Stem Cell Mobilization

METHODS

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MOLECULAR BIOLOGY™

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

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Stem Cell Mobilization Methods and Protocols

Edited by

Mikhail G. Kolonin and Paul J. Simmons Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA

Editors Mikhail G. Kolonin Institute of Molecular Medicine University of Texas Health Science Center at Houston Houston, TX, USA

Paul J. Simmons Institute of Molecular Medicine University of Texas Health Science Center at Houston Houston, TX, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-61779-942-6 ISBN 978-1-61779-943-3 (eBook) DOI 10.1007/978-1-61779-943-3 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2012943077 © Springer Science+Business Media, LLC 2012 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, 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 Humana Press is a brand of Springer Springer is part of Springer Science+Business Media (www.springer.com)

Preface Initially noted as a phenomenon accompanying the recovery of patients from myelosuppressive chemotherapy, stem cell mobilization represents a transient increase in the levels of circulating stem and progenitor cells. Observation of progenitor cell mobilization following the administration of certain factors, such as G-CSF, heralded a new era in hematological cell therapies with “mobilized blood” now essentially replacing bone marrow as the tissue of choice for hematopoietic reconstitution in cancer therapy. There is considerable interest in the phenomenon of mobilization in terms of understanding the underlying molecular mechanisms that drive the process of cell egress from the bone marrow. Recent studies have also revealed mobilization of progenitor cells from organs other than the bone marrow, although the importance of this phenomenon and the extent to which extramedullary cells contribute to the mobilized pool are yet to be understood. The notion that progenitor cell mobilization results in systemic redistribution of several cell populations that may participate in repair and regeneration has considerable clinical implications. While recruitment of systemically circulating stem cells may be beneficial in bone marrow reconstitution or wound healing settings, progenitor trafficking to lesions in cancer or other fibrotic conditions could have adverse effects. Therefore, development of reliable methods to quantify trafficking and read out activity of individual precursor cell types is highly important. This book aims to overview the current standing in cell mobilization methodology and to outline recent developments in the field for basic and biomedical research community. Specifically, clinical hematopoietic progenitor cell mobilization protocols and the experimental techniques used in animal models are covered in Chapters 1–11. The remaining part of the book addresses the frontiers in mobilization and analysis of non-hematopoietic progenitors, with specific emphases on endothelial progenitor cells (Chapters 12–14), mesenchymal progenitor cells (Chapters 15 and 20), monocyte-derived fibroblast progenitors (Chapter 16), and very small embryonic-like cells (Chapter 17). Advanced methodologies to analyze physiological and pathological functions of these distinct progenitor populations are also described (Chapters 14–15, 18–19). Houston, TX, USA

Mikhail G. Kolonin Paul J. Simmons

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Contents Preface ..................................................................................................................... Contributors............................................................................................................. 1 Mobilization of Hematopoietic Stem/Progenitor Cells: General Principles and Molecular Mechanisms................................................... Halvard Bonig and Thalia Papayannopoulou 2 Quantifying Hematopoietic Stem and Progenitor Cell Mobilization .................. Shiri Gur-Cohen, Kfir Lapid, and Tsvee Lapidot 3 Hematopoietic Stem Cell Mobilization with G-CSF .......................................... Chitra Hosing 4 Hematopoietic Stem Cell Mobilization with Agents Other than G-CSF ............. Jonathan Hoggatt and Louis M. Pelus 5 Hematopoietic Stem Cell Mobilization: A Clinical Protocol .............................. Gina Pesek and Michele Cottler-Fox 6 Monitoring Blood for CD34+ Cells to Determine Timing of Hematopoietic Progenitor Cells Apheresis..................................................... M. Louette Vaughn and Edmund K. Waller 7 Hematopoietic Progenitor Cell Collection ........................................................ S. Darlene Marlow and Myra House 8 Managing Apheresis Complications During the Hematopoietic Stem Cell Collection ......................................................................................... S. Darlene Marlow and Myra House 9 Hematopoietic Progenitor Cell Apheresis Processing ......................................... Eleanor S. Hamilton and Edmund K. Waller 10 Toxicities of Mobilized Stem Cell Infusion ........................................................ Jonathan L. Kaufman 11 Mobilization of Hematopoietic Stem Cells by Depleting Bone Marrow Macrophages ....................................................................................... Valérie Barbier, Ingrid G. Winkler, and Jean-Pierre Lévesque 12 Combinatorial Stem Cell Mobilization in Animal Models .................................. Simon C. Pitchford and Sara M. Rankin 13 Vascular Progenitor Cell Mobilization ............................................................... Kirsten A. Kienstra and Karen K. Hirschi 14 Evaluation of Circulating Endothelial Precursor Cells in Cancer Patients ........... Francesco Bertolini, Patrizia Mancuso, Liat Benayoun, Svetlana Gingis-Velitski, and Yuval Shaked

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15 Tracking Inflammation-Induced Mobilization of Mesenchymal Stem Cells ........ Erika L. Spaeth, Shannon Kidd, and Frank C. Marini 16 Differentiation of Circulating Monocytes into Fibroblast-Like Cells .................. Darrell Pilling and Richard H. Gomer 17 Enumeration of Very Small Embryonic-Like Stem Cells in Peripheral Blood ...... Rui Liu and Mariusz Z. Ratajczak 18 Generation of a Vascular Niche for Studying Stem Cell Homeostasis ................. Jason M. Butler and Shahin Rafii 19 Studying Vascular Progenitor Cells in a Neonatal Mouse Model ........................ Kirsten A. Kienstra and Karen K. Hirschi 20 Progenitor Cell Mobilization from Extramedullary Organs ................................ Mikhail G. Kolonin

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Index ................................................................................................................................ 253

Contributors VALÉRIE BARBIER • Mater Medical Research Institute, Aubigny Place, Raymond Terrace, South Brisbane, QLD, Australia LIAT BENAYOUN • Department of Molecular Pharmacology, Technion–Israel Institute of Technology, Haifa, Israel FRANCESCO BERTOLINI • Laboratory of Hematology-Oncology, European Institute of Oncology, Milan, Italy HALVARD BONIG • Department of Medicine/Hematology, University of Washington, Seattle, WA, USA JASON M. BUTLER • Weill Cornell Medical College and the Howard Hughes Medical Institute, Cornell University, New York, NY, USA SHIRI GUR-COHEN • Department of Immunology, Weizmann Institute of Science, Rehovot, Israel MICHELE COTTLER-FOX • Department of Pathology, University of Arkansas for Medical Sciences, Little Rock, AR, USA SVETLANA GINGIS-VELITSKI • Department of Molecular Pharmacology, Rappaport Faculty of Medicine, Technion–Israel Institute of Technology, Haifa, Israel RICHARD H. GOMER • Texas A&M University, College Station, TX, USA ELEANOR S. HAMILTON • Cellular Therapies Laboratory, Emory University Hospital, Atlanta, GA, USA KAREN K. HIRSCHI • Division of Neonatology, Department of Pediatrics, Baylor College of Medicine, Houston, TX, USA JONATHAN HOGGATT • Department of Microbiology and Immunology, Indiana University School of Medicine, Indianapolis, IN, USA CHITRA HOSING • Department of Stem Cell Transplantation and Cell Therapy, M.D. Anderson Cancer Center, Houston, TX, USA MYRA HOUSE • Center for Transfusion and Cellular Therapies (CTCT), Emory University Hospital, Atlanta, GA, USA JONATHAN L. KAUFMAN • Hematology and Medical Oncology, Winship Cancer Institute, Emory University School of Medicine, Atlanta, GA, USA SHANNON KIDD • Department of Leukemia, The University of Texas MD Anderson Cancer Center, Houston, TX, USA KIRSTEN A. KIENSTRA • Division of Neonatology, Department of Pediatrics, Baylor College of Medicine, Houston, TX, USA MIKHAIL G. KOLONIN • Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA KFIR LAPID • Department of Immunology, Weizmann Institute of Science, Rehovot, Israel TSVEE LAPIDOT • Department of Immunology, Weizmann Institute of Science, Rehovot, Israel JEAN-PIERRE LÉVESQUE • Mater Medical Research Institute, South Brisbane, QLD, Australia RUI LIU • Developmental Biology Program, James Graham Brown Cancer Center, University of Louisville Louisville, KY USA

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PATRIZIA MANCUSO • Laboratory of Hematology-Oncology, European Institute of Oncology, Milan, Italy FRANK C. MARINI • Institute for Regenerative Medicine, Comprehensive Cancer Center, Wake Forest University, Medical Center Blvd. Winston-Salem, NC S. DARLENE MARLOW • Center for Transfusion and Cellular Therapies, Emory University Hospital, Atlanta, GA, USA THALIA PAPAYANNOPOULOU • Department of Medicine/Hematology, University of Washington, Seattle, WA, USA LOUIS M. PELUS • Department of Microbiology and Immunology, Indiana University School of Medicine, Indianapolis, IN, USA GINA PESEK • Department of Pathology, University of Arkansas for Medical Sciences, Little Rock, AR, USA DARRELL PILLING • Texas A&M University, College Station, TX, USA SIMON C. PITCHFORD • Leukocyte Biology Section, Faculty of Medicine National Heart and Lung Institute, Imperial College London, London, UK SHAHIN RAFII • Weill Cornell Medical College and the Howard Hughes Medical Institute, Cornell University, New York, NY, USA SARA M. RANKIN • Leukocyte Biology Section, Faculty of Medicine National Heart and Lung Institute, Imperial College London, London, UK MARIUSZ Z. RATAJCZAK • Developmental Biology Program, James Graham Brown Cancer Center, University of Louisville, Louisville, KY, USA YUVAL SHAKED • Department of Molecular Pharmacology, Technion-Israel Institute of Technology, Haifa, Israel PAUL J. SIMMONS • Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA ERIKA L. SPAETH • Department of Leukemia, The University of Texas MD Anderson Cancer Center, Houston, TX, USA M. LOUETTE VAUGHN • Center for Transfusion and Cellular Therapies, Emory University Hospital, Atlanta, GA, USA EDMUND K. WALLER • Cellular Therapies Laboratory, Emory University Hospital, Atlanta, GA, USA INGRID G. WINKLER • Mater Medical Research Institute, Aubigny Place, Raymond Terrace, South Brisbane, QLD, Australia

Chapter 1 Mobilization of Hematopoietic Stem/Progenitor Cells: General Principles and Molecular Mechanisms Halvard Bonig and Thalia Papayannopoulou Abstract Hematopoietic stem/progenitor cell mobilization can be achieved by a variety of bone marrow niche modifications, although efficient mobilization requires simultaneous expansion of the stem/progenitor cell pool and niche modification. Many of the mechanisms involved in G-CSF-induced mobilization have been described. With regard to mobilization of hematopoietic stem/progenitor cells, challenges for the future include the analysis of genetic factors responsible for the great variability in mobilization responses, and the identification of predictors of mobilization efficiency, as well as the development of mobilizing schemes for poor mobilizers. Moreover, improved regimens for enhanced or even preferential mobilization of nonhematopoietic stem/progenitor cell types, and their therapeutic potential for endogenous tissue repair will be questions to be vigorously pursued in the near future. Key words: G-CSF, Mobilization, Hematopoietic stem/progenitor cell

1. Introduction Although mature hematopoietic cells are physiologically released from bone marrow to peripheral blood, their immature counterparts are found in circulation in very low frequencies. An enforced egress, referred to as “mobilization,” of a modest proportion of the latter cells from bone marrow to peripheral blood can be enacted by a variety of systemic “stressors.” Stem cell mobilization was uncovered mostly through empiric observations rather than rationally designed treatments. Why and how stem/progenitor cells physiologically escape the BM environment is not entirely clear, but it is very likely that the process of mobilization makes use of physiological molecular pathways leading to mobilization.

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_1, © Springer Science+Business Media, LLC 2012

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The considerable scientific interest in mobilization of immature cells is fuelled by its clinical relevance. Its importance in autologous repair mechanisms was demonstrated when after partial irradiation radiation-depleted marrow is repopulated from noncontiguous nonirradiated marrow sites, presumably by itinerant stem cells (1). Quantitatively, however, of greater clinical relevance at the current time, is the collection of mobilized cells by apheresis, enabling allogeneic transfer or temporary cryopreservation of autologous stem/progenitor cells for hematopoietic “stem cell” transplantation (2, 3). Protocols for several mobilization approaches are reported in this book and several recent comprehensive reviews have been published on clinical aspects or the cellular and molecular mechanisms of mobilization (4–8). This minireview focuses on issues relevant to G-CSF mobilization, because of its unique clinical importance and the plethora of studies on G-CSF mobilized cells. Mobilization by some other modalities is touched upon only because of their mechanistic insight and because they may display a synergistic or additive activity with G-CSF.

