µg/mL aprotinin, and 1 mM PMSF. Buffer is prepared without the final three components and stored at 4°C (see Note 1). 2. Needles, 30½ gauge. 3. Syringes, 1 mL. 4. Ultracentrifuge and tubes.

2.5

Raf-1 Activity Assay

1. Raf-1 activity assay kinase buffer: 25 mM Tris-HCl pH 7.2, 10 mM MgCl2, 2 mM DTT, 100 µM sodium vanadate, and 5 mM β-glycerophosphate. This is prepared as a 10× buffer and stored in aliquots at –20°C. 2. Recombinant protein G agarose (Invitrogen). 3. Thermomixer (Eppendorf, Westbury, NY, USA) or water bath. 4. Kinase mix: each reaction requires 20 µL kinase buffer diluted to 1× concentration with deionized water, with the addition of the following for each 20 µL: 200 µM ATP (Cell Signaling), 0.4 µg MEK1 (Upstate; stored at –70°C, and thawed on ice immediately prior to use), and 2 µg of kinase-inactive Erk2 (Cell Signaling; stored at –20°C). 5. Reducing sample buffer, 2×, diluted from 5× stock with deionized water.

2.6

Assay of RhoA Activation

1. RhoA activity cell lysis buffer: 50 mM Tris pH 7.2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 500 mM NaCl, 10 mM MgCl2, 60 mM n-octyl glucopyranoside, 2 µg/mL leupeptin, 10 µg/mL aprotinin, and 1 mM PMSF. Buffer is prepared without the last four components and stored at 4°C (see Note 1). Glucopyranoside is also made and added prior to each use. 2. Glutathione S-transferase (GST)–rhotekin Rho-binding domain (RBD) bound to glutathione–agarose (Upstate; or Cytoskeleton, Denver, CO, USA), stored at –70°C. 3. RhoA activity wash buffer: 50 mM Tris-HCl pH 7.2, 1% Triton X-100, 150 mM NaCl, 10 mM MgCl2, 2 µg/mL leupeptin, 10 µg/mL aprotinin, and 0.1 mM PMSF (see Note 1). 4. Primary antibody: mouse anti-RhoA (Santa Cruz Biotechnology). 5. SuperSignal West Femto Luminol/Enhancer Solution (Pierce, Rockford, IL, USA).

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209

Confocal Immunofluorescence for Raf-1 Membrane Localization

1. 2. 3. 4. 5.

Fixative solution: 37% formaldehyde solution, diluted to 3.7% in PBS. Permeabilization solution: 0.2% Triton X-100 in PBS. BSA, biotechnology grade (Bioshop). Donkey serum (Sigma–Aldrich) (see Note 4). Blocking buffer: 1% BSA, 3% donkey serum, 0.1% Triton X-100, and 0.1% sodium azide, which is added when serum or albumin is used to prevent bacterial growth. 6. Secondary antibodies: fluorochrome conjugated from Jackson ImmunoResearch (West Grove, PA, USA), adsorbed against the species from which the MC were obtained.

2.8

Confocal Immunofluorescence for F- and G-Actin

1. Blocking buffer: 1% BSA in PBS. 2. Rhodamine phalloidin for F-actin staining (Molecular Probes, Eugene, OR, USA). 3. DNase I-Alexa 488 for globular actin (G-actin) staining (Molecular Probes; see Note 5). 4. Coverslips, 22×22 mm. 5. Clear nail polish. 6. Vectashield mounting medium, with DAPI to visualize nuclei (Vector Laboratories, Burlingame, CA, USA).

2.9

Assessment of Fibronectin by ELISA

1. 96-well Flat-bottom plates (Sarstedt, Newton, NC, USA; or Costar Corning, Corning, NY, USA). 2. Multichannel pipetter. 3. Fibronectin from rat plasma (Sigma–Aldrich). 4. Microplate shaker. 5. Microplate reader. 6. Antibodies: mouse anti-fibronectin (BD Transduction, BD Biosciences, San Jose, CA, USA) or rabbit anti-fibronectin (Sigma–Aldrich), goat anti-mouse or antirabbit alkaline phosphatase-conjugated secondary antibody (Sigma–Aldrich). 7. Enzyme-linked immunosorbent assay (ELISA) coating buffer (ECB): carbonate-bicarbonate buffer, pH 9.6, reconstituted from capsules in 100 mL deionized water (Sigma–Aldrich). 8. Blocking buffer: 2% BSA, 0.05% Tween-20, and 0.05% sodium azide in PBS (diluted from 10× PBS, see Note 6).

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9. Dilution buffer: 0.25% BSA, 0.05% Tween-20, and 0.05% sodium azide in PBS. 10. Wash buffer: 0.05% Tween-20 and 0.05% sodium azide in PBS. 11. p-Nitrophenyl phosphate (pNPP) substrate (Sigma–Aldrich).

3

Methods

Erk is a component of the mitogen-activated protein kinase cascade, which consists of the sequential activation of three kinases, Raf-1, MEK1/2, and Erk1/2, with signal amplification occurring at each step (13). Detection of Erk activation by stretch is thus quite robust. Erk is activated by dual phosphorylation of Thr202/ Tyr204, and antibodies that detect phosphorylation at these sites provide a sensitive measure of Erk activation (14). Raf-1 activation requires the phosphorylation of Ser338, and this can be assessed by Western blotting using phosphorylation site-specific antibodies (15). However, incomplete correlation between phosphorylation of this residue and Raf-1 activation has been reported (16). Thus, the method for performing a Raf-1 activity assay is also presented. Additionally, since Raf-1 requires membrane targeting for full activation (17), two protocols assessing this are provided. The first protocol is the separation of cytosolic and membrane fractions with subsequent Western blotting for phospho-Raf-1 in the membrane isolates, and the second is immunohistochemistry, which allows visualization of Raf-1 location. Mechanical stretch has been shown to increase the formation of F-actin stress fibers, and disruption of the actin cytoskeleton prevents Raf-1/Erk activation (9, 11). A method for assessment of actin turnover by immunofluorescence is provided. RhoA is a central regulator of F-actin formation, requiring both membrane translocation and GTP binding for full activation (18, 19). Although assessment of membrane translocation has been used as a surrogate for RhoA activity, a more direct assessment of activity involves a pull-down assay of GTP-bound RhoA. Due to the intrinsic hydrolytic activity of RhoA, the GTP-bound state is relatively unstable and the assay must be performed quickly with attention paid to keeping the lysate in cold conditions throughout the procedure. Ultimately, an assessment of the requirement of signaling cascades or proteins for the stretch-induced production of matrix proteins is important. Although several methods can be utilized, secretion into the extracellular space is required for the laying down of matrix in vivo. Thus, assessment of secreted fibronectin by ELISA is presented.

3.1

Preparation of Primary Mesangial Cells for All Assays

1. MC (isolated from male Sprague-Dawley rats), are passaged at confluence with 1:3 trypsin. For early passage cells (up to six passages), plating at higher density is required for adequate growth (3–4×105 cells/100-mm dish). Otherwise,

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1–2 × 105 cells/100 mm is adequate for passaging. When plating 6-well BioFlex plates for experiments, cell density per well is equivalent to that used for 100mm plates. Cells are left to grow to full confluence, then rinsed once in PBS and serum deprived for 24 h. For all experiments except for longer-term stretch to assess matrix production, cells are placed into 0% FBS medium for serum deprivation. In the latter case, 0.5% FBS medium is used to improve matrix protein recovery and avoid cell death. 2. MC are used to a maximum of passage 20. If altered cell morphology is observed prior to this, cells are discarded. 3. Primary MC can be cryopreserved at early passage for later use. After trypsinization, MC are centrifuged at 4°C, 1,700×g for 4 min. They are resuspended at 1×106 cells/mL in prepared freezing medium. Cryovials with 1 mL of cells are encased in Styrofoam (used for 15-mL tubes) and left for 24–48 h at –70°C, then quickly transferred into liquid nitrogen storage. 4. Thawing of cryopreserved MC is performed by placing cryovials into a 37°C water bath for 1–2 min (until almost thawed), then gently pipetting to fully thaw and dividing one vial into two 100-mm plates. Medium is replaced the following day.

3.2

Working with the FX-4000TM System

The tension plus apparatus is a computer-regulated bioreactor that uses vacuum pressure to apply cyclic or static strain to cultured cells. To set up the apparatus, cells are grown on matrix-coated flexible-bottomed culture plates. Application of a vacuum stretches the cells by distorting the flexible culture surface. Subsequent release of the vacuum returns the culture plate bottom to its original configuration. The process is regulated by a microprocessor and solenoid values so that the magnitude, duration, and frequency of the applied force can be varied. Detailed instructions for the set up and operation of the system are provided by the manufacturer. The following is a list of points that may be helpful in its application to experiments. 1. Several waveforms are available depending on conditions being simulated. We use a sinusoidal waveform, which simulates pulsatile flow. 2. The degree of stretch can be varied, and the study of novel pathways activated by stretch can include dose–response studies. We have found maximal responses at 10% stretch for activation of the pathways discussed. Higher degrees of stretch are achievable, but result in MC detachment, particularly with stretch of longer duration. Minimum stretch is also programmed, and our laboratory uses 3%. 3. Frequency of stretch is chosen, and this varies significantly among different groups. To simulate pulsatile flow, we use 1 Hz, with 0.5 s of stretch and 0.5 s of relaxation. 4. A drying regimen is available, and it is suggested that this is used after stretch protocols lasting longer than 10–15 min. This helps to prevent moisture

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accumulation in the valves. The beads in the drying filter should be blue, and with water saturation, they become pink in color. The entire filter can be dried in an oven at 55°C; this requires overnight incubation. 5. The FlexLinkTM unit needs to be turned on before the computer. Failure to do this will result in an inability of the system to load a stretch regimen. 6. Control wells can be set up in a 6-well plate by placing rubber stoppers in the underside of the wells. These can be cleaned with 70% ethanol prior to insertion. Since inserting these can cause movement of the membrane to which the cells are attached, we place these at the time of serum deprivation (24 h before initiation of stretch). The plate is then placed into the baseplate in the incubator to keep it level. After stretch, the plates are washed and lysis buffer added, after which the stoppers are removed. 7. For short stretch time courses, one plate is stretched at a time to allow rapid plate processing to preserve the phosphorylation status of proteins.

3.3

Assay of Active Erk and Raf-1 by Western Blotting

1. After serum deprivation for 24 h, MC are stretched for the desired times. Maximal early Erk activation occurs at 10–20 min and Raf-1 at 2–5 min (Fig. 15.1a) (11). One well per condition provides sufficient protein quantity for assessment of activities. 2. After stretch, the plate is promptly removed from the baseplate and cells are washed three times with cold PBS (see Note 7).

Fig. 15.1 Stretch induces the activation of the Raf-1 and Erk. a MC are stretched for times up to 5 min, with Raf-1 and Erk activation assessed by Western blotting with phosphorylation-specific antibodies to Raf-1 Ser338 and Erk Thr202/Tyr204. Aliquots of samples are run for assessment of total Raf-1 and total Erk. b After stretch for up to 5 min, Raf-1 activity is assessed as described in Sect. 3.5. Equal immunoprecipitated Raf-1 is confirmed by reprobing membranes with the Raf-1 antibody. Con, control

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3. Lysis buffer at 40–50 µL is added to each well, ensuring that it is spread out evenly across the entire surface. Rubber stoppers are removed from any control wells and the plate is incubated on ice for 10 min (see Note 8). 4. Using a cell lifter, lysed cells are harvested into tubes precooled on ice, and pipetting up and down with a 200-µL tip assists in further breaking up the cells. 5. Lysates are cleared by centrifugation for 10 min at 16,200×g, 4°C. The supernatants are transferred to a new precooled tube. 6. Protein concentration is determined using the Bradford Protein Assay, measuring OD at 595 nm. The desired protein quantity is aliquoted into a new cold tube, and samples are boiled with 5× sample buffer for 5 min in a heat block set to 100°C. Twenty-five micrograms is sufficient to detect phospho-Erk Thr202/ Tyr204, but 50 µg is required for phospho-Raf-1 Ser338. 7. After boiling, samples can be stored at -20°C until they are run on an SDS-polyacrylamide gel electrophoresis (PAGE) using standard protocols. A 10% gel is used for both phospho-Erk and phospho-Raf-1 detection. 8. For Western blotting, the nitrocellulose membrane is blocked for 1 h at room temperature, then washed three times by swirling in TBST (enough to generously cover the membrane). 9. Prior to adding the primary antibody, all TBST is removed by suction. Primary antibody in dilution buffer (10 mL) is added to the membrane. For phospho-Erk Thr202/Tyr204, dilutions of 1:2,000 for mouse and 1:1,000 for rabbit antibodies work well. For phospho-Raf-1 Ser338, 1 µg/mL is used (see Note 9). 10. Membranes are incubated overnight at 4°C on a shaker at 50 rpm, after which the primary antibody is removed and membranes washed three times with TBST (enough to generously cover the membrane). Washes are 6 min each at 150 rpm on a shaker (see Note 10). 11. Membranes are then incubated with secondary antibody (1:5,000) diluted in blocking buffer for 1–1.5 h at room temperature, shaking at 50 rpm. 12. After this, membranes are washed with TBST as in step 10. 13. Membranes are exposed to ECL reagent. For each of the two portions of this reagent, 1 mL is aliquoted per membrane, the two are then mixed and applied to the membrane, ensuring complete coverage. Incubation is for 1 min at room temperature. 14. The membrane is removed from the ECL and wrapped in Saran wrap or placed between two transparency sheets. Film is exposed to the membrane for varying times in a dark room. For phospho-Erk Thr202/Tyr204, very short exposure times are required (usually 1–10 s). Phospho-Raf-1 Ser338 requires slightly longer exposures (30 s to 3 min) (see Note 11).

3.4

Assessment of Membrane Translocation of Raf-1

1. After stretch, cells are harvested in hypotonic lysis buffer as for Western blotting. One stretch plate per condition is used.

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Cells are then homogenized by passing 12 times through a 30½-gauge needle attached to a 1-mL syringe. Tubes are kept on ice during homogenization. The lysate is then subjected to two freeze–thaw cycles to ensure complete lysis. Lysates are frozen by brief immersion of tubes into liquid nitrogen. They are thawed by placing into a cold-water bath. The nucleus is removed by centrifugation at 500×g for 5 min at 4°C. The supernatant is then transferred to thick-wall polycarbonate Beckman ultracentrifuge tubes precooled on ice and ultracentrifuged at 100,000×g for 60 min at 4°C. For a Beckman swinging bucket rotor SW 55Ti, this corresponds to 33,000 rpm. The resultant pellet is small but visible at the bottom center of the tube. The supernatant is removed and if desired can be run as the cytosolic fraction. The pellet (membrane fraction) is gently washed three times with 0.5-mL cold PBS and resuspended in regular lysis buffer (40–60 µL per pellet). Resuspension requires repetitive pipetting. If a sonicator is available, brief (2 s) sonication is very effective in pellet resuspension. Protein concentration is obtained and equal amounts of protein are separated by 10% SDS-PAGE. Phospho-Raf-1 Ser338 is detected using the mouse antibody in section 3.3.