2. General Mobilization Principles

Under steady-state conditions, stem/progenitor cell location is almost exclusively restricted to the marrow, where these cells apparently reside in specific, supportive microenvironments (9–11). Environmental cues from stromal cells or matrix could influence cell fate, and are, under resting conditions, also responsible for their firm retention in the marrow. Active egress of stem/progenitor cells from bone marrow could be the default response when their restraining mechanisms are released, i.e., the HSPC could be inherently nomadic unless restrained. While this may appear to be a philosophical issue, the answer to this question could allow for a rational development of mobilizing agents. Currently available data on stem cell mobilization suggest that indeed the breakdown of retention mechanisms is sufficient for mobilization. Several common properties of mobilized hematopoietic cells have been emphasized irrespective of the mobilizing agent. Thus, mobilized immature cells are predominantly noncycling, in contrast to the cells left behind in the marrow (12–14), they express little VCAM-1, and low levels of many integrins (14–16). Specifically data generated with fast-acting mobilizing agents suggest that these phenotypic changes precede egress of cells from marrow, suggesting in turn that these properties are prerequisites for mobilization, rather than changes induced by the milieu in the peripheral blood (15). Likewise, gene expression patterns of mobilized immature subsets have been described; they differ markedly

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from their counterparts residing in unstimulated marrow (17, 18). Thus, in CD34+ cells from G-CSF mobilized blood, myeloid genes and cell cycle-associated genes were relatively up-regulated. These changes likely indicate differences in the heterogeneous mix of cells contained in the CD34+ fraction, entirely compatible with known effects of G-CSF, rather than necessarily pointing to molecular events involved in mobilization. In agreement with that, an extensive body of evidence has accumulated on differences in the ratio between primitive and more mature hematopoietic subsets, depending on the mobilizing agent. Thus, several publications have commented that AMD3100-alone mobilized immature cells are, on average, more functionally and phenotypically primitive than G-CSF- or G-CSF + AMD3100-mobilized ones (19, 20), resembling more closely the distribution in a steady-state marrow. This observation may be explained by the relative skewing of a G-CSF stimulated marrow towards less primitive (more mature) cells, i.e., the mobilized fractions are representative of marrow contents at the time of mobilization. As was reported many years ago, a G-CSF mobilized marrow is relatively depleted of immature hematopoietic subsets, and the marrow does not assume its normal cellular composition for several weeks after discontinuation of G-CSF (21). The precise locations from which mobilized immature cells originate, or the exact site of their egress, are not clear. A reasonable proposition is that egress into blood would require apposition to medullary blood vessels, most likely to medullary venous sinusoids. Mobilization by G-CSF is associated with a relative depletion of periosteal niches of hematopoietic stem cells, migration of stem cells to vascular niches where much of the proliferation occurs (5), followed by egress of both mature and immature subsets. With chemokine-induced mobilization the rapid kinetics likely do not allow for migration across significant distances, which may explain the relatively lower potency, and the synergism between G-CSF and AMD3100 (15). Of interest, data generated with the Gi protein inhibitor Pertussis toxin, which renders hematopoietic cells completely incapable of migration (22), clearly show that the ability to migrate is not a critical capacity of a mobilizable cell, i.e., that stem/progenitor cell pools might reside on the luminal side of medullary blood vessels. In aggregate, these data may indicate that although mobilized cells would at some time cross perivascular pools before they exit the bone marrow, they could initially originate from other bone marrow locations further away and that such movement to the perivascular space increases the number of mobilizable cells. How large is the fraction of stem cells that can be induced to leave the marrow? At first glance, extrapolation from the mouse model indicates that the efficiency of mobilization with G-CSF might be modest. After a similar mobilization regime as in humans

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(nine doses q12h) several thousand CFU-C per mL of blood, i.e., no more than 10,000 CFU-C in total, will be in circulation. The number of circulating CFU-C after G-CSF in the mouse (10,000 in a C57Bl/6 mouse, more in some other strains) must be compared against a CFU-C content of approximately 60,000 per steady-state (unmobilized) femur (15), which is estimated to represent 1/16 of the total marrow mass (23). Thus, the total number of CFU-C of the mouse is one million, 100 times the number that is found in circulation after G-CSF. However, this may not necessarily mean that only 1% of “stem cells” are mobilized by G-CSF, since other relevant variables in the equation are completely unfathomable. The transit time of mobilized cells is elusive (minutes to a few hours have been suggested) (24, 25), and their fate has not been completely elucidated. In other words, once the cells are in circulation, how long do they remain there, and when they leave the circulation, how many home back to marrow or are lost to other organs is unclear. Conceivably, many circulating cells could interact with and be siphoned off by nontarget organs. In that case the true number of mobilized cells would be much higher than the number in circulation suggests. What is the evidence for such “steal” effects? Experimental evidence has been provided that the spleen of a G-CSF mobilized mouse accumulates significant numbers of immature cells, so that mobilization of splenectomized mice is more pronounced (26). Trafficking of mobilized immature cells through the intestinal lymphoid system and to adipose tissue has also been shown (27, 28). The possibility that this likewise pertains to other organs must be entertained. Since unlike the spleen, other organs do not support immature hematopoietic cells, it is difficult to experimentally address this issue with currently available technology, but again, tracking experiments in transplanted animals demonstrate accumulation of progenitor cells in nonhematopoietic tissues (29). Thus, reliable estimates of the potency of G-CSF mediated mobilization cannot be given. How is mobilization quantified? Ultimately, the cell of interest in the context of hematopoietic cell mobilization is the stem cell. Yet the stem cell is defined functionally, as a cell capable of selfrenewal and long-term multilineage reconstitution in an appropriately conditioned host. It must be remembered that any other “stem cell” enumeration assay than the long-term engraftment assay (30) is measuring some surrogate parameter, so many caveats must be considered when interpreting the results of such assays. In addition to its tediousness, even a stem cell assay (limiting dilution transplantation and readout of long-term engraftment) has its limitations, since it tests at the same time stemness and transplantrelated properties like homing, niche-integration, retention, etc. Thus, if cells which would be capable of self-renewal and longterm repopulation in terms of their epigenetic status, i.e., are bona fide stem cells, are impaired in their ability to interact with the

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niche or to proliferate, the number of stem cells might be underestimated. In vitro colony assays have been used by most to quantify mobilization, or to compare mobilization efficiency (30–33). Since cells giving rise to colonies in colony assays are progenitor cells, i.e., more mature specimen, CFU-assays are not a true measure of stem cell mobilization. However, cumulative evidence indicates that during progenitor cell mobilization stem cells are always comobilized, but the relative frequency among the immature cells may vary, depending on the mobilizing agent. Thus, the CFU-C assay may be the most practicable assay for assessment of mobilization, but its shortcomings must be born in mind. Phenotypic analyses of “stem cells” using more or less complex surface marker panels have also been used. These assays are most problematic, because mobilizing agents can induce changes in surface phenotype (e.g., c-kit expression on immature cells is all but suppressed on G-CSF mobilized cells); thus, the stem cell phenotype of a stem cell in a steady-state marrow is likely different from that in mobilized peripheral blood (34, 35). With less complex surface marker panels (e.g., CD45/CD34), the relative mix between primitive and more mature subsets contained in this phenotypically defined, yet functionally heterogeneous population is not considered and can lead to misinterpretations of stem cell mobilization efficiency.

3. Mobilization by G-CSF The clinically most relevant mobilizing agent, G-CSF, expands the number of stem cells at the same time that it induces proliferation/ maturation towards the granulocytic lineage, and it causes marked alterations in the hematopoietic stroma in the marrow. Together these changes result in the release, or mobilization, of hematopoietic stem/progenitor cells. It seems clear that the summation of expansion and mobilization is responsible for the rather potent mobilization efficiency of G-CSF compared to other mobilizing agents. In humans, after a conventional course of G-CSF (5 μg/kg every 12 h, nine total doses) the number of circulating progenitor cells is increased approximately 60-fold, to 60–100 CD34+ cells/μL. Preliminary data indicate that other types of immature cells are comobilized alongside hematopoietic stem/progenitor cells, including endothelial and mesenchymal stroma cells (36–38). It is not unreasonable to hypothesize that the same changes which cause hematopoietic stem/progenitor cell mobilization are also involved in mobilization of these other stem cell specimen, but definitive data are lacking.

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Recently, significant advances have been made with respect to the molecular and cellular events involved in G-CSF mediated hematopoietic stem/progenitor cell mobilization. Informative data document that mobilization is not a direct effect of G-CSF on the stem cell proper. The receptor for G-CSF, G-CSFR, is conspicuously absent from hematopoietic stem cells (39). Indirect cues must therefore be responsible for numeric and spatial changes in the stem cell population. Recent data by the Link and Levesque laboratories suggest a chain of events starting with G-CSF mediated stimulation of certain marrow-resident macrophages which appear to relay signals to osteoblasts (also G-CSFR negative), which then downregulate SDF-1 gene transcripts (40–42). Proteolytic cleavage of SDF-1 off of stromal binding sites has also been demonstrated and functionally implicated, as truncation of SDF-1, resulting in nonfunctional SDF-1 molecules can compete with full-length SDF-1 for CXCR4 binding sites. The cellular and molecular architecture of a G-CSF treated bone marrow is significantly changed compared to a steady-state marrow. For instance, cleavage of a number of surface-bound chemokines, cytokines, receptors, etc. has been demonstrated. Some evidence has been provided that these changes are the work of proteases, which are elaborated during G-CSF mobilization, together with down regulation of protease-inhibitors during mobilization. However, the critical role of MMP9 emphasized in some studies (43) has not been confirmed, and even mice deficient in a whole panel of proteases responded to G-CSF with the expected efficiency (44). Further, deficiency in CD26 is associated with impaired mobilization by G-CSF (45). CD26 is a broad dipeptidase that (among many other putative target molecules) cleaves SDF-1 into a nonfunctional variant, which competes with SDF-1 for CXCR4 binding. It was proposed that the inability to cleave SDF-1 was responsible for the attenuated G-CSF responsiveness of the CD26deficient mice (45). At this point in time, a definitive contributory role of other proteases to mobilization cannot be pinpointed. As the marrow is exposed to G-CSF and the described profound changes in marrow architecture are happening, HSPC expand in regions located more centrally and closer to the blood vessel. Data from the Levesque laboratory suggest that this is at least in part a reflection of (a) greater oxygen needs of proliferating cells and (b) greater oxygen consumption in a proliferating marrow, i.e., during G-CSF stimulation, HSPC move towards higher oxygen concentrations (46). The potent mobilizing activity of certain chemotherapy drugs like cyclophosphamide has been solely attributed to endogenous G-CSF, since G-CSFR deficient mice treated with cyclophosphamide show the expected rebound proliferation in marrow, but egress of immature cells from marrow is virtually absent (47). The mechanisms involved in mobilization by cyclophosphamide would then likely be the same as during mobilization with exogenous G-CSF.