Raf-1 Activity Assay

After serum deprivation for 24 h, MC are stretched for 5 min for maximal Raf-1 activation (Fig. 15.1b). Three wells per condition are required. 2. After washing three times with cold PBS, cells are lysed with lysis buffer, with 40–50 µL used per well, and incubated on ice for 10 min. 3. Lysates are scraped with a cell lifter and placed in 1.5-mL tubes precooled on ice. Pipetting up and down with a 200-µL tip assists in cell disruption. 4. Lysates are cleared by centrifugation for 10 min at 4°C and the supernatants are transferred to new precooled tubes. 5. Protein concentration is obtained as in Sect. 3.3 and 200 µg per sample aliquoted to a new tube. Volume is equalized with lysis buffer (see Note 12). 6. Anti-Raf-1 rabbit antibody (2 µg) is added to each sample and tubes are placed in a rotator and incubated at 4°C overnight. 7. The next morning, 25 µL of recombinant protein G agarose is added and samples incubated at 4°C for 1.5 h with continued rotation (see Note 13). 8. Protein G agarose is then pelleted by centrifugation for 1 min at 1,300×g, 4°C and the supernatant is removed. This is then washed three times with 0.5 mL lysis buffer, followed by two washes with kinase buffer. To wash, buffer is added and the tubes are inverted multiple times by hand to allow complete resuspension of the pellet. At this time, the water bath or thermomixer is warmed to 30°C (see below). 9. Kinase mix is prepared for the number of samples to be assayed, adding one extra sample to account for small pipetting errors. Thus, for four samples,

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enough kinase mix is prepared for five samples. After carefully pipetting off any excess kinase buffer from the beads, 20 µL of the kinase mix is added to each sample by placing onto the beads and gently tapping the bottom of the tube to resuspend the beads. 10. The kinase reaction is carried out at 30°C for 30 min, either in a thermomixer (at 300 rpm) or a water bath. If a bath is being used, the bottom of each tube is gently tapped every 5–10 min to mix. 11. The reaction is stopped by placing the tubes on ice, then adding 25 µL of 2× sample buffer and incubating at 100°C for 5 min. The samples are vortexed immediately after boiling for 10 s at maximum speed and then centrifuged at 16,200×g for 1 min. The supernatant is resolved on 10% SDS-PAGE. 12. Membranes are probed with mouse anti-phospho Erk (Thr202/Tyr204). Without stripping, they are then reprobed with rabbit anti-Raf-1 to ensure its equal immunoprecipitation (see Note 14).

3.6

Assay of RhoA Activation

1. Cells are stretched after serum deprivation for 24 h. Maximal early RhoA activity in MC occurs at 1 min of stretch. One plate per condition is required to obtain enough protein for the activity assay (see Note 15). 2. After stretch, cells are immediately washed three times with ice-cold PBS. Lysis buffer (40 µL) is added to each well and the plate incubated on ice for 5 min, but not longer (see Note 16). 3. During incubation, glutathione–agarose-bound GST-RBD beads are removed from -70°C and allowed to thaw. After thawing for several minutes at room temperature, they are kept on ice. 4. Cells are scraped from the well and placed into prechilled tubes on ice. Lysate is pipetted up and down several times to further aid in lysis. 5. Samples are centrifuged at 16,200×g, 4°C for 2 min to pellet cell debris, and the lysates then transferred to prechilled tubes. GST-RBD slurry is mixed well by gently flicking the tube, and an aliquot of 30 µg is added with a yellow tip that has been cut about 3–5 mm from its end to allow efficient uptake of the beads (see Note 17). 6. The sample with GST-RBD is rotated at 4°C for 45 min to immunoprecipitate GTP-bound RhoA. Incubation periods of up to 1.5 h have been used. 7. Beads are then collected by centrifugation at 16,200×g for 30 s at 4°C. An aliquot (50 µL) of the supernatant is set aside to be run for total cellular RhoA (see Note 18). Beads are then washed three times in 0.5 mL wash buffer. 8. After the final wash, as much supernatant as possible is removed, and 30 µL of 2× sample buffer is added to each tube on ice. Samples are boiled for 5 min and immediately after boiling, they are vortexed for 10 s at top speed, then centrifuged for 1 min at 16,200×g. For samples set aside as total RhoA, 5× reducing sample buffer is added followed by boiling for 5 min.

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9. Supernatant is run on a 12.5% or 15% SDS gel. Equal aliquots of supernatant samples are run for total RhoA (40–60 µL of final boiled sample). Mouse antiRhoA antibody is used for Western blotting. Signal is visualized with SuperSignal ECL (see Note 19).

3.7

Confocal Immunofluorescence for Raf-1 Membrane Localization

1. After stretch, MC are washed three times with PBS warmed to 37°C. Washes are carried out as for protein harvesting. 2. Cells are fixed for 15 min at room temperature. 3. After three further washes with PBS, cells are permeabilized for 10 min at room temperature. 4. Nonspecific binding is blocked by incubation with blocking buffer for 45 min at room temperature. 5. Blocking buffer is removed by suctioning. Primary antibody is then applied, diluted in blocking buffer. For each well, 800–1,000 µL sufficiently covers the surface area. Dilutions are 1:25 for mouse anti-phospho-Raf-1 Ser338 and 1:50 for rabbit anti-Raf-1. Lids are replaced on the plates, and these are sealed with Parafilm. Plates are left at 4°C overnight on a shaker at 55 rpm. 6. The next day, antibodies are removed and cells are washed three times with PBS (2 min each on a shaker at 150 rpm) (see Note 20). 7. Cells are then incubated with fluorochrome-conjugated secondary antibody diluted at 1:100 in PBS for 45 min at room temperature on a shaker at 55 rpm. This should be carried out in the dark. This is followed by three further washes with PBS, as above. 8. If the microscope to be used for imaging is compatible with the stretch plates, the remaining PBS is removed by suctioning and the wells allowed to become semi-dry. A drop of mounting medium with DAPI (to visualize cell nuclei) is placed onto each well and covered with a 22×22-mm coverslip. The edges of the coverslip are sealed with clear nail polish. The wells can then be imaged. Alternatively, the wells can be removed from the plate. Using a scalpel, carefully cut around the inside edges of the well, which is then removed with forceps and placed onto a slide with the bottom of the well adhering to the slide. A drop of mounting medium placed onto the slide before adding the well will provide enough adherence. Another drop of mounting medium is placed onto the cells and a 22×22-mm coverslip added. This is sealed with clear nail polish and allowed to dry for 20–30 min (see Note 21). 9. Samples can then be imaged or stored at 4°C in the dark and visualized within 1 week (Fig. 15.2).

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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 (see Color Plate 8)

3.8

Confocal Immunofluorescence for F- and G-Actin

1. After steps 1 to 3 in Sect. 3.7, cells are incubated in blocking solution for 30 min at room temperature (see Note 22). 2. Cells are then incubated with probes for F-actin and G-actin for 40 min at room temperature in the dark (no rotation is required). For F-actin, 20 µL of rhodamine

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phalloidin (equivalent to 4 U) is used in a total of 800 µL PBS. The probe for Gactin is added to this at 1.5 µL (final concentration ~9 µg/mL) (see Note 23). Cells are then washed three times with PBS, with the plate kept in the dark during washes and all subsequent steps. Coverslips are added as in step 8 in Sect. 3.7, and the samples are imaged within 1 week, storing at 4°C in the dark. The absorption and emission spectra are as follows: for rhodamine phalloidin, 554 nm/573 nm; and for DNase I–Alexa 488, 495 nm/519 nm. For quantification of F:G actin ratio, which provides an estimate of F-actin turnover (assembly/disassembly), the slides are sampled and the optimal exposure settings for capturing images for each fluorochrome are chosen. All of the wells are then imaged using the same settings. Once attained, the fluorescence in the images (as tiff files) is quantified as pixel intensity using software such as Scion Image. The ratio of intensity between Factin and G-actin is calculated. An increase in ratio represents increased F-actin formation, as is found in stretched MC (10).

3.9

Assessment of Fibronectin by ELISA

1. Cells are serum deprived in 0.5% for 24 h, then stretched for 24 h; 1.5 mL of medium per well is used. After stretch, medium is harvested into 1.5-mL tubes (see Note 24). 2. Cellular debris is removed by centrifugation at 1,300×g, 4°C for 5 min. Supernatant is transferred to a new tube and the pellet is discarded. At this point, medium can be stored at -70°C for later use. 3. The wells of a 96-well flat-bottomed plate are coated with medium diluted in ECB to a total of 100 µL per well, reserving 24 wells for the standard (see below). For example, for a 1:6 dilution, which works well for fibronectin; 83.3 µL of ECB; and 16.7 µL of medium are used. ECB is dispensed using a multichannel pipetter. Medium is then added to each well individually with an unused tip and mixed with ECB by pipetting up and down. Each sample is plated in triplicate. The outer wells of the plate are not used (see Note 25). 4. A fibronectin standard is prepared by making serial dilutions of fibronectin in ECB as follows: 1025, 256, 64, 16, 4, and 1 ng/100 µL. Each dilution is plated in duplicate (see Note 26). 5. The plate is covered tightly with Saran wrap or a 96-well plate sealer (e.g., Fisher), then placed at 4°C overnight without shaking. 6. The next day, the cover is removed and the wells washed four times with wash buffer. To wash, buffer is added to each well with a squirt bottle, the plate is shaken by hand, then turned upside down over a sink to discard the contents. It is patted dry on a paper towel, particularly after the last wash. 7. Blocking buffer (200 µL) is added to each well and the plate incubated for 1 h at 37°C. 8. Each well is washed three times with wash buffer.

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9. Primary antibody in dilution buffer at 100 µL per well is added and incubated for 3 h at room temperature, shaking at 400–500 rpm. Dilutions that work well are 1:5,000 for the mouse antibody and 1:3,500 for the rabbit antibody. 10. The plate is washed three times with wash buffer. 11. Secondary antibody is added at 1:20,000 in dilution buffer at 200 µL per well and the plate incubated for 2 h at room temperature, shaking at 400–500 rpm. 12. The plate is washed three times with wash buffer. 13. To each well, 100 µL of pNPP is added and the reaction is allowed to develop until a yellow color appears (2–4 h). The plate is read at 405 nm. The curve generated from the fibronectin standard can be used to convert readings to approximate concentrations (see Note 27).

4

Notes

1. For all buffers, leupeptin, aprotinin, and PMSF are added when cells are harvested, with PMSF added immediately prior to use since its half-life is short, at approximately 30 min. 2. Loading stations are optional, with 25 mm providing the greatest strain range. These are covered in lubricant (provided with the stations) and placed beneath each well of the culture plate within the baseplate. They provide uniform stretch across the membrane overlying the area of the loading station. Note that the outer portion of the well will still undergo greater stretch than that programmed. 3. Various coatings are available. Collagen I is found primarily in pathologic settings, and collagen IV is a component of the normal basement membrane that may be produced in excess. For the signaling pathways discussed here, similar responses are seen when plates coated with either collagen I or IV are used. 4. The serum component should be specific to the species in which the secondary antibody was raised. 5. DNase I is highly sensitive to physical denaturation by shaking. It should be mixed by gentle inversion. Phalloidin and DNase I bound to other fluorochromes are also available. 6. A stock of 20% BSA can be made in water and kept at 4°C for several weeks. 7. To wash after stretch, tip the plate and gently pour cold PBS into the side of each well, allow the PBS to cover the well, then discard. After the last wash, suction out the remaining PBS, keeping the plate on ice. Ensure that all PBS is removed, otherwise the sample may become too diluted to load enough protein for assessment by Western blotting. 8. Fairly small amounts of lysis buffer are used in order to not dilute the samples. Lysis buffer is added immediately after suctioning out residual PBS. If the wells dry prior to addition of lysis buffer, the buffer will not spread out well. If this occurs, buffer will need to be redistributed by pipetting onto the dry areas. 9. Antibodies can generally be reused up to 10–15 times. They are stored at –20°C in 15-mL tubes with the number of uses tracked on the side of the tube. They are thawed at room temperature or in cold water. With greater reuse, longer film exposure times might be needed for signal detection. 10. For phospho-Erk Thr202/Tyr204, incubation with primary antibody for 1 h at room temperature is sufficient, and helps to preserve antibody longevity. 11. Film is exposed for the shortest time that will allow visualization of the signal; overexposure will saturate the signal and minimize differences between conditions. Western blotting for total Erk and Raf-1 will provide a loading control. Membranes can be reprobed for Raf-1 after stripping, but new samples need to be run for total Erk since the phospho-Erk antibody does not strip well.

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12. As little as 70 µg has been found sufficient to obtain stretch-induced Raf-1 activation. The final volume should not be less than 200 µL (equalized with lysis buffer) in order to ensure adequate mixing with immunoprecipitating antibody. 13. Ensure that the beads are completely resuspended. Retrieval of the beads is best done with a 200-µL pipette tip cut at its end. 14. The mouse anti-phospho-Erk antibody is required. If the rabbit antibody is used, the immunoglobulin bands from the immunoprecipitating antibody will obscure the phospho-Erk signal. Alternatively, the mouse antibody to Raf-1 from Santa Cruz Biotechnology can be used for immunoprecipitation, with the rabbit anti-phospho-Erk used for Western blotting. The mouse anti-Raf-1 antibody, however, is not good for subsequent detection of immunoprecipitated Raf-1 by Western; the rabbit antibody should be used. 15. We use 0% FBS for serum deprivation since serum has been shown to activate RhoA (20). Time is of significant importance in this procedure since RhoA possesses intrinsic hydrolytic activity. All steps are thus shorter than those in Sects. 3.3–3.5 and should be carried out as quickly as possible. For best results, one plate should be processed at a time to step 5; lysis buffer can be aliquoted and PMSF added immediately prior to harvesting each plate. 16. Ice-cold conditions are important throughout the procedure in order to reduce the intrinsic hydrolytic activity of RhoA. All tubes need to be precooled on ice. 17. Trials can be carried out with the quantity of GST-RBD beads used. We routinely use as little as 15–20 µg. 18. This can be done since only approximately 0.5–5% of total cellular RhoA is activated at a given time (21). 19. ECL as used in Sect. 3.3 may be adequate, although longer exposure times (up to overnight exposure) may be required. Use of SuperSignal ECL reduces exposure time to seconds to minutes. If there is difficulty in performing this assay, trials using a positive control can be used. After serum deprivation, 5 min of exposure to serum (i.e., full medium) works well as a positive control. 20. Antibodies can be kept at -20°C and reused up to five times, with the number of reuses marked on the tubes. 21. A single well StageFlexerTM is available to use for imaging in real time. This is a single 35-mm well form of the deformable membrane that can be placed on a microscope stage. 22. Methanol should not be used to fix cells as this disrupts the cytoskeletal structure. 23. The phalloidin probe is excellent and readily visualized, while the G-actin signal is much weaker. 24. Medium used to serum deprive and stretch cells should be 0.5% FBS since some serum is required for optimal recovery of secreted matrix proteins (4). 25. For other matrix proteins, the optimal dilution needs to first be established. A good starting point is trying dilutions of 1:2 to 1:10 in triplicate in conjunction with a standard. Readings should fall within the standard curve. 26. Do not store fibronectin diluted in ECB; prepare fresh for each ELISA. 27. Equal cell numbers are plated across conditions, and cell viability with this protocol is good. However, results can also be normalized for protein content per well if desired. Cells would be lysed at the time of harvesting medium and protein concentration measured as in Sect. 3.3. Acknowledgments This work was supported by funding from the Kidney Foundation of Canada and the Canadian Institutes of Health Research.

References 1. Dworkin LD, Feiner HD (1986). Glomerular injury in uninephrectomized spontaneously hypertensive rats. A consequence of glomerular capillary hypertension. J Clin Invest. 77:797–809.