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Several cytokines (including GM-CSF, FLT3 ligand and SCF, the c-kit ligand) mobilize HSPC and synergize with G-CSF in stem cell mobilization (47–50). These modalities have in common similarly slow kinetics as G-CSF mobilization, suggesting a combined effect of proliferation and mobilization, as with G-CSF. Some clinical data with GM-CSF and SCF, the latter predominantly in combination with G-CSF, have been reported, but their clinical relevance is modest. The mechanisms of mobilization with these cytokines have not been studied in any detail. Considering the role of the coagulation/complement cascade in hematopoietic cell trafficking (51–53) and the strong activation of this system by GM-CSF (54), a contribution of this pathway is conceivable. A different group of mobilizing agents has gained a lot of attention in the last few years, namely, CXCR4 antagonists of various chemistries. The effectiveness of this intervention has been demonstrated in mice, monkeys, dogs and humans. One CXCR4 antagonist, the bicyclam AMD3100 (Mozobil, Plerixafor) is licensed for clinical mobilization in combination with G-CSF + chemotherapy for patients failing to adequately mobilize with G-CSF + chemotherapy alone (15, 31, 55–60). Preliminary data from the Di Persio laboratory, reported at the 2010 ISBT meeting, indicate that when given alone, as with G-CSF, other species of immature cells are also comobilized by CXCR4 antagonists, although their nature has not been definitively elucidated. The mechanism of action of CXCR4 antagonists appears to be interference between stromal SDF-1 and CXCR4 on the HSPC surface. The kinetics is rapid, quite unlike those of G-CSF, and no conclusive evidence has been provided that these CXCR4 inhibitors elicit changes in the hematopoietic niche. Proliferation is not a feature of mobilization with CXCR4 antagonists, which was thought to explain the relatively low potency of these inhibitors. Preliminary data from our group indicates, however, that novel, more potent CXCR4 inhibitors can exceed the mobilization achieved with a 5-day course of twice-daily G-CSF, at least in mice (unpublished data). The frequency of stem cells in CXCR4-antagonist mobilized grafts among the cells with an immature phenotype is greater than after G-CSF. It appears that this reflects the frequencies within a steady-state marrow as opposed to a G-CSF-treated marrow, so this observation should not be surprising. Clearly mobilization with CXCR4 antagonists argues against a hypothesis put forth about mechanisms of G-CSF mobilization, i.e., inversion of an SDF-1 gradient, where mobilization of HSPC would be in response to greater concentrations outside the marrow than inside (supposedly because SDF-1 is cleaved from the stroma, to circulate in blood and bone marrow fluid). Because

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of the very short half-life of SDF-1 in plasma, this seemed unlikely, but definitive evidence against this hypothesis comes from the following observation: since CXCR4 antagonists effectively block SDF-1 directed migration, yet potently synergize with (rather than antagonize) G-CSF mobilization (15), it is clear that in mobilization cells do not respond to SDF-1, but are on the contrary made temporarily unresponsive. Data about mobilization with CXCR4 agonists are in line with this hypothesis (61), since they lead to downregulation of CXCR4 surface expression on HSPC. Thus, the SDF-1-CXCR4 axis acts as a retention pathway, which is disturbed by various means in mobilization with G-CSF and CXCR4 antagonists or agonists. Data generated in mice transplanted with CXCR4 deficient hematopoietic cells are in agreement with these observations (62). Considering that several authors have proposed interference of the CXCR4/SDF-1 pathway as the mechanism of action of G-CSF mediated mobilization, the well-documented synergism between G-CSF and CXCR4 antagonists is surprising. The simplest explanation may be (1) that after G-CSF, this pathway is only partially obstructed and (2) that CXCR4 antagonists may find a larger population to mobilize in a G-CSF treated marrow than in an untreated one, because of expansion of the pool and of relocation to perivascular regions. The inhibitor of Gi protein signals, including of SDF-1/ CXCR4 signals, Pertussis toxin was reported to elicit potent and protracted HSPC mobilization. Specifically, Pertussis toxin also synergized with G-CSF induced mobilization (22). Why Gi protein blockade leads to mobilization is not clear. While irrelevant from a clinical perspective, the implications for mobilization mechanics are of interest. Pertussis toxin mobilized HSPC are incapable of migration. This suggests that activation of migratory signals may not be required for mobilization. Similarly, GRO-β induced mobilization is seen despite the fact that it inhibits migration in vitro (32). These data could be interpreted to indicate that mobilizable pools of HSPC reside not in the marrow immediately adjacent to bone surfaces, CAR cells and other stromal elements, but in regions adjacent to the venous sinuses in marrow. Such a location would be equally compatible with the rapid kinetics of IL-8, GRO-β and CXCR4 antagonists—in either case distant transmarrow migration might not be feasible within the relevant time frame. Fenestrae in the septum segregating the spaces between the marrow space and the venous sinuses have been described as the site of passage of mature cells into blood. Conceivably, these fenestrae could also be used by immature hematopoietic cells during mobilization. This hypothesis would also be compatible with the observation that HSPC in G-CSF treated marrow are preferentially located close to blood vessels (46).

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VLA4 is an adhesion molecule expressed on HSPC. It is normally present in its low-affinity conformation, but affinity can be induced by a number of cytokines, and it changes during cell cycle transit (14). Like CXCR4 and SDF-1, VLA4 finds cognate ligands in the marrow stroma, including, but potentially not limited to, VCAM1, fibronectin, and osteopontin, thus serving as a stem cell retention pathway. Studies in mice, nonhuman primates or humans, using genetic, small-molecule- or antibody-mediated ablation of VLA4 or of several of its ligands in marrow have demonstrated mobilization of HSPC into peripheral blood (31, 63–67). The kinetics follow an intermediate time course. Although reduced expression of VLA4 is also a feature of G-CSF mobilized HSPC (14–16), suggesting VLA4 downregulation as another mechanism of G-CSF mobilization, VLA4 inhibition or genetic deletion was synergistic or at least superadditive with G-CSF. Similarly to what we postulated for synergism between CXCR4 antagonists and G-CSF, interference of G-CSF induced events with VLA4-mediated adhesion is likely incomplete, while direct targeting of the molecule is complete. As we have shown, VLA4 blockade is effective at mobilizing HSPC in mice, monkeys, and humans, albeit with low potency (31, 64, 68). As an indication that VLA4-inhibition and CXCR4 blockade are mobilizing HSPC by independent mechanisms, we and others have demonstrated synergism of the two modalities in monkeys and mice (31, 67). Lower VLA4 expression was also observed on HSPC mobilized with CXCR4 antagonists and with a variety of other mobilizing agents (15). This could either indicate downregulation of VLA4 under the influence of mobilizing agents and preferential mobilization of these VLA4dim cell populations, or assumption of a VLA4-dim phenotype during the transition from marrow to blood. Two other chemokines, GRO-β and IL-8, mobilize HSPC with very rapid kinetics. With respect to GRO-β, it was shown that this mobilization was dependent on MMP9, suggesting that the target cell may be a mature neutrophil (32, 69). For IL-8, contradictory results about the role of MMP9 have been reported, yet a dependence on G-CSFR likewise suggests a role for mediators released from mature neutrophils (47, 70, 71). The proposed chain of events leading to mobilization for these two molecules is granulocyte degranulation, release of proteases, severing of retention factors, resulting in stem cell release. Several other examples of stroma or niche modification have also been reported which resulted in stem cell mobilization. Several of these involved modification of stromal ligands for established retention factors. Examples include very different substances, such as Fucoidan, which displays competitive displacement of chemokines, including SDF-1 (which is present in the stem cell niche as a surface-bound molecule) (72, 73), anti-VCAM-1 antibodies (74)

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and VCAM-1-deleted (75) or osteopontin-deleted (33) mice, which blocked/ablated relevant VLA4 ligands in the stroma. Other mobilizing agents likely exerted their effects indirectly, through induction of endogenous G-CSF (e.g., parathyroid hormone) (76). Moreover, a variety of mediators have been associated with mobilization that seemingly have very little in common, and where the molecular mechanics are sometimes poorly defined. These include sympathomimetics (77, 78), cannabinoid receptor agonists (79), complement (52, 53), elevated lipoprotein levels (80), defibrotide (81), glycosaminoglycans (82), and endotoxin (83). None of these have gained any clinical relevance, but the abundance of mobilizing agents indicates the precariousness of the equilibrium between marrow retention and egress, and may in the future support the rational development of mobilizing strategies for poorly mobilizing patients, or for individuals who are intolerant to G-CSF. References 1. Nothdurft W, Kreja L (1998) Hemopoietic progenitor cells in the blood as indicators of the functional status of the bone marrow after totalbody and partial-body irradiation: experiences from studies in dogs. Stem Cells 16(Suppl 1): 97–111 2. Holig K, Kramer M, Kroschinsky F, Bornhauser M, Mengling T, Schmidt AH et al (2009) Safety and efficacy of hematopoietic stem cell collection from mobilized peripheral blood in unrelated volunteers: 12 years of single-center experience in 3928 donors. Blood 114:3757–3763 3. Chao NJ, Schriber JR, Grimes K, Long GD, Negrin RS, Raimondi CM et al (1993) Granulocyte colony-stimulating factor “mobilized” peripheral blood progenitor cells accelerate granulocyte and platelet recovery after high-dose chemotherapy. Blood 81:2031–2035 4. Levesque JP, Winkler IG (2008) Mobilization of hematopoietic stem cells: state of the art. Curr Opin Organ Transplant 13:53–58 5. Levesque JP, Helwani FM, Winkler IG (2010) The endosteal ‘osteoblastic’ niche and its role in hematopoietic stem cell homing and mobilization. Leukemia [E-pub 2010 Sep 23] 6. Papayannopoulou T, Scadden DT (2008) Stem-cell ecology and stem cells in motion. Blood 111:3923–3930 7. Pelus LM (2008) Peripheral blood stem cell mobilization: new regimens, new cells, where do we stand. Curr Opin Hematol 15:285–292 8. Greenbaum AM, Link DC (2010) Mechanisms of G-CSF-mediated hematopoietic stem and progenitor mobilization. Leukemia [E-pub 2010 Nov 16]

9. Lymperi S, Ferraro F, Scadden DT (2010) The HSC niche concept has turned 31. Has our knowledge matured? Ann N Y Acad Sci 1192: 12–18 10. Oh IH, Kwon KR (2010) Concise review: multiple niches for hematopoietic stem cell regulations. Stem Cells 28:1243–1249 11. Mendez-Ferrer S, Michurina TV, Ferraro F, Mazloom AR, Macarthur BD, Lira SA et al (2010) Mesenchymal and haematopoietic stem cells form a unique bone marrow niche. Nature 466:829–834 12. Scott MA, Apperley JF, Bloxham DM, Jestice HK, John S, Marcus RE et al (1997) Biological properties of peripheral blood progenitor cells mobilized by cyclophosphamide and granulocyte colony-stimulating factor. Br J Haematol 97:474–480 13. Williams CD, Linch DC, Watts MJ, Thomas NS (1997) Characterization of cell cycle status and E2F complexes in mobilized CD34+ cells before and after cytokine stimulation. Blood 90:194–203 14. Yamaguchi M, Ikebuchi K, Hirayama F, Sato N, Mogi Y, Ohkawara J et al (1998) Different adhesive characteristics and VLA-4 expression of CD34(+) progenitors in G0/G1 versus S + G2/M phases of the cell cycle. Blood 92:842–848 15. Bonig H, Chudziak D, Priestley G, Papayannopoulou T (2009) Insights into the biology of mobilized hematopoietic stem/ progenitor cells through innovative treatment schedules of the CXCR4 antagonist AMD3100. Exp Hematol 37:402–415

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16. Prosper F, Stroncek D, McCarthy JB, Verfaillie CM (1998) Mobilization and homing of peripheral blood progenitors is related to reversible downregulation of alpha4 beta1 integrin expression and function. J Clin Invest 101: 2456–2467 17. Graf L, Heimfeld S, Torok-Storb B (2001) Comparison of gene expression in CD34+ cells from bone marrow and G-CSF-mobilized peripheral blood by high-density oligonucleotide array analysis. Biol Blood Marrow Transplant 7:486–494 18. Steidl U, Kronenwett R, Rohr UP, Fenk R, Kliszewski S, Maercker C et al (2002) Gene expression profiling identifies significant differences between the molecular phenotypes of bone marrow-derived and circulating human CD34+ hematopoietic stem cells. Blood 99:2037–2044 19. Fruehauf S, Veldwijk MR, Seeger T, Schubert M, Laufs S, Topaly J et al (2009) A combination of granulocyte-colony-stimulating factor (G-CSF) and plerixafor mobilizes more primitive peripheral blood progenitor cells than G-CSF alone: results of a European phase II study. Cytotherapy 11:992–1001 20. Broxmeyer HE, Orschell CM, Clapp DW, Hangoc G, Cooper S, Plett PA et al (1995) Rapid mobilization of murine and human hematopoietic stem and progenitor cells with AMD3100, a CXCR4 antagonist. J Exp Med 201:1307–1318 21. Drize N, Chertkov J, Samoilina N, Zander A (1996) Effect of cytokine treatment (granulocyte colony-stimulating factor and stem cell factor) on hematopoiesis and the circulating pool of hematopoietic stem cells in mice. Exp Hematol 24:816–822 22. Papayannopoulou T, Priestley GV, Bonig H, Nakamoto B (2003) The role of G-protein signaling in hematopoietic stem/progenitor cell mobilization. Blood 101:4739–4747 23. Katayama Y, Hidalgo A, Furie BC, Vestweber D, Furie B, Frenette PS (2003) PSGL-1 participates in E-selectin-mediated progenitor homing to bone marrow: evidence for cooperation between E-selectin ligands and alpha4 integrin. Blood 102:2060–2067 24. Raghavachar A, Steinbach KH, Prummer O, Grilli G, Fliedner TM (1983) Survival of transfused cryopreserved granulocytic progenitor cells (CFU-C) in recipient circulation. Cell Tissue Kinet 16:303–311 25. Wright DE, Wagers AJ, Gulati AP, Johnson FL, Weissman IL (2001) Physiological migration of hematopoietic stem and progenitor cells. Science 294:1933–1936 26. Molineux G, Pojda Z, Dexter TM (1990) A comparison of hematopoiesis in normal and