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2. Meyer TW, Anderson S, Rennke HG, Brenner BM (1987). Reversing glomerular hypertension stabilizes established glomerular injury. Kidney Int. 31:752–759. 3. Riser BL, Cortes P, Yee J (2000). Modelling the effects of vascular stress in mesangial cells. Curr Opin Nephrol Hypertens. 9:43–47. 4. Riser BL, Cortes P, Zhao X, Bernstein J, Dumler F, Narins RG (1992). Intraglomerular pressure and mesangial stretching stimulate extracellular matrix formation in the rat. J Clin Invest. 90:1932–1943. 5. Yasuda T, Kondo S, Homma T, Harris RC (1996). Regulation of extracellular matrix by mechanical stress in rat glomerular mesangial cells. J Clin Invest. 98:1991–2000. 6. Hamaguchi A, Kim S, Izumi Y, Iwao H (2000). Chronic activation of glomerular mitogenactivated protein kinases in Dahl salt-sensitive rats. J Am Soc Nephrol. 11:39–46. 7. Ingram AJ, James L, Cai L, Thai K, Ly H, Scholey JW (2000). NO inhibits stretch-induced MAPK activity by cytoskeletal disruption. J Biol Chem. 275:40301–40306. 8. Ishida T, Haneda M, Maeda S, Koya D, Kikkawa R (1999). Stretch-induced overproduction of fibronectin in mesangial cells is mediated by the activation of mitogen-activated protein kinase. Diabetes. 48:595–602. 9. Harris RC, Haralson MA, Badr KF (1992). Continuous stretch-relaxation in culture alters rat mesangial cell morphology, growth characteristics, and metabolic activity. Lab Invest. 66:548–554. 10. Krepinsky JC, Ingram AJ, Tang D, Wu D, Liu L, Scholey JW (2003). Nitric oxide inhibits stretch-induced MAPK activation in mesangial cells through RhoA inactivation. J Am Soc Nephrol. 14:2790–2800. 11. Krepinsky JC, Li Y, Tang D, Liu L, Scholey J, Ingram AJ (2005). Stretch-induced Raf-1 activation in mesangial cells requires actin cytoskeletal integrity. Cell Signal. 17:311–320. 12. Amano M, Chihara K, Kimura K, et al.(1997). Formation of actin stress fibers and focal adhesions enhanced by Rho-kinase. Science. 275:1308–1311. 13. Kolch W (2000). Meaningful relationships: the regulation of the Ras/Raf/MEK/ERK pathway by protein interactions. Biochem J. 351 Pt 2:289–305. 14. Pearson G, Robinson F, Beers GT, et al.(2001). Mitogen-activated protein (MAP) kinase pathways: regulation and physiological functions. Endocr Rev. 22:153–183. 15. Mason CS, Springer CJ, Cooper RG, Superti-Furga G, Marshall CJ, Marais R (1999). Serine and tyrosine phosphorylations cooperate in Raf-1, but not B-Raf activation. EMBO J. 18:2137–2148. 16. Dhillon AS, Kolch W (2002). Untying the regulation of the Raf-1 kinase. Arch Biochem Biophys. 404:3–9. 17. Carey KD, Watson RT, Pessin JE, Stork PJ (2003). The requirement of specific membrane domains for raf-1 phosphorylation and activation. J Biol Chem. 278:3185–3196. 18. Bishop AL, Hall A (2000). Rho GTPases and their effector proteins. Biochem J. 348 Pt 2:241–255. 19. Burridge K, Wennerberg K (2004). Rho and Rac take center stage. Cell. 116:167–179. 20. Hill CS, Wynne J, Treisman R (1995). The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell. 81:1159–1170. 21. Ren XD, Kiosses WB, Schwartz MA (1999). Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J. 18:578–585.

Chapter 16

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

Abstract Fibrosis and sclerosis are widely recognized as hallmarks of progressive renal disease and are caused by the excessive accumulation of connective tissue, mostly collagen. The detection of collagen content, concentration (collagen content/dry weight tissue), and sub-types from kidney tissues is therefore an important part of determining the extent of renal fibrosis in ageing and diseased states. This chapter describes a colorimetric-based hydroxyproline assay used to estimate total collagen content and concentration. Based on the method of Bergman and Loxley (8), this spectrophotometric technique estimates total collagen by measuring the hydroxyproline content of tissue. The assay relies on the fact that the collagen triple helix is one of the few proteins that contain the amino acid hydroxyproline. The second part of this chapter describes the use of sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE) to isolate, detect and quantify changes in the soluble and insoluble interstitial collagen sub-types. This technique complements the hydroxyproline assay by providing a means of identifying which interstitial collagens are altered in renal disease. Keywords Hydroxyproline, Kidney collagen content, Kidney collagen concentration, Interstitial collagen subtypes

1

Introduction

Fibrosis and sclerosis is the final common pathway in all forms of progressive renal disease, regardless of etiology (1, 2). The progression of renal fibrosis/ sclerosis is the result of an excess accumulation of connective tissue, primarily collagen, which is over-expressed by renal vascular smooth muscle cells, mesangial cells, and interstitial myofibroblasts in response to repeated pathological stimuli and inflammation in the kidney (1, 2). Various collagen sub-types accumulate in this process, including both so-called interstitial (collagen I, III, and V) and basement membrane (collagen IV) proteins (3). Measuring changes

From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_16, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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in collagen content, concentration, and composition is therefore fundamental to our understanding of renal fibrosis. Several techniques have been established for this purpose. While morphometric techniques in particular are widely used, they are limited by the fact that they often only measure changes relative to normal/control tissue, and therefore do not measure absolute quantities. Such methodologies are therefore often unable to account for hypertrophy and atrophy of organs. The spectrophotometric-based hydroxyproline assay is one of the few assays that allows for the actual quantitation of collagen content and concentration in several organs, including the kidney. The assay was first described in 1950 (4), based on the fact that all collagens contain globular domains and share the common structural motif of triple helical segments. This triple helical structure is composed of three α- (polypeptide) chains, which range from 10 to 150 kDa per chain. Each chain consists of a repeating triplet amino acid sequence (Gly-X-Y)n, where X and Y can be any amino acid, but are often proline and hydroxyproline, respectively (5). Collagen is one of the few proteins containing the amino acid hydroxyproline. Thus, based on the absolute quantitation of hydroxyproline (which represents a fixed percentage of the amino acid composition of collagen in most mammalian organs (6, 7)), the amount of collagen content and concentration in tissues can also be derived. Since its original description (4), this method has been adapted and/or modified by several investigators to overcome a number of difficulties including reproducibility in optimal values between sample replicates, differences between standard curve values prepared on separate occasions, and interference by other components (particularly when the hydroxyproline concentration is low relative to other amino acids). While some of these issues are yet to be fully resolved, the method of Bergman and Loxley (8) has been established as one of the better assays for hydroxyproline determination, yielding optimal hydroxyproline values with reproducible standard curves. In this chapter, a 1:10 scaled-down version of the original Bergman and Loxley method (8) is described. This method has been used successfully in our laboratory to measure collagen content and concentration in the normal, ageing, and diseased kidney (3, 9, 10). Additionally, this chapter describes the extraction, detection, and quantitation of interstitial collagen sub-types (types I, III, and IV) by sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE), based on the original procedure described by Sykes and colleagues (11). Again this can been applied to kidney tissue from experimental (3, 9, 10) or clinical models. The SDS-PAGE method allows for both quantitation of the relative changes in interstitial collagen sub-types, and the amounts of soluble and insoluble (cross-linked) interstitial collagens in several organs, including the kidney. This technique is particularly useful for studying temporal changes in individual interstitial collagen subtypes.

16 Collagen Content, Concentration, and Sub-types in Kidney Tissue

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Materials Hydroxyproline Determination of Collagen Content and Concentration Liquid nitrogen. Hydrochloric acid 6 M and 0.1 M. Sodium hydroxide. Chloramine T (Sigma, St. Louis, MO, USA) is dissolved in distilled water to give a final concentration of 7% (w/v) and can be stored for up to 2 weeks at 4°C, in a dark bottle. Acetate/citrate buffer, pH 6.0 (per 100 mL): dissolve 5.7 g sodium acetate.3H2O (or 3.44 g sodium acetate anhydrous); 3.75 g tri-sodium citrate 2H2O; and 0.55 g citric acid in 38.5 mL of isopropyl alcohol and distilled water. The buffer can be stored at 4°C up to 6 months. Oxidation buffer is prepared (fresh before each assay) by mixing one part 7% chloramine T with four parts acetate citrate buffer and should be protected from light. Ehrlich’s reagent is prepared by dissolving 2.0 g para-dimethylaminobenzaldehyde (DMAB; Sigma) in 3.0 mL of 60% (v/v) perchloric acid and can be stored for up to 2 weeks at 4°C in a dark bottle. For assaying of larger sample numbers, multiple amounts of DMAB should be dissolved in the respective multiple volumes of perchloric acid. Analytical isopropanol reagent is prepared (fresh before each assay) by mixing 3 parts DMAB to 13 parts isopropanol. Hydroxyproline standard is prepared by dissolving cis 4-hydroxy-l-proline (Sigma) in 0.1 M hydrochloric acid (HCl) to a concentration of 1 mg/mL (w/v), should be protected from light, and can be stored at –20°C. Kimax™ screw-capped glass tubes (Kimble/Kontes, Vineland, NJ, USA). Disposable cuvettes (Lake Charles Manufacturing, Lake Charles, LA, USA). Lyophilizer. Heating block. Spectrophotometer. Microsoft Excel™ (or equivalent software program for plotting regression line).

SDS-PAGE Analysis of Interstitial Collagen Sub-types

1. Biorad Protean II™ xi gel system (Biorad, Richmond, CA, USA). 2. Neutral salt extraction buffer containing: 0.05 M Tris-HCl acid pH 7.5, 0.15 M sodium chloride, 10 mM protease inhibitors N-ethylmaleimide (Sigma), 10 mM

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3. 4. 5. 6. 7. 8. 9.

10. 11.

12. 13. 14. 15. 16.

17. 18.

3 3.1

C.S. Samuel

phenylmethylsulfonyl fluoride (Sigma), 1 mM benzamidine hydrochloride (Sigma), and 10 mM EDTA (Sigma). The protease inhibitors are added to prevent pro-collagen degradation during isolation. 0.5 M acetic acid diluted with distilled water. Pepsin (Boehringer Mannheim, Mannheim, Germany). Cyanogen bromide. Ammonium bicarbonate. Formic acid. Nitrogen (BOC Gases, Prahran, Australia). Separating gel I: 5% bis-acrylamide gel containing 0.375 M Tris-HCl pH 8.8, 2 M urea, and 10% SDS (Biorad). For large gels (using the Biorad Protean II xi gel system) add 150 µL of 10% (w/v) ammonium persulfate (APS; ICN Biomedicals Inc., Aurora, OH, USA) to aqueous gel contents; de-gas contents for 5 min by air vacuum; and then add 20–40 µL of TEMED (Sigma) before allowing gel to set/cross-link for 45–60 min (at room temperature). Separating gel II: 12.5% bis-acrylamide gel containing 0.375 MTris-HCl pH 8.8, 2 M urea, and 10% SDS. APS and TEMED can be added as described in Sect. 2.2.9. Stacking gel I: 3.5% bis-acrylamide gel containing 0.125 M Tris-HCl pH 6.8, 2 M urea, and 10% SDS. For large gels (using the Biorad Protean II gel system) add 50 µL of 10% (w/v) APS to aqueous gel contents; de-gas contents for 5 min by air vacuum; and then add 10–20 µL of TEMED before allowing gel to set/cross-link for 35–45 min (at room temperature). Stacking gel II: 4.5% bis-acrylamide gel containing 0.125 M Tris-HCl pH 6.8, 2 M urea and 10% SDS. APS and TEMED can be added as described in Sect. 2.2.11. Sample-loading buffer: 20% (w/v) sucrose/glycerol containing 0.05 M Tris-HCL pH 6.8, 2 M urea, 0.1% (v/v) SDS, and 0.1% (w/v) bromophenol blue (Biorad). Running buffer: 25 mM Tris-base, containing 0.192 M glycine and 0.35% (w/v) SDS. 10% (v/v) mercaptoethanol diluted in running buffer. Coomassie blue-R250: prepare a 0.1% (w/v) solution of Coomassie blue-R250 (Biorad) by dissolving in 45% methanol and 7% acetic acid for at least 60 min; then filter the solution through two layers of Whatmann 3m filter paper and chill at 4°C overnight. De-stain containing 20–30% methanol and 7% acetic acid. Densitometer with quantitative software (Biorad GS710 Calibrated Imaging Densitometer; Biorad).

Methods Preparation of Samples for Hydroxyproline Assay

1. Isolated kidney tissues should be weighed and can then be either snap-frozen in liquid nitrogen or stored immediately in dry ice. For analysis of more specific regions within each kidney tissue (i.e. cortex, medulla, renal pyramids, renal

16 Collagen Content, Concentration, and Sub-types in Kidney Tissue

2. 3. 4.

5.

6.

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papilla, etc.), these regions should immediately be dissected out in cold phosphate-buffered saline (PBS) and weighed before being frozen down. Each whole or regional kidney tissue should then be freeze-dried (lyophilized) down to dry weight and the dry weight tissue accurately measured. The dried tissues should then be re-hydrated in acetate/citrate buffer for 24 h at 4°C (see Note 1). The rehydrated tissue should then be transferred to Kimax™ screw-capped glass tubes (Kimble/Kontes) and submerged in 0.5–1.0 mL, 6 M HCl before being hydrolyzed at 110°C (in a heating block) for at least 18–24 h (see Note 2). The following day, the hydrolyzates should be cooled at 4°C, before being evaporated to dryness in the presence of a strong base (i.e. sodium hydroxide; to neutralize the acid) in a lyophilizer (overnight). The dried residues should finally be dissolved in 0.1 M HCl (as was performed for the hydroxyproline standard; approximately 1 mL per 100 mg wet weight tissue).

3.2

Hydroxyproline Determination (See Notes 3 and 4)

1. All assays should be conducted in Kimax™ screw-capped glass tubes, with the capacity to hold ≥5 mL of assay material. 2. Assay each sample in duplicate or triplicate by adding 10 µL of kidney tissue hydrolyzate (in 0.1 M HCl) to 90 µL of distilled water (in a separate Kimax™ glass tube per sample and replicate) (see Note 5). 3. In separate tubes, also establish a standard curve for each experiment by adding 0 (blank), 2, 4, 6, 8, and 10 µL of the 1 mg/mL hydroxyproline standard (in 0.1 M HCl) in separate Kimax™ glass tubes and adjusting the volume of each tube to 100 µL with distilled water. 4. To each of the standard curve and sample tubes (100 µL), add 200 µL of isopropanol, followed by 100 µL of the oxidation buffer (to give a volume of 400 µL). Immediately vortex all samples and allow them to stand at room temperature for 4 min (± 30 s). 5. Add 1.3 mL of the analytical isopropanol reagent to each tube and vortex each sample again (to give a volume of 1.7 mL), before tightly capping each tube. All samples should then be placed for 25 min in a shaking water bath or oven pre-set to 60°C (see Note 6). 6. The samples should then be cooled at 4°C for approximately 5–10 min (see Note 7). 7. Isopropanol (3.3 mL) should then be added to the cooled samples (to give a final volume of 5.0 mL), before the samples are thoroughly vortexed and decanted into disposable cuvettes (Lake Charles Manufacturing). 8. The absorbance of each standard curve sample/sample of interest should then be obtained at 558 nm in a spectrophotometer, using the “blank” to calibrate each assay. 9. A standard curve can then be generated using Ab558 nm (y-axis) vs micrograms of hydroxyproline added (x-axis) (Fig. 16.1). A line of best fit, equation (i.e. micrograms of hydroxyproline = Ab558 nm/arbitrary constant value), and R2-value can then be obtained (using Microsoft Excel™ software).