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splenectomized mice treated with granulocyte colony-stimulating factor. Blood 75:563–569 27. Han J, Koh YJ, Moon HR, Ryoo HG, Cho CH, Kim I et al (2010) Adipose tissue is an extramedullary reservoir for functional hematopoietic stem and progenitor cells. Blood 115:957–964 28. Massberg S, Schaerli P, Knezevic-Maramica I, Kollnberger M, Tubo N, Moseman EA et al (2007) Immunosurveillance by hematopoietic progenitor cells trafficking through blood, lymph, and peripheral tissues. Cell 131:994–1008 29. Watanabe T, Kajiume T, Takaue Y, Kawano Y, Kanamaru S, Okamura S et al (2001) Decrease in circulating hematopoietic progenitor cells by trapping in the pulmonary circulation. Cytotherapy 3:461–466 30. Szilvassy SJ, Humphries RK, Lansdorp PM, Eaves AC, Eaves CJ (1990) Quantitative assay for totipotent reconstituting hematopoietic stem cells by a competitive repopulation strategy. Proc Natl Acad Sci USA 87:8736–8740 31. Bonig H, Watts KL, Chang KH, Kiem HP, Papayannopoulou T (2009) Concurrent blockade of alpha4-integrin and CXCR4 in hematopoietic stem/progenitor cell mobilization. Stem Cells 27:836–837 32. Fukuda S, Bian H, King AG, Pelus LM (2007) The chemokine GRObeta mobilizes early hematopoietic stem cells characterized by enhanced homing and engraftment. Blood 110:860–869 33. Grassinger J, Haylock DN, Storan MJ, Haines GO, Williams B, Whitty GA et al (2009) Thrombin-cleaved osteopontin regulates hemopoietic stem and progenitor cell functions through interactions with alpha9beta1 and alpha4beta1 integrins. Blood 114:49–59 34. Randall TD, Weissman IL (1997) Phenotypic and functional changes induced at the clonal level in hematopoietic stem cells after 5-fluorouracil treatment. Blood 89:3596–3606 35. Rollini P, Faes-Van’t HE, Kaiser S, Kapp U, Leyvraz S (2007) Phenotypic and functional analysis of human fetal liver hematopoietic stem cells in culture. Stem Cells Dev 16:281–296 36. Kassis I, Zangi L, Rivkin R, Levdansky L, Samuel S, Marx G et al (2006) Isolation of mesenchymal stem cells from G-CSF-mobilized human peripheral blood using fibrin microbeads. Bone Marrow Transplant 37:967–976 37. Tatsumi K, Otani H, Sato D, Enoki C, Iwasaka T, Imamura H et al (2008) Granulocyte-colony stimulating factor increases donor mesenchymal stem cells in bone marrow and their mobilization into peripheral circulation but does not repair dystrophic heart after bone marrow transplantation. Circ J 72:1351–1358

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38. Zubair AC, Malik S, Paulsen A, Ishikawa M, McCoy C, Adams PX et al (2010) Evaluation of mobilized peripheral blood CD34(+) cells from patients with severe coronary artery disease as a source of endothelial progenitor cells. Cytotherapy 12:178–189 39. Liu F, Poursine-Laurent J, Link DC (2000) Expression of the G-CSF receptor on hematopoietic progenitor cells is not required for their mobilization by G-CSF. Blood 95:3025–3031 40. Semerad CL, Christopher MJ, Liu F, Short B, Simmons PJ, Winkler I et al (2005) G-CSF potently inhibits osteoblast activity and CXCL12 mRNA expression in the bone marrow. Blood 106:3020–3027 41. Winkler IG, Sims NA, Pettit AR, Barbier V, Nowlan B, Helwani F, et al (2010) Bone marrow macrophages maintain hematopoietic stem cell (HSC) niches and their depletion mobilizes HSC. Blood [E-pub 2010 Aug 16] 42. Christopher MJ, Liu F, Hilton MJ, Long F, Link DC (2009) Suppression of CXCL12 production by bone marrow osteoblasts is a common and critical pathway for cytokine-induced mobilization. Blood 114:1331–1339 43. Heissig B, Hattori K, Dias S, Friedrich M, Ferris B, Hackett NR et al (2002) Recruitment of stem and progenitor cells from the bone marrow niche requires MMP-9 mediated release of kit-ligand. Cell 109:625–637 44. Levesque JP, Liu F, Simmons PJ, Betsuyaku T, Senior RM, Pham C et al (2004) Characterization of hematopoietic progenitor mobilization in protease-deficient mice. Blood 104:65–72 45. Christopherson KW, Cooper S, Hangoc G, Broxmeyer HE (2003) CD26 is essential for normal G-CSF-induced progenitor cell mobilization as determined by CD26−/− mice. Exp Hematol 31:1126–1134 46. Winkler IG, Barbier V, Wadley R, Zannettino AC, Williams S, Levesque JP (2010) Positioning of bone marrow hematopoietic and stromal cells relative to blood flow in vivo: serially reconstituting hematopoietic stem cells reside in distinct nonperfused niches. Blood 116: 375–385 47. Liu F, Poursine-Laurent J, Link DC (1997) The granulocyte colony-stimulating factor receptor is required for the mobilization of murine hematopoietic progenitors into peripheral blood by cyclophosphamide or interleukin-8 but not flt-3 ligand. Blood 90:2522–2528 48. Brasel K, McKenna HJ, Charrier K, Morrissey PJ, Williams DE, Lyman SD (1997) Flt3 ligand synergizes with granulocyte-macrophage colony-stimulating factor or granulocyte colony-stimulating factor to mobilize

hematopoietic progenitor cells into the peripheral blood of mice. Blood 90:3781–3788 49. Gianni AM, Siena S, Bregni M, Tarella C, Stern AC, Pileri A et al (1989) Granulocytemacrophage colony-stimulating factor to harvest circulating haemopoietic stem cells for autotransplantation. Lancet 2:580–585 50. Molineux G, Migdalska A, Szmitkowski M, Zsebo K, Dexter TM (1991) The effects on hematopoiesis of recombinant stem cell factor (ligand for c-kit) administered in vivo to mice either alone or in combination with granulocyte colony-stimulating factor. Blood 78:961–966 51. Ratajczak J, Reca R, Kucia M, Majka M, Allendorf DJ, Baran JT et al (2004) Mobilization studies in mice deficient in either C3 or C3a receptor (C3aR) reveal a novel role for complement in retention of hematopoietic stem/progenitor cells in bone marrow. Blood 103:2071–2078 52. Wilschut IJ, Erkens-Versluis ME, Ploemacher RE, Benner R, Vos O (1979) Studies on the mechanism of haemopoietic stem cell (CFUs) mobilization. A role of the complement system. Cell Tissue Kinet 12:299–311 53. Molendijk WJ, van Oudenaren A, van Dijk H, Daha MR, Benner R (1986) Complement split product C5a mediates the lipopolysaccharideinduced mobilization of CFU-s and haemopoietic progenitor cells, but not the mobilization induced by proteolytic enzymes. Cell Tissue Kinet 19:407–417 54. Bonig H, Burdach S, Gobel U, Nurnberger W (2001) Growth factors and hemostasis: differential effects of GM-CSF and G-CSF on coagulation activation—laboratory and clinical evidence. Ann Hematol 80:525–530 55. Burroughs L, Mielcarek M, Little MT, Bridger G, Macfarland R, Fricker S et al (2005) Durable engraftment of AMD3100-mobilized autologous and allogeneic peripheral-blood mononuclear cells in a canine transplantation model. Blood 106:4002–4008 56. Larochelle A, Krouse A, Metzger M, Orlic D, Donahue RE, Fricker S et al (2006) AMD3100 mobilizes hematopoietic stem cells with longterm repopulating capacity in nonhuman primates. Blood 107:3772–3778 57. Devine SM, Vij R, Rettig M, Todt L, McGlauchlen K, Fisher N et al (2008) Rapid mobilization of functional donor hematopoietic cells without G-CSF using AMD3100, an antagonist of the CXCR4/SDF-1 interaction. Blood 112:990–998 58. Devine SM, Flomenberg N, Vesole DH, Liesveld J, Weisdorf D, Badel K et al (2004) Rapid mobilization of CD34+ cells following administration of the CXCR4 antagonist

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AMD3100 to patients with multiple myeloma and non-Hodgkin’s lymphoma. J Clin Oncol 22:1095–1102 59. DiPersio JF, Stadtmauer EA, Nademanee A, Micallef IN, Stiff PJ, Kaufman JL et al (2009) Plerixafor and G-CSF versus placebo and G-CSF to mobilize hematopoietic stem cells for autologous stem cell transplantation in patients with multiple myeloma. Blood 113:5720–5726 60. Liles WC, Broxmeyer HE, Rodger E, Wood B, Hubel K, Cooper S et al (2003) Mobilization of hematopoietic progenitor cells in healthy volunteers by AMD3100, a CXCR4 antagonist. Blood 102:2728–2730 61. Hattori K, Heissig B, Tashiro K, Honjo T, Tateno M, Shieh JH et al (2004) Plasma elevation of stromal cell-derived factor-1 induces mobilization of mature and immature hematopoietic progenitor and stem cells. Blood 97:3354–3360 62. Foudi A, Jarrier P, Zhang Y, Wittner M, Geay JF, Lecluse Y et al (2006) Reduced retention of radioprotective hematopoietic cells within the bone marrow microenvironment in CXCR4−/− chimeric mice. Blood 107:2243–2251 63. Craddock CF, Nakamoto B, Elices M, Papayannopoulou T (1997) The role of CS1 moiety of fibronectin in VLA mediated haemopoietic progenitor trafficking. Br J Haematol 97:15–21 64. Craddock CF, Nakamoto B, Andrews RG, Priestley GV, Papayannopoulou T (1997) Antibodies to VLA4 integrin mobilize longterm repopulating cells and augment cytokineinduced mobilization in primates and mice. Blood 90:4779–4788 65. Papayannopoulou T, Craddock C, Nakamoto B, Priestley GV, Wolf NS (1995) The VLA4/ VCAM-1 adhesion pathway defines contrasting mechanisms of lodgement of transplanted murine hemopoietic progenitors between bone marrow and spleen. Proc Natl Acad Sci USA 92:9647–9651 66. Papayannopoulou T, Priestley GV, Nakamoto B (1998) Anti-VLA4/VCAM-1-induced mobilization requires cooperative signaling through the kit/mkit ligand pathway. Blood 91:2231–2239 67. Ramirez P, Rettig MP, Uy GL, Deych E, Holt MS, Ritchey JK et al (2009) BIO5192, a small molecule inhibitor of VLA-4, mobilizes hematopoietic stem and progenitor cells. Blood 114:1340–1343 68. Bonig H, Wundes A, Chang KH, Lucas S, Papayannopoulou T (2008) Increased numbers of circulating hematopoietic stem/progenitor cells are chronically maintained in patients treated with the CD49d blocking antibody natalizumab. Blood 111:3439–3441