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Absorbance 558nm

2.5 2 1.5 1 0.5 0 0

2

4

6

8

10

12

Hydroxyproline (µg) Fig. 16.1 A typical standard curve for the hydroxyproline assay, which plots Ab558 nm (y-axis) versus micrograms of hydroxyproline (cis 4-hydroxy-l-proline) added (x-axis). A line of best fit, equation y = arbitrary constant/x, and R2 value can be obtained using Microsoft Excel software, as shown. Using the provided equation, the micrograms of hydroxyproline in each sample of interest can then be determined by dividing the corresponding Ab558 nm value by the arbitrary constant value

10. Based on the Ab558 nm readings, the generated equation (for each assay) should then be used to determine the amount of hydroxyproline (between 0 and 10 µg) in each sample (from 10 µL of starting material). The hydroxyproline values in duplicates or triplicates of the same sample should then be averaged out. 11. The total hydroxyproline content in each sample can be then extrapolated by multiplying the amount hydroxyproline (from the 10 µL of starting material) by the total volume of 0.1 M HCl added to the sample hydrolyzates (see Sect. 3.1.6)/10 µL. For example, if the sample hydrolyzates were dissolved in 1.0 mL (1,000 µL) of 0.1 M HCl, then the total amount of hydroxyproline in each sample is calculated by multiplying the amount of hydroxyproline from the 10 µL of assay material by 100 (Fig. 16.2a).

3.3

Determination of Collagen Content and Concentration (See Note 8)

1. The total (whole/regional) kidney collagen content can then be extrapolated by multiplying amount of total hydroxyproline content in each sample by a factor of 6.94, based on the fact that hydroxyproline represents 14.4% of the amino acid composition of collagen in most mammalian tissues (6, 7) (Fig. 16.2b).

Total hydroxyproline content (µg)/ kidney tissue

16 Collagen Content, Concentration, and Sub-types in Kidney Tissue

*** ###

200

*

150

100

50

0

Day 0

a

OB

Day 3 UO

Total collagen content (µg)/ kidney tissue

* 900

600

300

0

b Total collagen concentration (%)/ kidney tissue

Day 10 OB UO

*** ###

1200

c

229

Day 0

Day 3 OB UO

Day 10 OB UO

*** ###

4

3

*

2

1

0

Day 0

Day 3 OB UO

Day 10 OB UO

Fig. 16.2 Total hydroxyproline content (a), the corresponding collagen content (b), and collagen concentration (c) from the kidneys of control wild-type (C57B6Jx129SV) mice (at day 0); and from the obstructed (OB) and unobstructed (UO)/contralateral kidneys of mice 3 days and 10 days after unilateral ureteric obstruction. *p< 0.05; ***p< 0.001 versus values from day 0 kidneys; ###p<0.01 versus values from day 10 UO kidneys

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2. Collagen concentration (in the whole/regional kidney tissue), expressed as a percentage, can then be calculated by dividing the total collagen content of tissues (see Sect. 3.3.1) by the dry weight tissue (Fig. 16.2c).

3.4

Preparation of Samples for SDS-PAGE

1. Isolated whole/regional kidney tissues should be treated as described in Sect. 3.1.1 (see Note 9). 2. Kidney tissues should be powdered/chopped finely in the presence of liquid nitrogen and then placed into the neutral salt buffer (0.5–1.0 mL buffer/powdered kidney tissue, which extracts the newly synthesized soluble collagen) for 24 h at 4°C (with gentle agitation) (see Note 10). 3. Samples should then be centrifuged at ∼12,000×g for 45 min at 4°C (in a Beckman or Sorvall ultracentrifuge; or in an Eppendorf centrifuge with 2-mL Eppendorf tubes). The neutral salt supernatants can be frozen in dry ice and stored at –80°C or can be lyophilized down and 1) hydrolyzed in 6 M HCl (see Sect. 3.1.4) before being re-suspended in 50–100 µL of 0.1 M HCl (see Sect. 3.1.6) for hydroxyproline determination; or 2) re-suspended in Tris/HCl, pH 6.8 and sample loading buffer for SDS-PAGE analysis. 4. The neutral salt pellets can then be further extracted with 0.5 M acetic acid (which extracts the newly cross-linked soluble collagen) for 24 h at 4°C (with gentle agitation). 5. Samples should again be centrifuged and the acetic acid supernatants treated as described in Sect. 3.4.3. 6. The acetic acid pellets should also be freeze-dried/lyophilized down to dry weight (overnight). 7. One milligram of dry-weight acetic acid pellet should then be digested with 1:10 enzyme:substrate pepsin (which extracts the mature cross-linked insoluble collagen). The pepsin solution should originally be prepared as a 1 mg/mL (w/v) solution in 0.5 M acetic acid before being allowed to dissolve at 4°C for 60 min with gentle agitation. Following this 60-min period, a 1:10 dilution of the 1 mg/mL pepsin stock should be made with 0.5 M acetic acid (providing a 0.1 mg/mL pepsin solution); and 1 mL of this 0.1 mg/mL pepsin stock solution should be used to digest the insoluble collagen from 1 mg of dry weight acetic acid pellet for 24 h at 4°C (with gentle agitation). If required, larger volumes of 0.1 mg/mL pepsin can be used to extract the mature insoluble collagen from larger amounts of the lyophilized acetic acid pellet, as long as the enzyme:substrate ratio remains the same (see Notes 11 and 12). 8. Samples should again be centrifuged and the pepsin-digested supernatants treated as described in Sect. 3.4.3. 9. The final pepsin insoluble pellet can be lyophilized and stored at –80°C (over long-term periods) or can be cleaved with cyanogen bromide (CNBr) using the methods of Scott and Veis (12 , 13), which digests the remaining insoluble

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10. 11. 12. 13.

14. 15.

16. 17.

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kidney collagen (which is neutral salt, acetic acid, and pepsin insoluble). The procedures detailed in Sect. 3.4.10–3.4.15 should be carried out in a fume cupboard. The lyophilized pepsin-digested pellets should be dissolved in 100 mM ammonium bicarbonate, pH 8.0 (50 µL) for > 30 min (to inactivate the pepsin). Bubble 98% formic acid with nitrogen (to stop any oxygen from entering the system; as oxygen will interfere with the CNBr reaction). Weigh CNBr in a pre-weighed plastic tube and cap tightly. Dissolve CNBr at 70 mg/mL in nitrogen-saturated 98% formic acid for 10 min. Smaller/larger amounts of CNBr and nitrogen-saturated 98% formic acid may be used as long as the ratio of CNBr:formic acid remains constant. Digest samples at about 1 mL of 70 mg/mL CNBr per 1 mg protein for 4 h with gentle agitation. This reaction can be terminated by drying samples down in a Speedvac centrifuge under vacuum. Alternatively, CNBr and formic acid can be removed under a stream of nitrogen. Wash CNBr-digested pellets with 0.5 M acetic acid and lyophilize to remove any residual CNBr. Re-suspend washed CNBr-digested pellets with Tris-HCl, pH 6.8 and sampleloading buffer for SDS-PAGE analysis, or hydrolyze the pellet with 6 M HCl (see Sect. 3.1.4) before re-suspending it in 100–200 µL of 0.1 M HCl (see Sect. 3.1.6) for hydroxyproline determination.

SDS-PAGE Analysis of Interstitial Collagen Sub-types

1. Prepare 1× running buffer and chill at 4°C overnight. 2. Set up a Biorad Protein II™ xi (large) gel system for SDS-PAGE analysis (as the larger gel system allows for better separation of the collagen chains) (see Note 13). 3. Prepare the 5% separating gel (for neutral salt, acetic acid, and pepsin-soluble collagens) or 12.5% separating gel (for CNBr-digested collagens). Mark large plates 3.5 cm from the top (or 1.0 cm from the bottom of the sample wells). Also mark the plates 2.2 cm and 8.0 cm from the top of the 5% separating gels or 11.0 cm from the top of the 12.5% separating gels. Separating gels can be prepared a day before the addition of stacking gels; but should be covered with 1:3 ratio of Tris-HCl, pH 8.8:running buffer (10 mL), once set, and stored at 4°C overnight. The following day, gels can be thawed to room temperature while the stacking gel is being prepared. 4. Prepare the 3.5% stacking gel (for neutral salt, acetic acid, and pepsin-soluble collagens) or 4.5% stacking gel (for CNBr-digested collagens) with 10- to 15well sample combs (Biorad), which can hold up to 150 µL (15-well) to 200 µL (10-well) of sample. Once the stacking gel is set and combs removed, the sample wells should be washed three to six times with 1× running buffer (to equilibrate the samples with the running buffer).

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5. Heat neutral salt-, acetic acid-, pepsin-, and CNBr-extracted samples in Tris/ sample loading buffer at 60–65°C for 10 min before loading (with equivalent amounts of total protein) onto gels. 6. Once the gels tanks are filled with the appropriate amount of running buffer, run gels at a constant 25 mA through the stacking gel and 40 mA through the separating gel. For CNBr gels, samples should be run until they reach the 11.0-cm mark from the top of the 12.5% separating gel. For neutral salt/acetic acid/ pepsin gels, samples should be run and treated as described in Sects. 3.5.8–3.5.10 (see Note 14). 7. For neutral salt/acetic acid/pepsin gels: once samples have reached the 2.2-cm mark from the top of the separating gel, the gel should be stopped and the running buffer from the top of the gel and sample wells removed. 8. Ten percent (v/v) β-mercaptoethanol (diluted in running buffer) should then be added to the top of the sample wells of the separating gel for 90–120 min in a fume cupboard. This step is completed to separate the type I collagen α(α1(I)]chains from the type III collagen α(α1(III)]-chains. Under non-reducing conditions, these chains would normally migrate to the same position, making it difficult to distinguish them apart; however under reducing conditions (with β-mercaptoethanol), the type III collagen trimers/higher polymers are cleaved and the released α1(III) monomers migrate slightly more slowly than the α1(I) chains (see Note 15). 9. After 90–120 min the β-mercaptoethanol should be removed from the sample wells and replaced with 1× running buffer, so that the samples can be run (at 40 mA) to the 8.0-cm mark from the top of the separating gel. 10. Once all gels have been completed, they should be removed from the gel apparatus and stained overnight (in chilled Coomassie blue-R250; ∼200 mL/large gel) at 4°C. 11. Gels can then be de-stained (at room temperature) until the background stain is removed and the type I, III, and V collagen chains are visible (Fig. 16.3) and then washed in distilled water (to stop the de-stain reaction) (see Notes 16, 17, and 18). 12. For quantitation of the interstitial collagen monomers, dimers, and trimers (Fig. 16.3), the individual collagen chains can be analyzed by densitometry (using a Biorad GS710 Calibrated Imaging Densitometer and appropriate quantitative software; which allows for the quantitation of collagen bands from wet gels). Alternatively, gels can be soaked in 3% (w/v) glycerol for 30–45 min before being dried in gel-drying film (Promega, Madison, WI, USA) and then quantitated by densitometry.

4

Notes

1. After whole/regional kidney tissues are lyophilized for measurement of dry weight and rehydrated, they may be de-fatted in a mixture containing 2:1 chloroform:methanol for 24 h at 4°C before being re-hydrated for 24–48 h (at 4°C) and then hydrolyzed in 6 M HCl (see Sect. 3.1.4). This step is more suited to organs (such as the skin) that have a high fat content.

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γ

β11 β12

α1(III) α1(V) α2(V) α2(I)

α1(I)

Fig. 16.3 The pepsin-digested (mature cross-linked) interstitial collagens from the obstructed kidney of a wild-type (C57B6J) mouse, 10 days after unilateral ureteric obstruction. Shown are the type I collagen monomers, identified by the α1(I) and α2(I) subunits/chains; type III collagen monomers, identified by the α1(III) chains; and type V collagen monomers, identified by the α1(V) and α2(V) subunits/chains. Also shown are type I collagen dimers, β11: dimers of two α1(I) subunits; β12: dimers of α1(I) and α2(I) monomers; and trimers (γ)

2. During the hydrolysis step with 6 M HCl, it should be ensured that the Kimax glass tubes fit into the heating block (with no gaps between the external surface of the glass tube and the heating block; as this will affect the rate of hydrolysis). 3. The amount of hydroxyproline in collagen may vary between species and organs studied and should be checked before any calculations on collagen content and concentration are performed (see refs. (6, 7) as a guide). 4. Since the measurement of collagen content and concentration is extrapolated from the absolute hydroxyproline values of kidney tissues, other measures of collagen content and/or subtypes (by Western blotting, immunostaining, or SDS-PAGE analysis of interstitial collagen chains) are also recommended to confirm the findings of the hydroxyproline assay. 5. Ten to 20 µL of sample (hydrolyzate) is recommended for this assay as using larger amounts (up to 100 µL) does not provide a linear outcome (author’s unpublished observations). 6. While optimal yields of hydroxyproline (color reaction) will be obtained when samples are heated at 60°C for 25 min, these optimal yields will be lost if samples are heated for longer periods. 7. Samples need to be cooled down to stop the color reaction.

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8. If whole kidney tissues are used for analysis, then either collagen content or collagen concentration can be used as appropriate measures and comparisons between treatment groups. However, if only regional kidney tissues are used, then only collagen concentration should be used for comparisons between groups, as this measure corrects for the dry weight of the particular region of tissue used. It should be noted that collagen concentration is best calculated from the dry weight tissue (which represents the extracellular matrix material, primarily collagen); whereas deriving collagen concentration from the wet weight tissue is not as accurate, due to the fact that the water content of tissues may vary between organs and diseased conditions. 9. As with Note 1, whole/regional kidney tissues may be de-fatted in a 2:1 mixture of chloroform:methanol for 24 h at 4°C before being subjected to extraction with the neutral salt buffer. 10. Kidney tissues can be diced finely/powdered with a mortar and pestle or tissue chopper. 11. The pepsin solution should always be prepared fresh and dissolved in 0.5 M acetic acid, 1 h before use. If required, a second extraction of the mature insoluble collagen chains by pepsin can be performed, before CNBr treatment of pepsin-digested pellets. 12. All extractions, particularly with pepsin, should be carried out with gentle agitation, as more vigorous agitation may lead to the autolytic cleavage of the pepsin itself and hence, decreased efficiency of the pepsin in digesting the mature insoluble collagens. 13. SDS-PAGE analysis of the interstitial collagen chains can be achieved with Biorad mini-gels, but will result in poorer separation between the individual collagen -chains (see Fig. 16.3). 14. Due to the longer time required to run these larger SDS-PAGE gels, the gel running buffer should be chilled at 4°C overnight before use and the gel tanks cooled in ice or connected to a gel cooling system or the gels themselves run at 4°C to ensure a linear migration of all collagen bands. Once gels are heated, the outer samples (that are closer to the electrodes) will migrate at a slower rate compared with samples loaded in the middle of each gel. 15. Differences in the migration rate of the type III collagen a1(III) chains, under reducing conditions with b-mercaptoethanol, may be observed between species, with greater differences, between the a1(III) and a1(I) chains, being observed in the rat, mouse, calf, and rabbit than in humans (11). 16. Templates for CNBr-cleaved peptides from type I, II, III, and V collagen can be found in refs. (12–14). 17. As stated above, hydroxyproline assays can be used to quantitate the amounts of soluble and insoluble (cross-linked) collagens from the neutral salt-, acetic acid-, pepsin-, and CNBrdigested extracts/pellets, once they are lyophilized and re-suspended in 0.1 M HCl. 18. A limitation of this technique is that it cannot be used to identify collagen IV chains by peptide digestion. Thus, other techniques such as immunohistochemistry or Western blotting may be required for the detection of collagen IV. Acknowledgements The author is supported by Career Development Fellowships from the National Health & Medical Research Council (NHMRC) of Australia and the National Heart Foundation of Australia (NHFA).

References 1. Eddy, A.A. (2000) Molecular basis of renal fibrosis. Pediatr. Nephrol. 15, 290–301. 2. Becker, G.J. Hewitson, T.D. (2000) The role of tubulointerstitial injury in chronic renal failure. Curr. Opin. Nephrol. Hypertens. 9, 133–138. 3. Hewitson, T.D., Mookerjee, I., Masterson, R., Zhao, C., Tregear, G.W., Becker, G.J., et al.(2007) Endogenous relaxin is a naturally occurring modulator of experimental renal tubulointerstitial fibrosis. Endocrinology 148, 660–669.

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4. Neuman, R.E. Logan, M.A. (1950) The determination of hydroxyproline. J. Biol. Chem. 184, 299–306. 5. Bateman, J.F., Lamande, S.R., Ramshaw, J.A.M. 1996; Collagen superfamily. In: Comper WD, ed. Extracellular Matrix Vol. 2., Molecular Components and Interactions. Harwood Academic Publishers, Amsterdam, The Netherlands: 22–67. 6. Jackson, D.S. Cleary, E.G. (1967) The determination of collagen and elastin. Methods Biochem. Anal. 15, 25–76. 7. Gallop, P.M. Paz, M.A. (1975) Posttranslation protein modification, with special attention to collagen and elastin. Physiol. Rev. 55, 418–487. 8. Bergman, I. Loxley, R. (1963) Two improved and simplified methods for the spectrophotometric determination of hydroxyproline. Anal. Chem. 35, 1961–1965. 9. Samuel, C.S., Zhao, C., Bond, C.P., Hewitson, T.D., Amento, E.P., Summers, R.J. (2004) Relaxin-1-deficient mice develop an age-related progression of renal fibrosis. Kidney. Int. 65, 2054–2064. 10. Lekgabe, E.D., Kiriazis, H., Zhao, C., Xu, Q., Moore, X.-L., Su, Y., et al. (2005) Relaxin reverses cardiac and renal fibrosis in spontaneously hypertensive rats. Hypertension 46, 412–418. 11. Sykes, B., Puddle, B., Francis, M., Smith, R. (1976) The estimation of two collagens from human dermis by interrupted gel electrophoresis. Biochem. Biophys. Res. Commun. 72, 1472–1480. 12. Scott, P.G., Veis, A. (1976) The cyanogen bromide peptides of bovine soluble and insoluble collagens. I. Characterization of peptides from soluble type I collagen by sodium dodecylsulphate polyacrylamide gel electrophoresis. Connect. Tissue Res. 4, 107–116. 13. Scott, P.G. Veis, A. (1976) The cyanogen bromide peptides of bovine soluble and insoluble collagens. II. Tissue-specific cross-linked peptides of insoluble skin and dentin collagen. Connect. Tissue Res. 4, 117–129. 14. Bornstein, P., Sage, H. (1980) Structurally distinct collagen types. Ann. Rev. Biochem. 49, 957–1003.

Chapter 17

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

Abstract Protein profiling has numerous applications in renal research including the detection of protein biomarkers with aberrant expression levels during disease development. Such information is essential for early diagnosis and will aid the improvement of patient management and minimise the progression of disease. Further to this, data generated from these studies will assist the elucidation of the precise mechanisms of disease development and can lead to the discovery of potential drug targets. Surface enhanced laser desorption/ionisation time of flight mass spectrometry (SELDI-TOF MS), is emerging as a popular profiling tool for such studies. It incorporates the methods of solid-phase chromatography and TOF-MS in a single platform. This chapter provides a guide for establishing kidney profiling experiments using SELDI-TOF MS and will cover the following topics: 1) preparation of tissue extracts; 2) array processing, including optimisation of conditions for biomarker discovery; and 3) data acquisition/analysis. Keywords Biomarkers, Renal, Proteomics, Surface-enhanced laser desorption/ ionization time-of-flight mass spectrometry

1

Introduction

A global search of the proteome, called protein profiling, examines protein expression differences in biological systems under varying conditions. It has a vast array of applications in the field of renal research, including discovery of disease markers in nephropathology. This information not only aids to elucidate the mechanisms of disease, but also has important diagnostic implications as early detection is critical for improving patient management and minimising disease progression. Furthermore, information obtained from such studies can assist in the discovery of novel drug targets (1, 2). Protein profiling using surface enhanced laser desorption/ionisation time of flight mass spectrometry (SELDI-TOF MS) has become increasingly popular because of its sensitivity, ease of operation, and discriminatory power (3). From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_17, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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SELDI-TOF MS incorporates the methods of chromatography and TOF-MS into a single platform (4). ProteinChip™ arrays with specific chromatography properties (hydrophobic, ion exchange, and metal affinity capture) (5) are utilised to capture proteins with common biochemical properties, whilst remaining material is washed away. Further selectivity can be achieved by assessing different adsorption and wash buffers. The fraction of the proteome retained on the array can then be directly analysed by TOF-MS, resulting in a “profile” of proteins characterised by the mass to charge value (m/z) and signal intensities (4). SELDI-TOF MS profiling primarily involves two major steps, including 1) the discovery phase, the goal of which is to optimise conditions for biomarker discovery and search the proteome for putative biomarkers; and 2) the validation phase, during which the putative biomarkers are verified. Subsequent to this, it is important to purify the biomarker in preparation for sequencing to determine the identity of the unknown protein. SELDI-TOF technology can assist greatly in the purification process, after which point, standard MS/MS or Edman degradation sequencing methods are employed to identify the protein. This chapter provides a guide for establishing kidney profiling experiments using SELDI-TOF MS with a focus on biomarker discovery. Preparation of tissue extracts and processing of arrays will be described in detail. Please note that many of the concepts described in this chapter can also be applied to other tissues/ sample types.

2 2.1

Materials Reagents

1. Liquid nitrogen. 2. Microfuge (1.5-mL) tubes. 3. Extraction buffer: 10 mM Tris-HCl, 10 mM NaCl, 0.1% Triton X-100 pH 7.6, and protease inhibitor (complete mini EDTA free; Roche Diagnostics, Mannheim, Germany). 4. Bradford reagents and bovine serum albumin (BSA) standards (Quantichrome Protein Determination Assay; Bio Assay Systems, Hayward, CA, USA). 5. ProteinChip™ arrays: Q10, CM10, H50, and IMAC30 (Bio-Rad Laboratories, Hercules, CA, USA). 6. 100 mM sodium acetate (NaAc), pH 4. 7. Acetonitrile (ACN), HPLC grade. 8. Trifluoroacetic acid (TFA), HPLC grade. 9. Q10 binding/wash buffer: 100 mM Tris-HCl, pH 9. 10. CM10 binding/wash buffer: 20 mM NaAc, pH 5. 11. H50 binding/wash buffer: 5% ACN, 0.1% TFA.

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12. IMAC30 binding/wash buffer: phosphate-buffered saline (PBS) with 137 mM NaCl and 10 mM phosphate buffer, pH 7.4. 13. 100 mM Copper sulphate. 14. 1 mM HEPES, pH 7.2. 15. Kimwipes™ (Kimberly-Clarke, Missions Point, Australia). 16. Matrix/energy-absorbing molecule (EAM): alpha-cyano-4-hydroxy cinnamic acid (CHCA) and sinapinic acid (SPA), 5 mg/tube (Bio-Rad Laboratories).

2.2 1. 2. 3. 4. 5.

3 3.1

Equipment Tissue homogeniser. Mortar and pestle. Optical density (OD) micro plate reader (Asys Hitech, Nordstrasse, Austria). Bioprocessor (Bio-Rad Laboratories). ProteinChip™ reader, PBSIIc (Bio-Rad Laboratories).

Methods Tissue Collection

1. Collect tissue, ensuring handling is kept to a minimum. Use gloves to reduce the risk of contamination and protein degradation. 2. To distinguish protein profiles from specific regions within the kidney (i.e. cortex, medulla, renal pyramids, renal papilla, etc.), these regions should be immediately dissected out in cold PBS. 3. Weigh tissue and store in 1.5-mL tubes at –80°C as soon as possible (see Note 1).

3.2

Protein Extraction

1. Pulverise tissue into a fine powder using a mortar and pestle (in liquid nitrogen). 2. Immediately transfer the disrupted tissue to a 1.5-mL tube (see Note 2). Add extraction buffer to tissue (10 µL/mg tissue) and place on ice (see Note 3). Further homogenise the tissue using a rotor–stator homogeniser (10–20 s at maximum speed) or by passing extract through a 19-gauge needle 10 times.

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3. Leave homogenate on ice for 1 h with periodic mixing, followed by centrifugation at 7,840 × g at 4°C for 10 min. 4. Transfer the supernatant (containing total cellular proteins) into a fresh tube. 5. Aliquot samples, store at –80°C, and use only as required. The remaining material ideally should be discarded, however, if this is not possible, it is important to ensure that all samples experience the exact same number of freeze–thaw cycles (see Note 4).

3.3

Total Protein Assay

1. Perform a Bradford protein quantification assay to determine the total protein concentration of each sample. The assay described below is a Quantichrome Protein Determination Assay (Bio Assay Systems). 2. Make a BSA standard curve using the following concentrations: 0, 0.1, 0.2, 0.3, 0.4, 0.6, 0.8, and 1 mg/mL and load 10 µL of each standard into a well of a microplate. 3. Dilute kidney extract 1:10 into H2O and load 10 µL of sample per well. 4. Make working reagent by combining 1 volume of 5× stock and 5 volumes of H2O, then bring to room temperature. Add 200 µL of working reagent per well and mix with sample by pipetting up and down several times. 5. Read at 595 nm to determine the OD values of the samples and standards. Plot the BSA standard curve to generate the R2 value and slope intercept equation. The R2 value is used to gauge the models “goodness of fit”; it is displayed as a number between 0.0 and 1.0, with values closer to 1 reflecting a good-quality model. The slope–intercept equation (y = mx + b) is utilised to ascertain the concentration of the test sample, where y is the unknown value (concentration), m is the slope, x is the OD value of the test sample, and b is the y-intercept value.

3.4

Protein Standardisation

1. Prior to diluting samples into binding buffer, it is important to initially adjust the samples to the same concentration using extraction buffer (see Note 5). 2. Dilute samples into the appropriate binding buffer to a final concentration of 1 mg/mL (see Note 6).

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3.5

Optional Steps for Extract Preparation

3.5.1

Removal of Abundant Proteins

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As the arrays have a limited surface capacity, it can be advantageous to remove highly abundant proteins or fractionate samples to enhance the detection of low abundant proteins by limiting competition for the binding sites. Removal of abundant proteins can be achieved using a depletion approach in which the selected proteins are removed via binding to specific antibodies conjugated to resins and the remaining material is collected for further analysis. There are however a number of disadvantages to using the depletion approach, including the fact that biomarkers may also bind high abundant proteins and as a consequence will be removed. In addition, recent studies have revealed that some biomarkers are proteolytic-derived fragments of abundant proteins (6), which would also be discarded using this methodology. Fractionation segregates abundant proteins into limited fractions. Fractionation is commonly performed using anion exchange chromatography and a step-wise gradient from pH 9 through to pH 4 with a final organic wash. Size fractionation can also be achieved to improve detection of proteins within a particular molecular weight range using size exclusion chromatography or molecular weight cut-off columns.

3.6

Study Design

3.6.1

Discovery Phase

The discovery phase of the study is performed to determine the optimal conditions for biomarker detection. Using a small data set, various surfaces and adsorption/ wash buffers are assessed with the aim of finding conditions which yield the highest peak count and allow expression differences to be detected. The following combination of arrays/buffers provide a good starting point for optimisation: Anion exchange: Q10 arrays: Q10 binding/wash buffer (100 mM Tris HCl, pH 9). Cation exchange: CM10 arrays: CM10 binding/wash buffer (20 mM NaAc, pH 5). Reversed phase: H50 arrays: H50 binding/wash buffer (5% ACN and 0.1% TFA). Immobilised metal affinity capture (IMAC): IMAC30 arrays pre-coupled with 100 mM copper sulphate: IMAC30 binding/wash buffer (PBS). SPA is generally the energy-absorbing molecule of choice for profiling experiment as it works well for both peptides and proteins. If the mass range of interest

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falls below 30,000 Da, consider also assessing other matrices such as CHCA, which are particularly suitable for smaller molecules. If the conditions described above do not yield the desired results, further selectivity can be achieved by assessing different adsorption and wash buffers. H50 arrays: increasing the organic solvent content (0–50% ACN or methanol with 0.1–1% TFA) in the adsorption and wash buffers will increase selectivity on the H50 array surface. Proteins with greater hydrophobicity will be retained on the array surface in the presence of higher organic solvent content. IMAC30 arrays: increased selectivity on the IMAC30 surface can be achieved by adding imidazole (5–10 mM). Also consider assessing other metals such as zinc sulphate, which is a common metal used for profiling experiments. Galium can also be used to enrich for phosphorylated proteins. Ionic exchange arrays (Q10 and CM10 arrays): selectivity on the ionic exchange arrays can be enhanced by increasing the salt concentration of the buffer. Further to this, increasing or decreasing the pH of the binding buffer for the CM10 and Q10 arrays, respectively, will also increase surface selectivity. Recommended buffers for both array types include 10–100 mM Tris-HCl pH 7–9, 10–100 mM ammonium phosphate pH 6–8, and 10–100 mM sodium acetate pH 4–6.

3.6.2

Validation Phase

Once the optimal conditions have been determined, the validation phase of the study can be conducted. Larger data sets are now employed to verify the preliminary results. It is critical to employ the exact conditions used during the discovery phase, as any differences introduced may affect the resulting profile. Furthermore, it is crucial that all samples are treated in an identical manner to ensure that the data generated is reliable and reproducible.

3.7

Processing ProteinChip™ Arrays

Chip processing can be performed “on spot” or with a bioprocessor. “On spot” analysis involves directly loading buffer/samples onto the arrays, generally in 5-µL volumes. A bioprocessor allows larger volumes of up to 200 µL to be loaded onto the array’s surface, which can be beneficial if the sample is dilute. Furthermore, it allows up to 12 arrays to be processed simultaneously, making it amenable for highthroughput applications. For large-scale experiments, it is highly recommended that a bioprocessor is used to minimise chip-to-chip variation due to differential handling.

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3.7.1 3.7.1.1

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Q10, CM10, and H50 Arrays: Bioprocessor Pre-equilibration and Sample Binding

1. Slide arrays into the cassette, being careful not to touch the spots (see Note 7), place reservoir on top of the cassette, and assemble in the bioprocessor. 2. Pre-equilibrate arrays in the appropriate binding buffer; load each well with 100 µL of binding buffer and cover with Parafilm. Centrifuge the unit for 1 min at 140×g to remove air bubbles and ensure that the buffer comes into direct contact with the surface of the array (unless otherwise stated, the bioprocessor is centrifuged after the addition of buffer/sample). Incubate for 5 min on a shaking table. 3. Decant buffer and repeat step 2 for a total of two washes. 4. Load 100 µL of sample per well and incubate for a minimum of 60 min (see Note 8).

3.7.1.2

Washes

1. Decant samples, load 100 µL of binding/wash buffer per well, and incubate for 5 min on a shaking table. Repeat for a total of three washes. 2. Load 100 µL of 1 mM HEPES, pH 7.2 per well and incubate for 1 min on a shaking table (see Note 9). Repeat for a total of two washes. 3. Decant buffer and remove the array cassette from the bioprocessor. Use a Kimwipe™ to remove excess buffer from each spot. 4. Air-dry the array for ∼10 min in preparation for EAM addition (see Note 10).