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69. King AG, Horowitz D, Dillon SB, Levin R, Farese AM, MacVittie TJ et al (2001) Rapid mobilization of murine hematopoietic stem cells with enhanced engraftment properties and evaluation of hematopoietic progenitor cell mobilization in rhesus monkeys by a single injection of SB-251353, a specific truncated form of the human CXC chemokine GRObeta. Blood 97:1534–1542 70. Laterveer L, Lindley IJ, Hamilton MS, Willemze R, Fibbe WE (1995) Interleukin-8 induces rapid mobilization of hematopoietic stem cells with radioprotective capacity and long-term myelolymphoid repopulating ability. Blood 85:2269–2275 71. Laterveer L, Lindley IJ, Heemskerk DP, Camps JA, Pauwels EK, Willemze R et al (1996) Rapid mobilization of hematopoietic progenitor cells in rhesus monkeys by a single intravenous injection of interleukin-8. Blood 87:781–788 72. Sweeney EA, Papayannopoulou T (2001) Increase in circulating SDF-1 after treatment with sulfated glycans. The role of SDF-1 in mobilization. Ann N Y Acad Sci 938:48–52 73. Sweeney EA, Lortat-Jacob H, Priestley GV, Nakamoto B, Papayannopoulou T (2002) Sulfated polysaccharides increase plasma levels of SDF-1 in monkeys and mice: involvement in mobilization of stem/progenitor cells. Blood 99:44–51 74. Kikuta T, Shimazaki C, Ashihara E, Sudo Y, Hirai H, Sumikuma T et al (2000) Mobilization of hematopoietic primitive and committed progenitor cells into blood in mice by anti-vascular adhesion molecule-1 antibody alone or in combination with granulocyte colony-stimulating factor. Exp Hematol 28:311–317 75. Ulyanova T, Priestley GV, Nakamoto B, Jiang Y, Papayannopoulou T (2007) VCAM-1 ablation in nonhematopoietic cells in MxCre + VCAM1f/f mice is variable and dictates their phenotype. Exp Hematol 35:565–571 76. Calvi LM, Adams GB, Weibrecht KW, Weber JM, Olson DP, Knight MC et al (2003) Osteoblastic cells regulate the haematopoietic stem cell niche. Nature 425:841–846 77. Maestroni GJ, Conti A (1994) Modulation of hematopoiesis via alpha 1-adrenergic receptors on bone marrow cells. Exp Hematol 22: 313–320 78. Katayama Y, Battista M, Kao WM, Hidalgo A, Peired AJ, Thomas SA et al (2006) Signals from the sympathetic nervous system regulate hematopoietic stem cell egress from bone marrow. Cell 124:407–421 79. Jiang S, Alberich-Jorda M, Zagozdzon R, Parmar K, Fu Y, Mauch P, et al (2010) Cannabinoid receptor 2 and its agonists mediate

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hematopoiesis and hematopoietic stem and progenitor cell mobilization. Blood [E-pub 2010 Nov 9] 80. Gomes AL, Carvalho T, Serpa J, Torre C, Dias S (2010) Hypercholesterolemia promotes bone marrow cell mobilization by perturbing the SDF-1:CXCR4 axis. Blood 115:3886–3894 81. Carlo-Stella C, Di NM, Magni M, Longoni P, Milanesi M, Stucchi C et al (2002) Defibrotide in combination with granulocyte colony-stimulating factor significantly enhances the mobilization of primitive and committed peripheral

blood progenitor cells in mice. Cancer Res 62:6152–6157 82. Albanese P, Caruelle D, Frescaline G, Delbe J, Petit-Cocault L, Huet E et al (2009) Glycosaminoglycan mimetics-induced mobilization of hematopoietic progenitors and stem cells into mouse peripheral blood: structure/ function insights. Exp Hematol 37:1072–1083 83. Cline MJ, Golde DW (1977) Mobilization of hematopoietic stem cells (CFU-C) into the peripheral blood of man by endotoxin. Exp Hematol 5:186–190

Chapter 2 Quantifying Hematopoietic Stem and Progenitor Cell Mobilization Shiri Gur-Cohen, Kfir Lapid, and Tsvee Lapidot Abstract Allogeneic donor blood cells and autologous peripheral blood leukocytes (PBL), obtained following clinical mobilization procedures, are routinely used as a major source of hematopoietic stem and progenitor cells (HSPC) for transplantation protocols. It is, therefore, essential to evaluate and to quantify the extent by which the HSPC are mobilized and enriched in the circulation in correlation with their longterm hematopoietic reconstitution capacity. In this chapter, we describe quantitative methods that measure the number of mobilized HSPC according to specific criteria, as well as their functional properties in vitro and in vivo. The described assays are useful for assessment of progenitor cell mobilization as applied to both human and murine HSPC. Key words: Mobilization, G-CSF, Hematopoietic stem and progenitor cells, SDF-1/CXCR-4

1. Introduction The process by which adult stem and progenitor cells are recruited from their supportive microenvironment during stress conditions and enter the blood stream is defined as mobilization. The major organ in which repopulating hematopoietic stem and progenitor cells (HSPC) reside is the bone marrow (BM); however, low levels of HSPC can also be localized in distinct organs, such as the spleen, liver, and muscle (reviewed in Schulz et al. (1)). Although the vast majority of progenitor cells reside in the BM reservoir of immature and maturing leukocytes, a small amount of HSPC continuously egress from the BM to the peripheral blood (PB). These small numbers of circulating HSPC can be dramatically increased during stress conditions, including injury, mild bleeding, physical exercise, inflammation due to bacterial and viral infections, and DNA

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_2, © Springer Science+Business Media, LLC 2012

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damage (reviewed in Lapidot and Kollet (2)). Several clinical protocols mimicking stress conditions due to alarm situations were developed to enable the use of PB mobilized HSPC, rather than direct retrieval of the cells from the bones, in order to reconstitute hematopoiesis in ablated patients. A wide array of mobilizing agents have been discovered in the previous decades, including chemokines, cytokines, and chemotherapeutic agents, among which the most commonly used is the cytokine granulocyte-colony stimulating factor (G-CSF) either alone or following chemotherapy (3). The chemokine SDF-1 (also termed CXCL12) and its major receptor CXCR-4 play key roles in both maintenance and motility of HSPC. Of interest, the SDF-1/CXCR-4 signaling axis is extensively studied in the context of the mobilization process, HSPC homing to the BM and steady-state egress (4). The interactions between SDF-1 and CXCR-4 are crucial for balancing retained HSPC in a quiescent noncycling mode in the BM microenvironment (5, 6). More than two decades ago, the BM stromal microenvironment which robustly expresses SDF-1, was revealed as a negative regulator of HSPC proliferation and differentiation, believed to be a part of the mechanism to preserve quiescent HSPC pool in the BM (7). Maintenance of HSPC retention in the BM involves adhesion interactions and signals, which are mediated by BM stromal supporting cells (reviewed in Wilson et al. (8)). Tight regulation of SDF-1 in the BM promotes the expression of adhesion molecules such as VLA-4/5, LFA-1 (9), and CD44 (10), which mediate HSPC attachment to BM matrix components and stromal supporting cells. Breaking the fine-tuned SDF-1/CXCR-4 balance under stress conditions allows HSPC proliferation, differentiation and subsequent mobilization (11). The major roles of SDF-1 in HSPC mobilization can be inferred from the observation that, following G-CSF treatment, the levels of SDF-1 mRNA (12) and protein (11–13) are dramatically reduced in the BM, while the levels of CXCR-4 are increased, enabling motility of HSPC toward the blood (11). Indeed, increased plasma levels of SDF-1 attract HSPC and thereby increase cell mobilization to the circulation (14). Active disruption of SDF-1/CXCR-4 dynamic interactions by the CXCR-4 antagonist AMD3100 serves as another clinically used protocol to induce HSPC mobilization (15). While G-CSF induced mobilization requires repetitive daily stimulations, AMD3100 is a rapid mobilizing agent that triggers HSPC recruitment to the circulation within a few hours after administration (15, 16). Of interest, combined treatment of G-CSF and AMD3100 synergistically augment HSPC recruitment to the circulation (15, 17). Stress-induced mobilization is a multievent process that involves activation of bone-resorbing osteoclasts (18, 19), regulation of osteoblasts by β-adrenergic signals (20), neutrophil activation (11), proteolytic enzyme activity (21, 22), reactive oxygen species (ROS) signaling (23) and various cytokines and chemokines. All of these factors facilitate the detachment of HSPC from

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their anchored microenvironment, their proliferation and differentiation, migration through the mechanical barrier of the bone-blood endothelial barrier and finally HSPC recruitment to the blood circulation (11, 24–26). Recently, interactions between SDF-1 and CXCR-4 were demonstrated by a rare mesenchymal stem cell (MSC) subpopulation identified by Nestin expression, which represent approximately 4% of the murine CD45− BM nonhematopoietic cells and 0.08% of total BM cells. Nestin+ MSC, functionally defined by exclusive colony-forming units-fibroblast (CFU-F) content in the murine BM, were found to express very high levels of SDF-1 and are physically associated with SLAM CD150+ CD41− CD48− hematopoietic stem cells (HSC). Interestingly, transplanted HSC rapidly home and lodge in proximity to mesenchymal Nestin+ stem cells, whereas in vivo depletion of Nestin+ cell significantly reduces the homing capacity of transplanted HSPC. Selective inhibition of SDF-1 transcription by Nestin+ MSC cells and suppression of their proliferation during G-CSF stimuli exemplifies the disruption of the SDF-1/CXCR-4 axis during HSPC mobilization (27). Not only HSPC undergo mobilization, but also other immature cell types, including endothelial progenitor cells (EPC) (28–30). Stromal progenitor cells (or CFU-F) were also found to undergo expansion as a consequence of increased bone turnover due to accelerated osteoclast activity during G-CSF induced mobilization (31). Notably, both BM derived HSPC and EPC have significant clinical benefits, share similarities and overlapping mobilization mechanisms, involving altered balance of the SDF-1/CXCR-4 interactions and RANKL induced osteoclast activation (28–30). In order to evaluate mobilization of immature leukocytes, several in vivo and in vitro assays have been developed for characterization and quantification of both progenitors and more primitive hematopoietic stem cell levels. Herein, we describe in vitro techniques and the use of functional in vivo models to assess murine, as well as a preclinical model for human HSPC mobilization in immune deficient NOD/SCID chimeric mice.

2. Materials 2.1. HSPC Marker Characteristics: Flow Cytometry (FACS) Assay

1. FACS buffer: DPBS−/− 10× (Dulbecco’s Phosphate Buffered Saline 10×, without calcium and magnesium) supplemented with heat-inactivated 5% fetal calf serum (FCS) and 0.1% sodium azide. 2. Antibodies: For murine SKL staining-FITC conjugated antilineage markers (CD4, NK, GR-1, B220, CD8a, and CD11b— additional markers can be included, such as Ter119 for erythrocytes), PE conjugated anti-Sca-1 and APC conjugated anti c-Kit (BioLegend). For human CD34+/CD38− staining,

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FITC conjugated anti-CD34 and PE conjugated anti-CD38 (BD Biosciences) (see Note 1). 3. FACS tubes (BD Biosciences). 4. Flow cytometry analyser (e.g., FACSCalibur with CellQuest software). 2.2. Cell Cycle Analysis

1. FACS buffer (see Subheading 2.1). 2. Antibodies: For murine SKL staining-FITC conjugated antilinage markers (CD4, NK, GR-1, B220, CD8a, and CD11b), PE conjugated anti-Sca-1 and APC conjugated anti-c-Kit (BioLegend). For human CD34+/CD38− staining, FITC conjugated anti-CD34 and PE conjugated anti-CD38 (BD Biosciences) (see Note 1). 3. FITC conjugated anti-KI-67 and 7-AAD (BD Biosciences). 4. Hematopoietic progenitor enrichment kit and BD IMagnet (BD Biosciences). The kit contains IMag buffer supplied as 10× stock, blocking antibody, cocktail of biotinylated anti mouse lineage depleting antibodies and streptavidin particles. 5. Fixation/permeabilization kit (BD Biosciences). The kit contains two reagents, fixation/permeabilization solution and the Perm/Wash Buffer supplied as 10× stock solution, which is diluted in FACS buffer to obtain 1× buffer. 6. FACS tubes (BD Biosciences). 7. Flow cytometry analyser (e.g., FACSCalibur with CellQuest software).