3.7.1.3

Energy-Absorbing Molecule Addition

1. Make 50% ACN, 0.5% TFA by combining 200 µL of ACN with 200 µL of 1% TFA. 2. Make 100% saturated EAM by adding 200 µL of 50% ACN and 0.5% TFA to a tube containing 5 mg of CHCA or SPA. 3. Vortex the matrix solution and incubate for 5 min. 4. Centrifuge the matrix solution (1 min at 13,250×g) to pellet undissolved matrix, collect 100 µL of supernatant, and combine with 100 µL of 50% ACN and 0.5% TFA to make 50% saturated EAM. 5. Load 1 µL of 50% saturated EAM to each spot, dry the arrays, apply a second application of EAM, and dry again (see Note 11). Introduce arrays into the ProteinChip™ reader in preparation for data acquisition.

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3.7.2

IMAC Arrays: Bioprocessor

3.7.2.1

Precoating Arrays with Copper Sulphate

1. Assemble the IMAC arrays in the bioprocessor. Apply 100 µL of 100 mM copper sulphate solution per well and incubate on a shaking table for 5 min. 2. Decant the buffer and repeat step 1 for a total of two copper sulphate washes. 3. Discard the metal solution and neutralise the copper sulphate-coated IMAC array by the addition of 100 mM sodium acetate, pH 4 (100 µL/well) for 5 min. 4. Wash spots with 100 µL of 1 mM HEPES pH 7.2 for 5 min, repeat for a total of two washes. 5. Arrays can now be equilibrated in binding buffer; remaining steps are performed as described above in Sects. 3.7.1.1–3.7.1.3.

3.8

Data Acquisition Settings

For each new experiment it is important to optimise the data acquisition settings including the laser intensity and detector sensitivity. Sub-optimal laser settings can significantly affect the quality of the resulting spectra. If the laser energy is set too low, proteins will not be adequately extracted from the surface of the array leading to a low peak count. In contrast, laser intensities set too high will generate broad off-scale peaks and an unstable baseline. In the latter, accurate assessment of peak expression differences will not be possible as the detector will be saturated. It is therefore important to adjust the laser settings to yield peaks that are sharp and well resolved. In order to provide adequate coverage of the whole molecular weight range, it is advisable to optimise the laser energy settings for the low (< 20,000 Da) and higher mass ranges (> 20,000 Da). Larger proteins require additional laser energy to “fly”, hence the laser settings should be adjusted accordingly and increased during data acquisition within the higher mass range. Data subsequently generated can be represented in spectra or virtual gel view. A summary of the array processing/data acquisition method is shown in Fig. 17.1.

3.9

Data Analysis

Once data has been collected, prepare spectra for biomarker assessment by performing the following: baseline subtraction, calibration, noise definition, and normalisation.

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a

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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™ technology training course (Bio-Rad Laboratories) (see Color Plate 9)

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Baseline Subtraction

Baseline “noise” is a combination of electronic and chemical (EAM) noise and is controlled by the baseline subtraction function. Generally, the default settings are sufficient to stabilise the baseline, however, to ensure the baseline closely follows the spectrum, the fitting width values can be reduced.

3.9.2

Calibration

In order to ensure reliable mass accuracy, it is important to calibrate each spectrum. Create a calibration equation using standards that span the appropriate mass range combined with the corresponding matrix used for sample analysis. Apply the calibration equation to relevant spectra.

3.9.3

Noise Definition

Define the area in which local noise will be calculated. Typically the EAM portion of the spectra is excluded from the calculation; hence local noise is usually defined from ∼1,500 Da onwards.

3.9.4

Normalisation

Normalisation by total ion current is commonly employed for profiling studies. It involves linearly scaling the intensities to account for slight variations between spectra. The EAM portion of the spectrum is excluded from such calculations; hence the lower m/z limit is set to ∼1,500 Da onwards.

3.9.5

Biomarker Analysis

Peak clustering tools are used to search the spectra for potential biomarkers. Clustering tools group peaks of the same molecular weight across all samples and display the differences in expression between sample groups in the form of scatter or box and whiskers plots. P values are also ascertained to display the statistical significance in protein expression between groups (Fig. 17.2). Clusters with P values < 0.05 should be inspected manually to ensure that the software is not clustering on noise and the peak has been correctly labelled with the centroid function in the centre of the peak, and not to the side of the peak.

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Fig. 17.2 SELDI-TOF MS analysis of kidney tissue from healthy mice (control) and mice with unilateral ureteric obstruction (UUO). SELDI-TOF MS analysis of normal tissue and tissue from kidneys in which the ureter has been obstructed for 3 days (a, b). Data is represented in both spectra and virtual gel view. A potential biomarker upregulated in the disease group is highlighted and represented in a scatter plot (c), which displays the expression differences between sample groups

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3.10

Purification and Identification of Biomarkers

As a first step in the biomarker purification process, it is initially recommended to perform retentate chromatography with the ProteinChip™ array used to discover the biomarker. Retentate chromatography involves applying the sample containing the peak of interest to every spot on an array. Each spot is then washed with a buffer of increasing stringency, for example if the biomarker was identified on Q10 arrays using pH 9 buffer, the following buffers can be used during the washing procedures: Spot A: pH 9, Spot B: pH 8, Spot C: pH 7, etc. This approach will determine when the peak of interest elutes from the surface. These conditions can then be replicated on spin columns. As an extra confirmatory step, it is recommended to analyse the eluant on an array to ensure the elution fraction contains the peak of interest. It may be necessary to complete several column runs, particularly if the protein of interest is not present in high concentrations. Elution fractions can subsequently be pooled/ concentrated using a vacuum concentrator or freeze drier and electrophoresed on a 1D sodium dodecyl sulphate (SDS) polyacrylamide gel electrophoresis (PAGE) gel. After excising the band and eluting the protein from the gel, it is recommended to analyse a fraction of the sample on a NP20 array to confirm the band is the correct molecular mass (see Note 12). Once purified, standard sequencing methodologies such as Edman degradation, peptide mass fingerprinting or tandem mass spectrometry (matrix-assisted laser desorption ionisation [MALDI]-TOF/TOF) can be employed to determine the identity of the unknown protein.

4

Notes

1. It is important to employ a standard operating procedure for collection and handling of tissue to ensure that differences detected between sample groups are a reflection of the phenotypes being examined rather than a consequence of variations introduced during tissue processing. 2. Use consumables with low protein binding plastics such as polypropylene. Ensure the same consumables are utilised throughout the duration of the study. 3. Most common buffers, salts, and non-ionic detergents are compatible with the SELDI-TOF MS system, however, certain chemicals should be avoided. EDTA will strip metal ions from IMAC surfaces. High concentrations of ionic detergents such as SDS will suppress ionisation. DTT and PEG can also be problematic and hence should be avoided. 4. Subjecting samples to several freeze–thaw cycles can generate degradation products/oxidative modification (+16 Da mass shift) and alter the resulting profile. 5. It is critical to ensure that all samples are identical in regards to protein concentration and the final buffer constituents. The protein concentration affects peak number and intensity. The final buffer constituents of the sample (pH, salt concentration, etc.) affect the binding stringency and resulting profiles. 6. For each new experiment it is important to initially determine the optimal protein concentration by performing a series of titrations in replicates. The conditions that yield the highest peak count and reproducibility should be selected. In general it is recommended to use protein extracts within a concentration range of 50–2,000 µg/mL. As the capacity of the array varies

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9.

10. 11. 12.

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for each chip type, depending on the surface chemistry, it can be advantageous to optimise the protein concentration for each type of array. During chip handling use non-latex powder-free gloves such as nitrile. Latex contamination can generate a series of peaks complicating spectra analysis. Sample addition: if processing large numbers of samples, it is advisable to dispense samples in a 96-well polypropylene plate using the same format that will be applied to the bioprocessor. After equilibrating arrays, the samples can simply be transferred from the plate into the bioprocessor using a multichannel pipette. Samples should be incubated on the array for a minimum of 60 min to ensure the reaction has reached equilibrium, this will improve reproducibility. It is also advisable to randomise samples to reduce systematic bias and include replicates to assess assay reproducibility. Washing arrays with 1 mM HEPES, pH 7.2 buffer is conducted to remove contaminants such as detergent and salts. Detergents create a series of low molecular weight peaks that can complicate data interpretation. The presence of salts e.g. Na+ and K+ can lead to the generation of salt adducts that can interfere with data analysis and decrease sensitivity. Ensure that the EAM drying time does not exceed 20 min, extended incubations can decrease peak intensity and negatively influence assay reproducibility. Use a 2-µL calibrated pipette for matrix addition. Ensure the method of EAM application, including drying time, is consistent as this can affect the assay reproducibility. During development of the purification strategy it is beneficial to screen various arrays to determine if the biomarker binds to other surfaces. This may provide another means of separating and enriching for the protein of interest.

Acknowledgments This work was supported in part by a National Health & Medical Research Council (NHMRC) R.D.Wright/National Heart Foundation of Australia (NHFA) Career Development Fellowship to C.S.S and an Ian Potter Foundation grant to the Howard Florey Institute.

References 1. Janech, M.G., Raymond, J.R., and Arthur, J.M. (2007). Proteomics in renal research. Am. J. Physiol. Renal Physiol. 292: F501–F512. 2. Bonventre, J.V. (2002). The kidney proteome: A hint of things to come. Kidney Int. 62: 1470–1471. 3. Issaq, H.J., Veenstra, T.D., Conrads, T.P., and Felschow, D. (2002). The SEDLI-TOF MS approach to proteomics: protein profiling and biomarker identification. Biochem. Biophys, Res. Com. 292: 587–592. 4. Wiesner, A. (2004). Detection of tumor markers with ProteinChip technology. Curr. Pharm. Biotechnol. 5: 45–67. 5. Tang, N., Tornatore, P., and Weinberger, S.R. (2004). Current developments. In: Seldi Affinity Technology. Mass Spectrometry Reviews. 23: 34–44. 6. Fung, E.T., Yip, T.T., Lomas, L., Wang, Z., Yip, C., Meng, X.Y., Lin, S., Zhang, F., Zhang, Z., Chan, D.W., and Weinberger, S.R. (2005). Classification of cancer types by measuring variants of host response proteins using SELDI serum assays. Int. J. Cancer. 115: 783–789.

Chapter 18

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

Abstract Small synthetic interfering RNA duplexes (siRNAs) can selectively suppress gene expression in somatic mammalian cells without the nonselective toxic effects associated with double-stranded RNA (dsRNA). However, in vivo delivery of siRNA targeting the kidney has been described in only a few reports. We have found that injection of synthetic siRNAs via the renal artery, followed by electroporation, can be therapeutically effective in silencing the expression of specific genes in the glomerulus. Here we provide details of an experimental protocol showing that 1) delivery of siRNA targeting enhanced green fluorescent protein (EGFP) to the kidney in the transgenic “green” rat reduces endogenous EGFP expression, mainly in the glomerular mesangial cells, and that 2) delivery of siRNA targeting transforming growth factor (TGF)-β1 to the kidney significantly suppresses messenger RNA (mRNA) and protein expression of TGF-β1, thereby ameliorating the progression of matrix expansion in experimental glomerulonephritis. In addition, we describe the application of vector-based RNA interference (RNAi) (small hairpin RNA [shRNA]), which also inhibits TGF-β1 expression in vivo. Keywords: Electroporation, Enhanced green fluorescent protein; RNA interference; Small interfering RNA; Small hairpin RNA

1

Introduction

RNA interference (RNAi) is initiated by the introduction of double-stranded RNA (dsRNA) into the cell and leads to the sequence-specific destruction of endogenous RNA [1]. RNAi-induced gene-specific silencing has proven successful in Caenorhabditis elegans and plants. However, the use of long dsRNA in vertebrates is problematic because they induce a generalized suppression of protein synthesis and cell death by activating the interferon pathway. Tuschl et al. made a crucial breakthrough, finding that small synthetic interfering RNA duplexes (siRNAs) can selectively silence the expression of complementary genes in somatic mammalian cells without the nonselective toxic effects associated with long dsRNAs [2]. These observations in mammalian cells not only permitted the next generation of From: Methods in Molecular Biology, Vol. 466: Kidney Research DOI: 10.1007/978-1-59745-352-3_18, Edited by: T.D. Hewitson and G.J. Becker © Humana Press, Totowa, NJ

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genetic research to occur, but also allowed the development of new therapeutic approaches. An important step in realizing the potential of RNAi as a therapeutic tool is determining whether this mechanism can be triggered in vivo. Recently, we developed an electroporation-mediated siRNA transfer system targeting the kidney, whereby siRNA is injected via the renal artery, then electric pulses are applied using tweezers-type electrodes [3]. This system allows us to deliver siRNA mainly to the glomeruli, a region central to the inflammatory response in the initiation and progression of various kidney diseases. To prove the therapeutic application of transfected siRNA in an animal disease model, we targeted transforming growth factor (TGF)-β1, a potent cytokine that plays an important role in the fibrogenic phase of various diseases, including glomerulonephritis. Also, we examined the efficacy of a siRNA-producing DNA-based vector system (small hairpin RNA [shRNA]). Here, we provide details of the protocol used to conduct siRNA or shRNA transfer via the renal artery.

2

Materials

2.1

2.1.1

siRNA-Mediated Inhibition of Enhanced Green Fluorescent Protein (EGFP) Expression in Glomeruli Preparation of siRNA Targeting EGFP

1. siRNAs (21 nucleotides long) targeting EGFP and the scrambled genes were chemically synthesized as 2′ bis (acetoxyethoxy)-methyl ether-protected oligonucleotides, deprotected, annealed, and purified by Dharmacon Research (Lafayette, CO, USA). The strands of siRNA targeting EGFP are 5′-P. GGCUACGUCCAGGAGCGCACC-3′ (sense) and 5′-P.UGCGCUCCUGGACGUAGCCUU-3′ (antisense) [4]. Aliquots of 20 µM siRNA solution in 1× annealing buffer (100 mM KOAc, 30 mM HEPES-KOH pH 7.4, and 2 mM MgOAc) can be stored at –20°C.

2.1.2 1. 2. 3. 4. 5.

Transfer of siRNA Targeting EGFP Via the Renal Artery

siRNA dilution buffer: nuclease-free phosphate-buffered saline (PBS). Catheter: 24-gauge (cat no. SR-OT2419CW; Terumo, Tokyo, Japan). Injection needle: 26-gauge (Terumo). Syringe: 1 mL and 2.5 mL with lock system (Terumo). Animals: individuals of the “green” transgenic rat line SD-TgN (act-EGFP) Osb4 8 [5], weighing approximately 200 g (Japan SLC, Inc., Hamamatsu, Japan) (see Note 1).

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6. Pentobarbital: 50 mg/mL (Dainippon Sumitomo Pharma, Osaka, Japan). 7. Shaver (commercial). 8. Povidone–iodine solution. 9. Papaverine hydrochloride: 40 mg/mL (Dainippon Sumitomo Pharma). 10. “Pillow”: fold a paper towel into a pad measuring approximately 5×20×3 cm and wrap with aluminum foil. 11. Aneurysm clip: Sugita aneurysm clip (cat nos. 07-940-01 or 07-940-96; Mizuho-ika Kogyo, Tokyo, Japan). 12. Forceps for aneurysm clip (cat nos. 07-941-01 or 07-942-01, straight type; Mizuho-ika Kogyo). 13. Other anatomical instruments, including scissors and retractor. 14. Surgical glue: Aron Alfa A “Sankyo” (Toagosei, Tokyo, Japan). 15. Cotton swab. 16. Ampicillin sodium (cat no. 012-20162; Wako, Osaka, Japan): dissolved in PBS at a concentration of 20 mg/mL. 17. Skin stapler (Igarashi-ika Kogyo, Tokyo, Japan). 18. [Optional] Oval-shaped tweezer-type electrodes; 15×10 mm, specifically ordered from BTX (San Diego, CA, USA). 19. [Optional] Electric pulse generator (Electro Square Porator T820M; BTX). 20. [Optional] Switch box (MBX-4; BTX). 21. [Optional] Graphic pulse analyzer (Optimizor 500; BTX).