2.3. In Vitro Colony Assays (CFU-C) in Semi Solid Media (Methylcellulose)

1. Methylcellulose preparation: Boil 500 ml deionized water (ddH2O) and add 20 g of methylcellulose powder (SigmaAldrich), while stirring. Cool to RT and add 500 ml DMEM (concentrated 2×) with 1% penicillin and streptomycin antibiotics. Aliquots can be stored at −20°C. The mixture is stable at 4°C for up to 1 month. 2. Supplements: 30% FCS, 50 ng/ml SCF, 5 ng/ml IL-3, 5 ng/ ml GM-CSF (R&D Systems) and 2 μl/ml erythropoietin (Orto BioTech). 3. Tissue culture dishes of 35 × 10 mm and 100 × 20 mm, Nunclon Surface. 4. Syringe, 16 gauge blunt end needle is recommended.

2.4. In Vitro Transwell Migration Assays

1. Costar transwells (6.5 mm diameter, 5 μm pore). 2. Migration assay medium: RPMI supplemented with heat-inactivated 10% FCS, 2 mM L-glutamine, and 1% penicillin and streptomycin antibiotics. 3. SDF-1α 125 ng/ml (PeproTech), keep at −20°C and store at 4°C before use (stable at 4°C for up to 2 weeks).

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4. FACS tubes (BD Biosciences). 5. Flow cytometry analyser (e.g., FACSCalibur with CellQuest software). 2.5. Proteolytic Enzyme Activity

1. Plastic gel casting cassettes, 0.75 mm thick, 10 × 10 cm. 2. 30% Acrylamide. 3. 1% (w/v) gelatin: Dissolve 40 mg gelatin in 1 ml of ddH2O and heat the solution at 60°C in water bath for at least 20 min; mix well. Make sure that the gelatin is completely dissolved. Cool the gelatin solution to room temperature before use. Prepare it fresh. 4. 10% SDS. 5. 10% (w/v) ammonium persulfate. 6. N,N,N¢,N¢-Tetramethylethylenediamine (TEMED). 7. Running buffer stock (10×): Prepare 1 l of 0.25 M Tris base and 1.92 M glycine, pH 8.3. Adjustment of the pH is not required. Store at room temperature. Dilute the running buffer 10× stock with dH2O to make 1 l and supplement with 5 ml of 20% (w/v) SDS to a final concentration of 0.1% (w/v). Store solution at room temperature. All buffers are stable for months. 8. Sample buffer (4×): Prepare 10 ml of 250 mM Tris–HCl, pH 6.8, 40% (v/v) glycerol, 8% (w/v) SDS, and 0.01% (w/v) bromophenol blue. Store at −20°C. Before use, warm solution to dissolve the SDS. 9. Collagenase buffer stock (10×): Mix 60.6 g Tris base, 117 g NaCl, 5.5 g CaCl2, complete to 900 ml with ddH2O. Adjust to pH 7.6 with concentrated HCl, top up to 1 l with ddH2O and store at 4°C. For an 1× working solution, dilute the running buffer stock 10× with ddH2O to a volume of 1 l and add 670 μl 30% (w/v) Brij-35 (Sigma-Aldrich). 10. Developing buffer: Prepare 50 mM Tris–HCl, pH 8, 10 mM CaCl2, 1 mM ZnCl2 and 1% Triton X-100. 11. Coomassie Brilliant Blue staining R250. 12. Destaining solution: 5% Acetate, 10% methanol in dH2O. 13. Image analysis program (e.g., ImagJ or NIH Image processor) and a scanner.

2.6. Functional HSC Engraftment, Repopulation and Serial BM Transplantation

1. Mobilized mouse cell transplantation: (a) Recipient mice: B6.SJL (CD45.1) mice, 8–12 weeks old. (b) Cell donors: C57BL/6 (CD45.2) mice, 8–12 weeks old. 2. For mobilized human cell transplantation: (a) Mononuclear cells or CD34+ enriched cells collected from human mobilized blood. (b) Recipient immunodeficient mice: NOD/SCID or NOG mice, 8–12 weeks old.

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3. RPMI medium supplemented with 10% heat-inactivated FCS, 2 mM L-glutamine, and 1% penicillin and streptomycin antibiotics. 4. 16-Gauge, flat bore needles with syringe. 5. Phosphate-buffered saline (PBS 1×). Dilute DPBS−/− 10 × (-) CaCl2 (-)MgCl2. 6. Ficoll-Hypaque (light sensitive). 7. Anti-mouse antibodies: FITC conjugated anti-CD45.2, PE conjugated anti-CD45.1, and IgG isotype control (BioLegend). 8. Anti human antibody: FITC conjugated anti-CD34 (BD Biosciences). 9. FACS buffer: PBS−/− (DPBS−/− 10− (-)CaCl2 (-)MgCl2) supplemented with 5% heat-inactivated FCS and 0.1% sodium azide. 10. FACS tubes (BD Biosciences). 11. Ciprofloxacin antibiotic (Bayer). 12. Flow cytometry analyser (e.g., FACSCalibur with CellQuest software). 13. Irradiator. 14. Heat lamp (recommended).

3. Methods 3.1. Phenotypic Characterization of Mobilized Hematopoietic Stem and Progenitor Cells 3.1.1. HSPC Marker Characteristics: Flow Cytometry (FACS) Assay

HSPC respond to stress-induced signals, such as bleeding and inflammation, by detachment from their BM microenvironment and recruitment to the circulation. Clinical mobilization protocols mimic these stress signals by administration of mobilizing agents, such as repetitive G-CSF stimulations (3). Mobilized HSPC from the PB can be easily identified, characterized and quantified by a molecular signature of extracellular antigen sets. The heterogeneity of cell surface markers enables one to distinguish primitive stem cells from other uncommitted and lineage-restricted progenitors. Over the years, numerous flow cytometry based methods have been established to identify and distinguish human and murine stem cells (32). Enriched murine HSPC are identified as Sca1+Lineage−c-Kit+ (33, 34) cells (SKL) or CD34−Sca-1+Lineage−cKit+ (SKL/CD34−), while lineage + marker combinations identify differentiating leukocytes. Most SKL cells are considered as progenitor cells, while the SKL/CD34− cells are more primitive, retaining long-term repopulating potential (35). Recently, the expression of SLAM family antigens, CD150+ CD41− CD48−, has been reported as another tool to identify and characterize primitive

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HSC (36). Human HSPC are enriched in the CD34+ fraction, which can be further purified into CD34+/CD38− or CD34+/ CD38−/Lin−. Herein, we describe the use of a well-known set of markers for the identification and quantification of both murine and human HSPC. Beside those conventional sets of markers, others molecules that highly associate with mobilized HSPC function can be determined since their levels change and correlate with mobilization. These include CD44 (10), which is cleaved as part of the mobilization process, as well as the membrane-bound MT1MMP which its increased levels, correlate with clinical mobilization protocols (37). Both CD44 and MT1-MMP are also important for human CD34+ HSPC homing capacity and motile behavior (37, 38). Assessment of HSPC mobilization can be usually done by identifying murine SKL cells or human CD34+/CD38− in the PB. 1. For the detection of mobilized HSPC, murine or human PB mononuclear cells (MNC) (at least 1 × 106 cells, for the separation of PB MNC by ficoll see Subheading 3.4.1) should be centrifuged at 200 × g for 5 min. 2. Wash the cell pellet in 1 ml of FACS buffer (see Subheading 2), and centrifuge at 200 × g for 5 min; 4°C is recommended. 3. Discard supernatant and resuspend the cells in 100 μl of antibody mix. For detection of murine SKL cells, mix 0.5 μg from each of the lineage antigen marker FITC conjugated antibodies (CD4, NK, GR-1, B220, CD8a, and CD11b) and 0.5 μg from the PE conjugated anti-Sca-1 and APC conjugated antic-Kit antibodies for each sample. For detection of human HSPC, add 1 μg of anti-CD34 and anti-CD38 antibodies for each sample of 1 × 106 cells. 4. Mix the cells by gently vortexing and incubate for 30 min at 4°C. After incubation, wash in 1 ml of FACS buffer and centrifuge at 200 × g for 5 min. 5. Resuspend the cells in 300 μl FACS buffer and quantify mobilized HSPC percentage in comparison to controls by FACS. 3.1.2. Cell Cycle Analysis

HSC are maintained primarily in a homeostatic quiescent state, mostly due to a complex tight regulation by the BM microenvironment (5, 19, 39, 40). Following treatment with mobilizing agents, HSPC appear in the circulation in either the G0 or G1 phase of the cell cycle, entering the blood only after the M phase (41), while HSPC that reside in the BM and spleen are more actively cycling (41, 42). What dictates the noncycling state of mobilized HSPC in the PB of both mouse and human is not fully understood; however, it can be speculated that noncycling HSPC with condensed chromatin will migrate from the BM to the circulation more efficiently than cycling cells. Recent evidence suggests that cleavage of the potent cyclin-dependent kinase inhibitor p21 by

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proteinase 3 during stress-induced mobilization is responsible for cell cycle progression into the S phase in the BM and spleen as compared to noncycling circulating cells (43). The cell cycle status of mobilized HSPC can be assessed using DNA intercalating molecules, such as BrdU labeling, Ki-67 nuclear antigen staining, Pyronin Y, or 7-amino actinomycin D (7-AAD). This section describes analytical intracellular staining method for cell cycle marker Ki-67 and 7-AAD with membrane staining for multiple HSPC markers. Ki-67 is a nuclear antigen that is strictly associated with cell proliferation during all active phases of the cell cycle (G1, S, G2, M) but is absent from quiescent, noncycling cells (G0). 7-ADD is a nuclear acid stain with a strong affinity for GC-rich regions. The combination of Ki-67 and 7-ADD staining provides the ability to distinguish between the cell cycle stages in primitive subpopulations. 1. For the assessment of cell cycle stages of mobilized HSPC, centrifuge ficoll-separated MNC from murine or human PB (at least 5 × 106 cells, see Note 2) at 200 × g for 5 min (for the separation of PB MNC see Subheading 3.4.1). 2. Wash the pellet in 1 ml of FACS buffer (see Subheading 2), and centrifuge at 200 × g for 5 min; 4°C is recommended. 3. For murine cells, discard supernatant and continue with depletion of lineage positive cells using the hematopoietic progenitor enrichment kit accordingly to the instructions supplemented with the kit. 4. After the depletion procedure of murine cells or following the washing of human cells, discard supernatant and resuspend the pellet in 100 μl of antibody mix to identify murine or human HSPC, as depicted in Subheading 4.11. 5. This method combines both extracellular and intracellular staining; therefore, it is important to first stain the cell surface antigens and then fix and permeabilize the cells for intracellular staining of nuclear DNA content. After performing extracellular staining, wash the cells in 1 ml of FACS buffer and centrifuge at 200 × g for 5 min. Discard the supernatant, resuspend the cell pellet with 100 μl of fixation/permeabilization solution and incubate for 30 min at 4°C (this can be done also at room temperature). After incubation, wash the cells with 1 ml Perm/Wash buffer 1× and centrifuge at 200 × g for 5 min. It is important to use the Perm/Wash buffer in all subsequent washing steps in order to keep cells permeabilized for intracellular staining. 6. Add 20 μl of Ki-67 to each tube, mix the cells by gently vortexing and incubate for 30 min at 4°C. 7. Wash the cells with 1 ml Perm/Wash buffer 1× and centrifuge at 200 × g for 5 min.