2.1.3

Assessment of EGFP Expression in the Glomeruli

1. Syringe: 20 mL. 2. Four percent (w/v) paraformaldehyde (PFA) solution (pH 7.4). Store at 4°C. Use within 1 week after preparation. 3. Ten percent (also 20% and 30%) sucrose in PBS: dissolve 10 g (or 20 or 30 g) of sucrose in 100 mL of PBS and autoclave at 105 ° C for 1 min. Store at 4°C. 4. OCTTM compound (Tissue-Tek ΤΜ, Sakura Finetek USA, Inc., Torrance, CA, USA). 5. Liquid nitrogen. 6. Wash buffer (TBST): 20 mM Tris-HCl (pH 7.5), 137 mM NaCl, and 0.1% Tween 20 (v/v). 7. Blocking solution: 5% (v/v) normal horse serum in TBST. 8. Primary antibody: clone OX-7 (gift from Dr. Ken-ichi Isobe and Prof. Seichi Matsuo, Nagoya University, Nagoya, Japan) (see Note 2). Dilute in blocking solution at 1:100 just before use. 9. Texas red-conjugated anti-mouse IgG (Vector Laboratories, Inc., Burlingame, CA, USA). Dilute in blocking solution at 1:200 just before use. 10. Mounting medium: PermaFluor ΤΜ Aqueous Mounting Medium (cat no. 434990; Thermo ELECTRON Corp.).

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siRNA-Mediated Inhibition of TGF-b1 in the Glomeruli

2.2.1 2.2.1.1

Preparation of siRNA or siRNA Expression Vector Targeting TGF-b1 siRNA

1. siRNAs (21 nucleotides long) targeting TGF-β1 and the scrambled genes are chemically synthesized as 2′ bis (acetoxyethoxy)-methyl ether-protected oligonucleotides, deprotected, annealed, and purified by Dharmacon Research. The strands of siRNA targeting TGF-β1 are 5′-P.GUCAACUGUGGAGCAACACdTdT3′ (sense) and 5′-P.GUGUUGCUCCACAGUUGACdTdT-3′ (antisense) [3]. Aliquots of 20 µM siRNA solution in 1× annealing buffer (100 mM KOAc, 30 mM HEPES-KOH pH 7.4, and 2 mM MgOAc) can be stored at –20°C.

2.2.1.2

siRNA Expression Vector

1. A pair of oligodeoxynucleotides (64 nucleotides long), encoding shRNAs targeting TGF-β1 and with a BamHI or HindIII site are synthesized by Bex (Tokyo, Japan). The sequences of the oligodeoxynucleotides are shown in Fig. 18.3a. 2. siRNA expression vector featuring the human U6 RNA pol III promoter: pSilencer 2.0-U6 siRNA expression vector (0.1 mg/mL; Ambion, Inc., Austin, TX, USA). 3. 5× Annealing buffer: 0.5 M Tris-HCl (pH 7.4) and 0.35 M MgCl2. 4. TE: 10 mM Tris-HCl and 1 mM EDTA (pH 8.0). 5. 4× Ligation buffer: 200 mM Tris-HCl (pH 7.6). 6. Ligation enzyme: solution I in the DNA Ligation Kit Ver. 2.1 (Takara, Shiga, Japan). 7. Plasmid DNA purification kit: High Purity Maxiprep ΤΜ Plasmid Purification Kit (Marligen Biosciences, Urbana Pike Ijamsville, MD, USA). 8. [Others] LB plate, competent Escherichia coli cells (JM109).

2.2.2

Animals and Induction of Thy-1 Nephritis

1. Sprague-Dawley (SD) rats (Japan SLC, Inc.), weighing approximately 200 g (see Note 1). 2. Pentobarbital: 50 mg/mL (Dainippon Sumitomo Pharma). 3. Monoclonal antibody (mAb) 1-22-3 (kind gift from Dr. H. Kawachi of Niigata University) (see Note 3).

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Transfer of siRNA or siRNA Expression Vector Targeting TGF-b1 Via the Renal Artery (See Sect. 2.1.2)

2.2.4

Assessment of TGF-b1 Expression in the Glomeruli

1. Sieving mesh: pore sizes 125 µm, 180 µm, and 75 µm (Bunsekifurui, Tokyo, Japan). 2. Tissue grinder (1 mL Dounce Tissue Grinder; Wheaton Science Products, Millville, NJ, USA). 3. Lysis buffer: 50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1%(v/v) Nonidet-P40, 10% (v/v) glycerol, 1 mM PMSF, 1 µg/mL aprotinin, 1 µg/mL leupeptin, and 0.5 mM sodium orthovanadate. 4. 3× Laemmli buffer. 5. 15% Sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE) gel. 6. 10× SDS-PAGE running buffer: 250 mM Tris, 1.92 M glycine, and 1% (w/v) SDS in 1 L distilled water. Store at room temperature. Dilute with distilled water just before use. 7. 1× Transfer buffer: 25 mM Tris, 192 mM glycine, 20% methanol in 1 L distilled water. Adjust pH to 8.3. Store at 4°C. 8. Polyvinylidene difluoride membrane (Hybond-P PVDF Membrane; Amersham Biosciences). 9. TBST: 20 mM Tris-HCl (pH 7.5), 137 mM NaCl, and 0.1% Tween 20 (v/v). 10. Blocking solution: 5% (v/v) bovine serum albumin in TBST. 11. Primary antibody: polyclonal antibody against TGF-β1 (Promega, Madison, WI, USA). 12. Secondary antibody: horseradish peroxidase-conjugated goat anti-rabbit IgG (Dako, Glostrup, Denmark). 13. Chemiluminescent reagent: SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL, USA). 14. X-ray film: Hyperfilm ΤΜ (Amersham Biosciences). 15. BCA ΤΜ protein assay reagent (Pierce).

3

Methods

We usually use electroporation when transfecting siRNA, antisense oligodeoxynucleotides (ASODN) or DNA vectors (including siRNA expression vectors) via the renal artery because electroporation increases the transfection efficiency, especially for DNA vectors [6], but we and other groups have demonstrated that a substantial amount of siRNA or ASODN can be successfully transfected into the glomeruli without electroporation (data not shown), with hydrodynamic pressure as one of the

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main driving forces. Electroporation devices are very costly, and the electrode must be specially ordered. Therefore, we recommend that this protocol is first attempted without electroporation; here we have described the protocols that are related to electroporation as being optional.

3.1

3.1.1

siRNA-Mediated Inhibition of EGFP Expression in the Glomeruli Preparation of siRNA Targeting EGFP

1. Dilute 50 µg of siRNA solution for EGFP and the scrambled gene in 500 µL of nuclease-free PBS, respectively. Store the solution at room temperature until transfection and use within several hours (see Note 4).

3.1.2

Transfer of siRNA Targeting EGFP Via the Renal Artery

1. Fill two or more 1-mL syringes with PBS. Attach the 26-gauge needle to one syringe. Expel the air from the syringes. 2. Remove the flashback chamber of a 24-gauge catheter and flush with PBS (keep the chamber off hereafter). 3. Fill the 2.5-mL syringe with the siRNA solution and expel the air carefully. 4. Anesthetize EGFP-transgenic rats via intraperitoneal injection of pentobarbital (50 mg/kg). 5. Shave the hair around the rats’ abdomens and disinfect with povidone–iodine or ethanol. 6. Make a midline incision from the bladder to the xiphoid process. 7. Set the retractor and place the “pillow” beneath the body. 8. Move the digestive tract downward and to your left side to reveal the left kidney and renal artery. 9. Peel off the retroperitoneum around the root of the left renal artery using a cotton swab and/or small forceps (Fig. 18.1a). 10. Expose the aorta at the level of the left renal artery and its proximal site and remove excessive connective tissue (see Note 5). 11. Drop a small amount of papaverine hydrochloride around the outcrops of the aorta to avoid vasoconstriction. 12. Move to the head side of the rat. Puncture the aorta (not the renal artery) using a 24-gauge catheter where it branches into the left renal artery using your right hand while creating some tension in the renal artery using your other hand (see Notes 6 and 7) (Fig. 18.1b). Observe for “flash back” as blood slowly fills the catheter. After insertion of both the stylet and the catheter, advance only the catheter ~5 mm further into the left renal artery and remove the needle. 13. Fill the end of the catheter hub with a small amount of PBS, using a syringe with needle to avoid infusing air (repeat this step whenever a new syringe is connected to the catheter).

Fig. 18.1 Method used to transfer the siRNA via the renal artery. a Peel off the retroperitoneum around the root of the left renal artery (dotted line). b Puncture the aorta with a 24-gauge catheter. Transverse section is also shown (aorta, arrow; catheter, arrowhead). c Advance the catheter into the left renal artery and clamp the aorta with an aneurysm clip (clamp 1; arrow). d Connect the syringe filled with siRNA solution. e Infuse the siRNA solution using a single shot. Immediately after injection, clamp at the renal hilum (clamp 2; arrow). f When you attempt the electroporation method, clamp the renal vein and artery together with catheter (clamp 3; arrow). g Sandwich the left kidney between a pair of electrodes and deliver electric pulses. h After removal of the clamp(s) except clamp 1, pipette a small amount of surgical glue onto the puncture.

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14. Connect the 1-mL syringe filled with PBS to the catheter and inject ~500 µL of PBS. If the kidney is perfused thoroughly (i.e., if the kidney becomes yellow), the catheter is inserted properly. Disconnect the syringe. If the kidney is not perfused thoroughly (for example only the upper portion is perfused), it is likely that the catheter is inserted beyond the branching portion of the renal artery, so retract the catheter carefully and repeat the perfusion (see Note 8). 15. Clamp the aorta with an aneurysm clip proximal to the left renal artery (clamp 1) (Fig. 18.1c). 16. Repeat step 14 to completely remove the red and white blood cells. 17. Connect the 2.5-mL syringe (with lock system) filled with siRNA solution to the catheter while fixing the tube with forceps to avoid accidental removal (Fig. 18.1d). 18. Retract the catheter and syringe carefully to the greatest extent possible. 19. Infuse the siRNA solution into the left kidney using a single shot (within 1 s). 20. [Assistant] Clamp the renal artery and vein together with the retroperitoneum with an aneurysm clip at the renal hilum immediately after injection (clamp 2) (Fig. 18.1e). 21. [Optional] Clamp the renal vein together with the catheter (and renal artery) with an aneurysm clip at the proximal portion of the left renal artery (clamp 3) so that the catheter does not come out of the renal artery when the electric pulse is delivered in the next step (Fig. 18.1f). 22. [Optional] [Operator] Hold the rat on the table with both hands, because the rat will move during electroporation. [Assistant] Sandwich the left kidney between a pair of oval-shaped tweezer-type electrodes (Fig. 18.1g). Deliver electric pulses. Administer six pulses of 100 V at a rate of one pulse per second, with each pulse being 50 ms in duration. 23. Remove one or two clamps (clamp 2 and clamp 3 if applied), but not the clamp at the most proximal site (clamp 1). 24. [Operator] Place your fingers over the proximal and distal puncture sites and remove the catheter. Pipette a very small amount of surgical glue onto the puncture sites immediately after wiping the blood away using a cotton swab (Fig. 18.1h). Wait about 30 s, then remove your fingers slowly. After confirming hemostasis, remove clamp 1 (see Note 9). 25. Place 20 mg of ampicillin solution into the peritoneal cavity. 26. Suture the abdominal muscle and skin separately by using a skin stapler.

3.1.3

Assessment of EGFP Expression in the Glomeruli

1. Seven days after transfection, perfuse the rats with 20 mL of ice-cold PBS via the abdominal aorta. 2. Remove the kidneys, then slice them about 1-mm thick and immerse in icecold 4% neutral-buffered PFA solution for 4–6 h (see Note 10).

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3. Wash the kidney sections in PBS several times and place in 10% sucrose in PBS (see Note 11). 4. Replace the 10% sucrose solution with a 20% sucrose solution ~5 h later. 5. Replace the 20% sucrose solution with a 30% sucrose solution ~5 h later. 6. The next day, embed the kidney sections in OCT ΤΜ compound on a metal block chilled in the liquid nitrogen, and then freeze the sections to -80°C. 7. Using a cryostat, section the kidney further at 4-µm thick, and then air-dry the sections on glass slides at room temperature for 1 h. 8. Wash the sections in distilled H2O three times for 5 min each. 9. Wash the sections in wash buffer for 5 min. 10. Block each section with ~100 µL blocking solution for 30–60 min. 11. Remove the blocking solution and add ~100 µL of diluted primary antibody to each section. Incubate overnight at 4°C. 12. Remove the antibody solution and wash the sections in wash buffer three times for 5 min each. 13. Add ~100 µL of Texas red-conjugated secondary antibody to each section. Incubate for 30 min at room temperature. 14. Wash the sections in wash buffer for 5 min. 15. Mount cover slips on the slides using mounting medium. Store slides at room temperature in the dark. For longer periods, store at 4°C. 16. Photograph the green fluorescence of EGFP and red fluorescence on the same film using a double exposure. Examples of the photographs are shown in Fig. 18.2.

3.2

siRNA or shRNA-Mediated Inhibition of TGF-b1 in the Glomeruli

3.2.1 Preparation of siRNA and siRNA Expression Vector Targeting TGF-b1 3.2.1.1

siRNA

1. Dilute 50 µg of siRNA for TGF-β1 and scrambled gene solution in 500 µL nuclease-free PBS, respectively. Store the solution at room temperature until transfection and use within several hours (see Note 4).

3.2.1.2

Cloning Hairpin siRNA Inserts into siRNA Expression Vector

1. Dissolve each hairpin siRNA oligonucleotide (64 nt) in TE at 250 pmol/µL. 2. Annealing: assemble 5 µL of each of the sense and antisense hairpin siRNAencoding oligonucleotides in an Eppendorf tube (total 10 µL) and heat the mixture to 90°C for 10 min. 3. Add 2.5 µL of 5× annealing buffer and incubate at 90°C for 5 min.

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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 siRNA-transfected (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 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]) (see Color Plate 10)

4. Cool the solution slowly to room temperature. The annealed siRNA insert can be stored at -20°C for future ligation. 5. Dilute 1 µL of the annealed siRNA insert in 100 µL nuclease-free water. 6. Ligation: assemble 20 µL of ligation mix (1 µL of pSilencer vector [linearized, 0.1 mg/mL], 0.5 µL of the diluted annealed siRNA insert, 5 µL of 4× ligation buffer, 3.5 µL of nuclease-free water, and 10 µL of solution I) and incubate at 16°C for 1 h. 7. Transform 50 µL of transformation-competent E. coli cells with 5 µL of the ligation mix. 8. Plate cells onto LB plates containing 50–200 mg/mL ampicillin overnight at 37°C.