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8. Discard the supernatant and add 20 μl of 7-AAD to each sample and incubate for 10 min at room temperature. 9. Resuspend the cells in 300 μl FACS buffer and quantify mobilized HSPC in comparison to control cells. After reading the samples by FACS, gate on the murine Sca-1 and c-Kit positive cells or the human CD34+/CD38− subpopulation to display the Ki-67 vs. 7-AAD dot plot. The lower left quadrant (cells negative for Ki67 and 7-AAD) represents cells in G0 and the upper right quadrant represents cells in the S, G2 and M phases (cells positive for Ki67 and 7-AAD). 3.2. In Vitro Assays 3.2.1. In Vitro Colony Assays (CFU-C) in Semi Solid Media (Methylcellulose), for Mobilized Human and Murine, as well as Enriched Human CD34+ Cells

Colony-forming cell assays are widely used as a functional method to identify myeloid lineage restricted progenitor cells. While these assays are easy to perform, they cannot identify true HSC and cannot be used to measure the rate of mobilization in real time. Colony-forming cells (CFC) or colony-forming unit cells (CFUC) are committed stem and early multipotent progenitor (MPP) cells that are able to proliferate and differentiate into more mature cells under appropriate conditions. The in vitro hematopoietic colony assay was developed at the beginning of the 1960s, making it possible to investigate and distinguish hematopoietic cell populations at different stages of differentiation. The multi-potent progenitors CFU-GEMM give rise to all the myeloid lineages; granulocyte–macrophage colony-forming cells (CFU-GM) are progenitor cells that can give rise to colonies containing a heterogeneous population of macrophages and granulocytes; lineagecommitted progenitors include burst-forming unit erythroid (BFU-E), colony-forming unit erythroid (CFU-E), which are primitive erythroid progenitors, and megakaryocyte precursors (CFU-Mk). In murine studies, assessment of mobilized progenitors can be performed with cells obtained from different sources, whole leukocytes or purified mononuclear cells (MNC) from the BM, PB or spleen. 1. Colony forming assays are performed by seeding hematopoietic cells (according to Table 1, while the use of PB and spleen MNC is enough for mobilization assessment) into a cell culture dish containing prewarmed semisolid matrix methylcellulose supplemented with 30% FCS, 50 ng/ml SCF, 5 ng/ml IL-3, 5 ng/ml GM-CSF, and 2 μl/ml erythropoietin. 2. The mixture should be vigorously vortexed before plating. Wait approximately 10 min until all air bubbles disappear from the vortexed cell suspension. 3. Collect the mixture with a syringe (a 16 gauge blunt end needle is recommended, since the solution is viscous) and plate 1 ml of the mixture in a 35 × 10 mm culture dish, spreading it out to cover the entire surface of the dish.

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Table 1 Number of cells to be seeded for colony formation assay Source

Cells per dish

Human PB-MNC Mobilized PB-MNC Mobilized PB CD34+ enriched cells

1–2 × 105 1 − 5 × 104 5 × 102 to 2 × 103

Murine Total crude BM cells Total crude spleen cells PB-MNC Mobilized PB-MNC

1.5 × 103 2 × 105 2 × 104 2 × 104

4. Duplicates of colony culture dishes should be incubated together with additional 35 × 10 mm dish containing sterile water but without a lid to maintain high humidity. The dishes are placed together in a 100 × 25 mm dish and incubated at 37°C in a humidified atmosphere (>96%), containing 5% CO2 for 7 days (murine colonies) or 14 days (human colonies). 5. In order to assess CFU-C frequency, place the dish containing cell colonies on a 60 mm gridded scoring dish, and count the colonies of interest using an inverted microscope. Moreover, the absolute number of CFU-C can be counted per 1 ml of blood by taking into account the frequency of CFU-C, the blood volume and the number of PB-MNCs after ficoll enrichment (e.g., (PBMNC number × 1 ml × CFU − C number) ). (blood volume (ml) × seeded cells) 3.2.2. In Vitro Transwell Migration Assays (Directional and Spontaneous Migration)

HSPC mobilization is modulated and controlled by multiple factors, including chemokines, cytokines, growth factors and hormones. Particularly, the SDF-1/CXCR-4 axis plays major roles in HSPC maintenance, homing and mobilization. During G-CSF induced mobilization, SDF-1/CXCR-4 signaling is altered, resulting in reduced SDF-1 levels in the BM and upregulation of CXCR-4 expression, leading to HSPC mobilization. In addition to the pivotal role played by SDF-1 and CXCR-4 in HSPC motility, the mobilization process was found to be tightly regulated by the cytokine hepatocyte growth factor (HGF), and its receptor c-Met, involving elevation in intracellular ROS signaling. Of interest, c-Met inhibition reduced HSPC mobilization following G-CSF treatment and interfered with their chemotactic migration towards SDF-1 (23). C-Met inhibition also reduced MT1-MMP elevation on mobilized human CD34+ HSPC (38). As SDF-1/CXCR-4 signaling is essential for directional HSPC migration and can predict clinical repopulation outcome in autologous transplantation (44).

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In vitro migratory capacity of CD34+ cells is related to hematopoietic recovery after autologous stem cell transplantation (44). An assay for SDF-1 induced transmigration evaluates the motility properties of mobilized HSPC. Recruitment of HSPC from the BM microenvironment to the blood stream and vice versa, is a multistep process that involves adhesion, directed migration and proteolytic enzymes, which degrade the extracellular matrix barrier. Coating the transwell filter with the barrier of interest, such as fibronectin, endothelial cells, or matrigel (which contains extracellular matrix proteins that partially mimic the extracellular basement membrane), can be applied to evaluate the ability of cells to penetrate through an artificial barrier under semi-physiological conditions. 1. For the transmigration assay, add 600 μl migration medium to the lower chamber supplemented with or without 125 ng/ml SDF-1α. The herein described method using human SDF-1 can be applied both to human and murine cells, because the SDF-1 sequence is highly conserved and there is a crossreactivity between human and mouse SDF-1. 2. Load 100 μl of 50,000–100,000 human enriched CD34+/ murine SKL cells or 100,000–200,000 BM/PB MNC gently into the upper chamber of each transwell (take special care to avoid air bubbles in the chamber and perforation of the membrane). 3. In parallel, add 100 μl of cells into a separate tube containing 500 μl of migration medium. This tube serves as the migration index (see Note 3). 4. After loading the cells, transfer the upper chamber into the lower chamber (avoid air bubbles that may interfere with the transmigration of cells). 5. Place the transmigration plate in a humidified 37°C containing 5% CO2 incubator for 2–4 h (see Note 4), and make sure it is not disturbed. 6. Stopping the transmigration experiment is done by transfer the upper filter chamber into an empty well very gently, avoiding media transfer between the upper and lower compartments of the transmigration wells. 7. Collect 300 μl of media from the lower chamber into a FACS tube and evaluate the migration capacity by counting the cells at high speed for 60 s using FACS. To calculate the percentage of cells migrating (the migration capacity), take into account the migration index (see Note 3). Migrating fractions can be also stained for murine Lin-c-Kit+, human CD34 (see Subheading 3.1.1), as well as examined for CFU-C potency (see Subheading 3.2.1). These last two assays are useful mainly for the detection of hematopoietic progenitor cells (e.g., Lin−cKit+, CD34+, and CFU-C) with high migration capacity.

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3.2.3. Proteolytic Activity (Zymography Assay)

A major group of proteases has been directly associated with HSPC mobilization from the BM microenvironment to the circulation following stress signals. Neutrophil accumulation and activation in the BM and other BM stromal cells play a pivotal role in HSPC motility, following G-CSF-induced mobilization, through the release of active neutrophil proteases, such as elastase, cathepsin G, and matrix metalloproteinase-9 (MMP-9) (21). Circulating human CD34+ cells (45), murine BM osteoclasts (46) and hematopoietic progenitor cells (45), secrete metalloproteinases MMP-2 or MMP-9. These active proteases are released into the BM cavity and selectively cleave chemokines, cytokines, and their receptors, as well as adhesion molecules, that are essential for HSPC retention in the BM microenvironment, such as VCAM-1(21), the receptor c-Kit (13, 21), and SDF-1 (21). Indeed, there is an accelerated induction of proteolytic activity in the BM following G-CSF administration, accompanied by a decrease in SDF-1 levels and subsequently HSPC mobilization to the circulation (11, 21). CD26/DPPIV is a murine progenitor and human CD34+ associated peptidase that has an important role in HSPC mobilization by facilitating SDF-1 cleavage in the BM (47). Notably, it has been shown that mice deficient either in MMP-9, neutrophil elastase, cathepsin-G, or CD26 are able to respond to G-CSF-induce mobilization normally, suggesting redundancy in the activity of different proteases (48). In addition, the membrane type 1–MMP (MT1-MMP) has been reported to be highly expressed on mobilized HSPC, regulating their motility and mobilization (21, 37, 49). Hereby, we provide a detailed protocol to measure MMP-9 activity in the mouse BM and plasma by gelatin zymography. The gelatin zymography assay is an easy yet powerful technique to detect the presence of MMP-9 (and MMP-2) in biological samples by identifying gelatindegrading activity. Protein extraction can be done from mouse BM supernatant. 1. For mouse BM MMP-9 expression, extract the BM from the femurs and tibias by cutting off the tips of the bones and flushing the marrow out of the bone cavity, using a 1 ml syringe containing PBS. 2. Centrifuge the samples at 180 × g for 10 min at 4°C and transfer the BM supernatant to a new Eppendorf tube. 3. For mouse plasma MMP-2 and MMP-9 expression, collect PB and centrifuge at 180 × g for 10 min at 4°C, and transfer plasma (the supernatant) into a new Eppendorf tube. 4. Samples can be maintained for long term storage at −20°C. 5. Prepare 10% SDS-polyacrylamide gels supplemented with 1 mg/ml gelatin. For the resolving gels (two gel preparations), mix 3 ml of dH2O, 3.3 ml of 40% acrylamide, 100 μl of 10% (w/v) SDS, 100 μl of ammonium per-sulfate and 2.5 ml of

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1.5 M Tris buffer pH 8.8 and 1 ml of gelatin (as mentioned in Subheading 2). Add 5 μl TEMED to the resolving gel solution to initiate polymerization and rapidly swirl the solutions, while avoiding air bubbles. Immediately, transfer 5 ml of the resolving gel solution into the running cassette, then overlay the separating gel solution with dH2O, and let the gel polymerize for at least 30 min at room temperature. After discarding the overlying water, add 5 μl TEMED to the stocking gel solution containing 3.15 ml of dH2O, 0.66 ml of 40% acrylamide, 50 μl of 10% (w/v) SDS, 100 μl of ammonium per-sulfate and 1.25 ml of 1.5 M Tris buffer pH 6.8. Gently swirl and rapidly transfer to the upper running cassette. Insert immediately the appropriate comb and let the stacking gel polymerize for 1 h at room temperature. 6. Mix volumes of BM supernatant equivalent to 2 μg protein or 5 μl of plasma with 7.5 μl nonreducing sample buffer 4× (without mercaptoethanol) and dilute to a final volume of 30 μl of collagenase buffer without heat treatment, loading 15–30 μl per lane (see Note 5). Use conditioned medium from HT1080 (human fibrosarcoma) cell line culture as a positive control for MMP-9 and MMP-2 expression. 7. Let the gels undergo electrophoresis at 300 V for 1 h and 40 min in running buffer. 8. Gently remove the gels from the running cassette, transfer the gels to a new container and wash them in a 2.5% Triton X-100 solution for 30 min (see Note 6). 9. Wash the gels 2–3 times in dH2O for 10–15 min. 10. Incubate the gels in developing buffer 1× with gentle shaking for 30 min at room temperature. 11. Discard used developing buffer 1× and incubate the gels in fresh developing buffer 1× over night in 37°C (see Note 7). Protease activity can be visualized as a clear zone with Coomassie Brilliant Blue staining within 3 min of incubation. 12. Add destaining solution with gentle shaking for 30 min. Discard used destaining solution and incubate the gels in ddH2O until clear bands appear on the gels. 13. Scan gels using a flat bed scanner and analyze gelatinase activity using an image processor. 3.3. Functional In Vivo Assays: HSC Engraftment, Repopulation and Serial Transplantation

Mobilized HSPC can be identified based on their in vivo functional capacity to repopulate host BM with high levels of maturing myeloid and lymphoid cells, while the “stemness” property is maintained by a small pool of undifferentiated stem cells with the potential to repeat the entire process in serially transplanted recipients. The gold standard for measuring HSC function following the mobilization process is the long-term repopulation assay. This assay

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determines the potential of HSC to fully differentiate into all blood lineage-restricted cells and their longevity by preserving the selfrenewal potential, which enables the preservation of the HSC pool for many years. The repopulation assay utilizes transplantation of mobilized blood MNC or sorted immature cells in preconditioned irradiated hosts. Short-term engraftment is initiated by differentiating progenitors (SKL Sca-1+c-Kit+Lin− cells in mouse or CD34+CD38+ cells in human), usually evaluated after four weeks up to a couple of months. Long-term multilineage engraftment is carried out by “true” stem cells (E-SLAM EPCR+CD48−CD150+ (50) cells or SKL CD34− cells in mouse and CD34+CD38− cells in human), evaluated after 4–6 months in mice and years in patients (51). HSPC transplantation requires preconditioning of the host with total body irradiation (TBI), thus the homing ability and engraftment potential of mobilized HSPC is not examined under physiological conditions. A more physiological relevant assay, using parabiotic mice with a shared blood circulation, has revealed that HSPC are constitutively circulating, trafficking from the BM to blood stream through the blood–bone barrier and return to the BM of the parabiont partner, and enabling functional engraftment of unconditioned BM (52–54). While the blood and spleens of these mice are equally repopulated by both parabionts, the BM is mostly host derived due to the blood–bone barrier. Of interest, HSPC mobilization by G-CSF (53) or AMD3100 (54) in parabiotic mice, which has been done in a more physiological context, is accompanied by a dramatic increased engraftment of the partner BM with primitive cells, revealing that mobilization and homing are sequential events with clinical relevance. Assessment of murine HSC engraftment and repopulation following stress induced mobilization requires congenic strains of donors and hosts, for example by identification of different alleles of the hematopoietic cell marker CD45. Assessment of human cell mobilization requires sublethally irradiated (300–375 cGy) immune-deficient recipient mice that are able to tolerate human xenografts, such as the NOD/SCID or the NOG strains (owing to the complete absence of T, B, and NK cell activity, and reduced function of innate immunity), allowing multilineage reconstitution of human hematopoiesis (55). Notably, some reports indicate that mobilized CD34+ cells, as compared to cord blood, can provide early multilineage reconstitution though nonsustained long-term multilineage engraftment of immunodeficient mice (56). The most stringent test for stemness is the serial transplantation assay, wherein only the most primitive HSC can yield longterm multilineage repopulation. In this assay, mobilized human MNC or enriched CD34+ cells are transplanted into primary recipients, then harvested after 4 weeks and transplanted again into secondary recipients. Obviously, subsequent serial transplantations can be done.