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9. Pick colonies, isolate the plasmid DNA, and use restriction digestion analysis to confirm the presence of the siRNA insert. 10. Purify the siRNA-expressing plasmid for transfection using the High Purity Maxiprep Plasmid Purification Kit. Store the plasmid solution at -20°C until transfection. 3.2.2

Induction of Thy-1 Nephritis

1. Anesthetize SD rats via intraperitoneal injection of pentobarbital (50 mg/kg). 2. Induce anti-Thy-1 glomerulonephritis by a single injection of 700 µg of mAb 1-22-3 via the tail vein (see Note 3). 3.2.3

Transfer of siRNA or siRNA Expression Vector Targeting TGF-b1 Via the Renal Artery

1. Twenty-four hours after induction of nephritis, transfect the siRNA (or siRNA expression vector) via the renal artery in accordance with the protocol described in Sect. 3.1.2 (see Note 4). 3.2.4

Assessment of TGF-b1 Expression in the Glomeruli

1. Four days after transfection, kill the rats. 2. After perfusion with 20 mL of ice-cold PBS via the abdominal aorta, remove the kidneys. 3. Collect the glomeruli in accordance with the conventional sieving technique using 125-, 180-, and 75-µm meshes. 4. Homogenize the collected glomeruli in 1 mL of lysis buffer using a tissue grinder on ice. Keep the lysate on ice for 20 min. 5. Centrifuge the lysate at 15,000 rpm (~20,000×g) for 20 min at 4°C. 6. Assay the protein concentration of the supernatant by using the BCA ΤΜ assay. 7. Mix the soluble lysates 1:2 with 3× Laemmli buffer and heat for 10 min at 95°C. 8. Load lysate onto a 15% SDS-PAGE gel (20 µg per lane), resolve, and transfer onto a polyvinylidene difluoride membrane. 9. Block membranes with 5% bovine serum albumin in TBST for 30 min at room temperature and then immunoblot with polyclonal antibodies against TGF-β1 in blocking buffer overnight at 4°C. 10. Wash membranes with TBST three times for 5 min each. 11. Incubate membranes with horseradish peroxidase-conjugated goat anti-rabbit IgG for 30 min. 12. Wash membranes with TBST three times for 5 min each. 13. Visualize membranes using chemiluminescence substrate in accordance with the manufacturer’s instructions. 14. Obtain X-ray films of the blots. An X-ray film for the siRNA expression vector is shown in Fig. 18.3b.

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Hind III

U6 promoter

sense

antisense

5⬘ GATCCCGTCAACTGTGGAGCAACACTTCAAGAGAGTGTTGCTCCACAGTTGACTTTTTTGGAAA 3⬘ 3⬘ GGCAGTTGACACCTCGTTGTGAAGTTCTCTCACAACGAGGTGTCAACTGAAAAAACCTTTTCGA 5⬘

a

loop pSU6TGF-β1

DC

N

TGF-β1 12.5kD pSU6Scramble

DC

N

TGF-β1 12.5kD b Fig. 18.3 shRNA-mediated silencing of TGF-β1 expression in vivo. a Construction of the siRNA expression vector targeting TGF-β1. To overcome the shortcomings of chemically synthesized siRNA, a DNA vector-mediated mechanism to express substrates that can be converted into siRNA was developed. Here, we used an siRNA expression vector featuring the human U6 RNA pol III promoter. Two oligodeoxynucleotides containing 21-nucleotide sense and antisense sequences of siRNA for TGF-β1, a 9-nucleotide loop sequence, and a transcription termination signal of six thymidines were annealed and inserted downstream of the U6 promoter. b After transferring this expression vector (200 µg) into rat kidney, we checked for shRNA-mediated silencing of TGF-β1 expression in the glomeruli of the anti-Thy-1 nephritis model using Western blot analysis. The siRNA expression vector for TGF-β1 (PSU6 TGF-β1) efficiently inhibited expression of TGF-β1 in vivo when compared with the contralateral diseased kidney (DC, diseased control) whereas the control vector (PSU6 scramble) did not. N, normal control. (Reproduced from ref. [3] with permission)

4

Notes

1. Rats weighing approximately 200 g are appropriate for this procedure because a catheter cannot be inserted via the renal artery in rats weighing less than 150 g, and the presence of too much fat can cause difficulties with operative precision in rats weighing more than 250 g. 2. Anti CD90 (Thy1.1) antibody that is applicable for immunohistochemistry studies is commercially available (e.g., Chemicon MAB1406). 3. Thy-1 nephritis can be reportedly induced using other commercially available mAbs (e.g., the OX-7 clone).

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4. In transferring siRNA or a DNA vector via the renal artery, we routinely use 50 µg of siRNA or 100–200 µg of a DNA vector. However our own data has shown that 5 µg of siRNA is sufficient in vivo [3]. As using a large amount of siRNA may produce an “off-target” effect, consider using a smaller amount of siRNA to minimize any nonspecific effects. 5. It is not necessary to peel off the peritoneum around the distal side of the left renal artery and the left kidney; doing so may lead to unexpected tissue damage. 6. Puncturing the renal artery directly will lead to unwanted ischemic change. 7. Keeping the bevel of the stylet downward will minimize the bleeding (Fig. 18.1b). 8. When the siRNA solution is infused with the catheter located at the distal site, siRNA will not be delivered to the whole kidney. 9. Successful hemostasis depends on any blood being completely removed. The operator and the assistant need to cooperate to ensure that this occurs. 10. Four-percent PFA can also be perfused after PBS perfusion (perfusion fixation), but the snap freezing (fresh freezing) method is not recommended because the EGFP is not fixed. 11. Immersion in the 10% and 20% sucrose solutions can be omitted without loss of quality.

References 1. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. (1998) Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature. 391, 806–811. 2. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T. (2001) Duplexes of 21nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature. 411, 494–498. 3. Takabatake Y, Isaka Y, Mizui Μ, Kawachi H, Shimizu F, Ito T, Hori Μ, Imai E. (2005) Exploring RNA interference as a therapeutic strategy for renal disease. Gene Ther. 12, 965–973. 4. Novina CD, Murray MF, Dykxhoorn DM, Beresford PJ, Riess J, Lee SK, Collman RG, Lieberman J, Shankar P, Sharp PA. (2002) siRNA-directed inhibition of HIV-1 infection. Nat Med. 8, 681–686. 5. Ito T, Suzuki A, Imai E, Okabe Μ, Hori Μ. (2001). Bone marrow is a reservoir of repopulating mesangial cells during glomerular remodeling. J Am Soc Nephrol. 12, 2625–2635. 6. Tsujie Μ, Isaka Y, Nakamura H, Imai E, Hori Μ. (2001) Electroporation-mediated gene transfer that targets glomeruli. J Am Soc Nephrol. 12, 949–954.

Index

A Actin cytoskeleton, 4, 205–206, 209–210, 217–218 Adenine nephrotoxicity, animal model, 47–48 Adriamycin nephrotoxicity, animal model, 46 Alkaline phosphatase, immunohistochemistry, 142–143 Aminonucleoside nephrosis, animal model, 46 Animal ethics, 41–42 Animal models, see Adenine nephrotoxicity, see also Adriamycin nephrotoxicity see also Aminonucleoside nephrosis, see also Anti-GBM nephritis, see also Cisplatin nephrotoxicity, see also Diabetes, experimental, see also Folic acid nephropathy, see also Gentamicin nephrotoxicity, see also Ischemic acute renal failure, see also Mercuric chloride nephrotoxicity, see also Obesity, see also Sub-total nephrectomy, see also Streptozotocin induced diabetes, see also Thy-1 nephritis, see also UUO Anti-GBM nephritis, 49–51, 50 (Fig. 4.3), 114 Antigen retrieval, tissue sections enzyme digestion,143, 168 microwave pretreatment,125, 143–144, 190 Anti-thymocyte serum nephritis, see Thy-1 nephritis APES coated slides, see silane coated slides Apoptosis, 4, 12, 176–181, 178 (Fig. 13.3) detection of, 175–192, 177 (Fig.13.1, 13.2) Autoradiography, 129–130 Avidin-Biotin complex, immunohistochemistry, 140–142, 134 (Fig. 10.1)

B Biomarkers, 237–249

C Caspase-3, marker of apoptosis,189 Cell cultures, see fibroblast, see also mesangial cells, see also myofibroblast, see also proximal tubules Cell death see Apoptosis, detection see also Necrosis Cell stretch, see Stretch, cell culture Cisplatin nephrotoxicity, animal model, 46–47 CM10 arrays, 241–243 Collagen, measurement of total collagen concentration, 226–229, 229 (Fig. 16.2) measurement of collagen sub-types, 230–232, 233 (Fig. 16.3) lattices, 193–203 Colloidal gold, preparation, 153–154 Confocal microscopy, see imaging, confocal Contraction, collagen lattices, 193–203 Counting, cell cultures, 198 Creatinine clearance, measurement in mice, 69 Cytochemistry, see Immunocytochemistry

D db/db mouse, 54 Diabetes, experimental models, 51–53, 51 (Fig. 4.4), 52 (Fig. 4.5), 56, 67 DNA fragmentation, see Apoptosis, detection of cDNA synthesis, 91–92 DNase treatment, Polymerase chain reaction, 78, 91 265

266 Dot Blot hybridization, 127–128 Double labelling, histochemistry, 142–143, 189–190

E Electron Microscopy immuno electron microscopy, 149–156 lectin histochemistry, 156–157 transmission electron microscopy, 176–178, 177 (Fig. 13.1b), 149–150, 150 (Fig. 11.1) Electrophoresis, 213, 231, 236 Electroporation, 256–258 Embedding of tissue, in paraffin wax, 139 in resin, 155 Enhanced green fluorescent protein (EGFP), 258–259, 260 (Fig. 18.2) Erk, 210 Expression vector, construction of vector for gene silencing, 259–261

F Fibroblast characterization, 32–35, 33 (Table 3.2, Fig. 3.2) culture, 31–35, 198 isolation, 30–31, 30 (Fig. 3.1) Fibronectin, Elisa assay, 218–219 immunohistochemical staining, 49 (Fig. 4.2) 5/6 Nephrectomy, see Sub-total nephrectomy, experimental model Fixation, collagen lattices, 200 electron microscopy,152, 185 general options,135–136, 139, 185–186 in situ hybridization,124–125 light microscopy, 135–136, 185–186 perfusion, 165–167, 164 (Fig. 12.3), 166 (Fig. 12.4) Fixatives, mercuric formalin,136 methyl carnoys,136 neutral buffered formalin,136 paraformaldehyde, 136, 163, 182 paraformaldehye-lysine-periodate (PLP), 152 Folic acid nephropathy, animal model, 48 Freezing, cultured cells, 11, 35, 211 tissue, see Tissue, freezing

Index G Gene expression, quantitative real time PCR,83–107 in situ hybridization, 119–132 Gene silencing, 251–263 Gentamicin nephrotoxicity, animal model, 47 Glomerular basement membrane nephritis, see Anti-GBM nephritis Glomerular filtration rate (GFR), measurement in mice, 61–72 Glomeruli, isolation of, 7–8 Glomerulonephritis, leukocyte infiltration, 109–110

H H&E, tissue stain, 186 H50 arrays, 241–243 Histochemistry, see Electron microscopy, Immuno electron microscopy see also Immunoflurorescence see also Immunohistochemistry see also In situ hybridization Histogene, tissue stain, 75 Housekeeping genes, 97 Hydrolysis, of probe, 126 Hydroxyproline, measurement, 226–229, 229 (Fig. 16.2) Hypoxia, in situ localization, 161–174

I IMAC30 arrays, 241–243 Imaging, confocal, 216–217, 217 (Fig. 15.2) in vivo, 109–117 Immunoblot, see western blotting Immunocytochemistry, 34–35 Immunofluorescence, 34–35, 189–190, 200–201, 253 Immunohistochemistry, paraffin emebedded tissue, 141–142, 141 (Fig. 10.2), 168, 200–201 controls for, 144 Inflammation, glomerular, 109–117 Intravital microscopy, see Imaging, in vivo In situ hybridization, 119–132 Inulin clearance, see Glomerular filtration rate Ischemic acute renal failure, animal model, 45

Index L Laser capture microdissection, 73–82, 76 (Fig. 6.1) Lectin histochemistry, see Lectin staining Lectin, staining electron microscopic level, 156–157 light microscopic level, 144–146, 145 (Fig. 10.3c,d) Leucocytes, 109–117

M Mass spectrometry, 245 (Fig. 17.1) Media, for cell culture fibroblast culture, 27–28, 196 mesangial cell culture, 5, 6 (Table 1.1), 7, 11, 206 proximal tubule culture, 20, 85, 89 Mercuric chloride nephrotoxicity, animal model, 47 Mesangial cell, commercial sources, 13 (Table 1.3) characterization, 10–11, 10 (Table 1.2) culture, 9–10, 11–13, 210–211 isolation, 6 (Table 1.1), 7–8 role in glmerulosclerosis, 4 stretch, 205–221 mRen–2 rat, 53 Multiplexing, 97–98 Myofibroblast, characteristics of mesangial cells, 3–4, 194 culture, 31–32 identification in culture, 33 (Table 3.2) identification in tissue, 145 (Fig. 10.3b)

N Necrosis, 176–181, 177 (Fig. 13.2b), 178 (Fig. 13.3)

O Obesity, animal models, 53–54 Osmotic mini-pumps, insertion and use of, 63 Oxygen gradients, in kidney, 162 (Fig. 12.2)

P PAN nephropathy, see Aminonucleoside nephrosis, animal model Percoll gradient, for isolation of proximal tubules, 21

267 Periodic acid-Schiff, tissue stain, 187 Pimonidazole, 161–174, 162 (Fig. 12.1) Polymerase chain reaction (PCR), see Quantitative real time PCR Primer design, polymerase chain reaction, 93–94 Protein, total protein (bradford) assay, 240 Protein-A gold, conjugation, 153–154 Protein-chip arrays, 242–244 Proteomics, 137–249 Proximal tubule, characterization, 23 culture, 22 isolation, 21 Puromycin aminonucleoside nephrosis, see Aminonucleoside nephrosis

Q Q10 arrays, 241–243 Quantitative real time PCR, 83–107

R Raf–1, activity assay, 214–215 Real-time PCR, see quantitative real time PCR Renal function, measurement, see Glomerular filtration rate, measurement Rho-A, activation assay, 215–216 Riboporbes, use in in situ hybridization, 119–132 RNA extraction from cultured cells, 90 from tissue, 77–78, 79, 88–89 RNA interference, see Small hairpin RNA (shRNA), see also Small interfering RNA (siRNA) cRNA probe labeling, Biotin labeling,127 Digoxigenin (DIG) labeling, 127 isotopic labeling, 126 RNA quantitation, 80, 90–91

S Sectioning, paraffin embedded tissue, 125, 140 SELDI-TOF, 237–249, 247 (Fig. 17.2) Small hairpin RNA (shRNA), 251–163 Small interfering RNA (siRNA), 251–263, 260 (Fig. 18.2) Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), 231–232, 233 (Fig. 16.3), 261

268 Silane coated slides, 125, 139–140 Spontaneously hypertensive rat (SHR), 53 Streptozotocin induced diabetes, animal model, 51–52, 51 (Fig. 4.4), 52 (Fig. 4.5), 67 strain susceptibility in mice, 53 (Fig. 4.6) Stretch, cell culture, 205–221 STZ diabetes, see Streptozotocin induced diabetes, Sub-total nephrectomy, experimental model, 44, 67

T Thawing, cultured cells, 11, 35, 211 Thy-1 nephritis, animal model, 4, 48–49, 49 (Fig. 4.2), 254 Tissue, fixation, see Fixatives freezing, 75 homogenization, 87–88, 239–240 Toluidine blue, stain for resin sections, 187

Index Transforming Growth Factor Beta effect on epithelial cells,100–101 inhibition of expression, 259–262, 262 (Fig. 18.3) TUNEL, 188-190 Tyramide, signal amplification of biotinylated probes, 131

U Unilateral ureteric obstruction, see UUO Urinary inulin clearance, see Glomerular filtration rate, measurement in mice UUO, 43, 112

W Wax, see Embedding of tissue Western blotting, 212–213, 212 (Fig. 15.1a), 261

Z Zucker rat, 54