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1. Irradiate host mice 1 day prior to transplantation. Recipient mice for human grafts (NOD/SCID) should be irradiated with a sublethal dose of ~350 cGy, and recipient mice for mouse grafts (B6.SJL CD45.1) should be irradiated with a lethal dose of ~900 cGy, depending on the irradiation device. 2. Add 16 μg/ml Ciprofloxacin antibiotics to the drinking water after irradiation. Replace the antibiotic-containing water once a week for 3 weeks.

3.3.2. Donor Mobilized MNC Preparation

1. Once mobilization with a mobilizing agent has been assessed in human or in the mouse, dilutes mobilized cells at least 1:1 with sterile PBS and separate mononuclear cells by a ficoll gradient. 2. Prepare falcon tubes with 1 ml ficoll reagent. Load 2 ml of diluted mobilized blood cells slowly on the top of ficoll, using a 1 ml pipette at a 45 angle against the wall of the tube. 3. Carefully transfer the tubes to the centrifuge without mixing the layers. Centrifuge at 220 × g for 25 min. A fraction of enriched mononuclear cells is found at the middle of the tube arranged in an annular-like shape. 4. Discard the plasma from the top of the tube without disturbing the MNC fraction. 5. Transfer the MNC fraction to a new PBS containing tube. 6. Wash out ficoll reagent by centrifuging the tube at 200 × g for 5 min, discard the supernatant and resuspend the cell pellet in 1 ml of full-RPMI. 7. Prepare 20 million MNC (both for human or mouse cells) in 500 μl of RPMI per mouse and inject the cells immediately without incubation.

3.3.3. Transplantation

1. Mice are irradiated approximately 24 h prior to transplantation. 2. When ready, place the recipient mouse in a restrainer and inject 20 million cells per mouse intravenously. Placing recipient mice under a heating lamp for about 2 min might help to find the lateral tail veins more easily.

3.3.4. Evaluation of Engraftment

1. Engraftment is measured 1 month after transplantation for short-term engraftment and 6 month for long term engraftment. Assessment of the engraftment levels can be monitored from the PB by mild bleeding, while keeping the animal alive for further examination, or by sacrificing the animal and extracting total BM cells. Although both PB and the BM undergo engraftment, the best way to analyze reconstitution is by monitoring the level of engraftment in the BM of recipients. In addition, analysis of the BM rather than PB is recommended, since it has previously been demonstrated that

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bleeding results in accelerated hematopoiesis and mobilization of HSPC from the BM to the circulation (18). 2. Remove the femur and the tibia of the mice and extract total BM cells into 1 ml of PBS by cutting off the tips of the bones and flushing the marrow out of the bone cavity using a 1 ml syringe. 3. Transfer at least one million cells to a new FACS tube and resuspend the cells in 1 ml of FACS buffer. 4. Centrifuge the cells at 200 × g for 5 min, discard supernatant and resuspend the cell pellet in 100 μl of antibody mix containing CD34 for human or CD45.1 and CD45.2 for mouse. 5. Mix the cells by gently vortexing and incubate for 30 min at 4°C. After incubation, wash the cells in 1 ml of FACS buffer and centrifuge at 200 × g for 5 min. 6. Discard the supernatant, resuspend the cells in 300 μl FACS buffer and quantify the percentage of engrafted mobilized HSPC in the host. 3.4. Concluding Perspectives and Other Approaches to Analyzing HSPC Mobilization

Although an extensive body of literature describes the nature of HSPC mobilization, which is accelerated following additional stress conditions, still much has to be learned in order to better understand the molecular mechanisms by which HSPC are dynamically guided to navigate from their anchored microenvironment in the BM to the circulation. There is a constant demand for improvement of mobilization strategies, so as to overcome “poor mobilization” difficulties, by increasing the number of primitive stem cells, as well as their functional homing and engraftment efficacies for clinical transplantation. Numerous mobilization assays have been developed during the years, some of which have been detailed above, in order to evaluate the efficacy of novel mobilizing agents and to analyze the long-term repopulation potential of the mobilized stem cells. Despite the multiple established assays for mobilization assessment, critical distinction between the source organs for murine HSPC (e.g., BM vs. the spleen) remains obscure. However, recently, a novel technique of in situ perfusion, which is installed on the murine hind limb, enables a direct in vivo assessment of mobilization from the BM (57). This elegant approach contributes to the progress in the field by allowing detection of the ability of new mobilizing agents to induce HSPC recruitment directly from the BM cavity to the circulation, and thereby might assist in developing improved transplantation protocols. Extensive research in the field has uncovered key roles for microenvironmental cues in inducing mobilization and augmenting HSPC motility. For example, bone-resorbing osteoclasts, which are essential for homeostatic bone turnover, are activated in response to G-CSF stimulations, while bone-lining osteoblasts are suppressed. This dynamically coordinated and regulated microenvironment “clears the way out” for mobilized HSPC together with other myeloid

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cells, such as neutrophils. Osteoclast differentiation from monocyte precursors under RANK-ligand regulation was also associated with moderate HSPC mobilization (18). Of note, osteoclast activity releases active TGF-β1 from bone matrix, forming a gradient that recruits Sca-1+ MSC to the endosteum region, where they undergo osteoblast differentiation (58). G-CSF administration has been demonstrated to deplete a population of trophic endosteal macrophages (osteomacs) that support osteoblast function (59). Inhibition of osteoclast by bisphosphonates on the other hand, impaired HSC levels and quiescence and was demonstrated to induce delayed hematopoietic recovery following transplantation (60). Multiple studies have shed light on the putative role of BM stromal supporting cells in preserving the immature-primitive phenotype of HSC. Notably, osteoblast lineage cells robustly express the key molecule SDF-1, thus providing a unique supportive microenvironment for HSC maintenance, contributing to longterm hematopoiesis (61). SDF-1 is presented by Annexin-2, which is expressed by BM osteoblasts. Thus, CXCR4+ HSC are also anchored directly to the internal bone surface. Indeed, higher rates of G-CSF-induced mobilization of HSPC were observed in Annexin-2 deficient mice as compared to wild-type animals (62). HSPC mobilization also requires signals from the sympathetic nervous system, which plays a major role by affecting HSPC directly (e.g., catecholamines) and indirectly (e.g., suppression of osteoblasts or daily rhythmic regulation of SDF-1 synthesis) (20, 63–65). Neurotransmitters can be transmitted to HSPC through the blood stream or directly secreted from nerve endings in the BM. The catecholaminergic receptors were found to be dynamically expressed on human HSPC, while G-CSF stimulations enhance their expression on primitive human CD34+CD38−/low cells (65). Acute stress, mimicked by norepinephrine stimulation actively induced rapid release of SDF-1 and subsequently HSPC to the blood stream. Treatment with β2-adrenergic antagonist was demonstrated to inhibit HSPC mobilization in both steady-state and following AMD3100 administration (inducing SDF-1 release to the peripheral blood) (66). Recent evidence suggests that endocannabinoids secreted by BM stromal cells, signaling through the CB2 receptor that is functionally expressed by human and murine HSPC, induced mobilization of murine HSPC with short- and long-term repopulating abilities. Moreover, G-CSF-induced mobilization of HSPC was significantly decreased by CB2 antagonists (67, 68). Bone remodeling processes, including bone formation by osteoblasts and degradation by osteoclasts, and HSPC mobilization are sequential events which are both controlled by β2-adrenergic signals. In particular, Vitamin D receptor that is controlled by β2adrenergic signals was found to be essential for G-CSF induce mobilization, leading to suppression of osteoblast activity and upregulation of RANKL (69). This complex picture of the dynamic

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“brain-bone-blood” triad may be exploited in future designing of the exact timing of clinical mobilization protocols (70) and therapeutic transplantation procedures. Of note, it is yet unknown how to guide a particular progenitor cell type (e.g., hematopoietic or endothelial) egress and mobilization from the BM. Better understanding of the factors involved in cell mobilization will pave the way for better regenerative therapies, including the use of enriched EPC for neovascularization as part of regaining organ function, enriched MSC to restore skeletal and muscle tissue damage, as well as the use of enriched HSC for hematopoietic recovery following high dose radiotherapy and chemotherapy.

4. Notes 1. Any other set of conjugated antibodies can be suitable for the HSPC quantification. 2. Since intracellular staining of DNA requires permeabilization, more cells per staining procedure are needed. 3. The migration index equals 100% migration capacity, and therefore it is a reference point for migrating cells in the transwell migration assay. 4. Duration time of the migration experiment should be considered based on cell-intrinsic motility properties. 2–4 h migration is recommended. 5. Do not heat the samples, since gelatin zymography examines proteolytic activity, and do not load more than 30 μg total protein per lane. 6. Do not exceed 30 min incubation, as prolonged incubation can damage the protease. 7. Cover the gel container to avoid vaporization of buffer. References 1. Schulz C, von Andrian UH, Massberg S (2009) Hematopoietic stem and progenitor cells: their mobilization and homing to bone marrow and peripheral tissue. Immunol Res 44:160–168 2. Lapidot T, Kollet O (2002) The essential roles of the chemokine SDF-1 and its receptor CXCR4 in human stem cell homing and repopulation of transplanted immune-deficient NOD/SCID and NOD/SCID/B2m(null) mice. Leukemia 16:1992–2003 3. Metcalf D (1990) The colony stimulating factors. Discovery, development, and clinical applications. Cancer 65:2185–2195

4. Lapid, K., Vagima, Y., Kollet, O., Lapidot, T. (2009). Egress and mobilization of hematopoietic stem and progenitor cells. In: StemBook Cambridge (MA): Harvard Stem Cell Institute. 5. Sugiyama T, Kohara H, Noda M, Nagasawa T (2006) Maintenance of the hematopoietic stem cell pool by CXCL12-CXCR4 chemokine signaling in bone marrow stromal cell niches. Immunity 25:977–988 6. Nie Y, Han YC, Zou YR (2008) CXCR4 is required for the quiescence of primitive hematopoietic cells. J Exp Med 205:777–783

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7. Zipori D, Sasson T (1980) Adherent cells from mouse bone marrow inhibit the formation of colony stimulating factor (CSF) induced myeloid colonies. Exp Hematol 8:816–817 8. Wilson A, Laurenti E, Trumpp A (2009) Balancing dormant and self-renewing hematopoietic stem cells. Curr Opin Genet Dev 19:461–468 9. Peled A, Kollet O, Ponomaryov T, Petit I, Franitza S, Grabovsky V, Slav MM, Nagler A, Lider O, Alon R, Zipori D, Lapidot T (2000) The chemokine SDF-1 activates the integrins LFA-1, VLA-4, and VLA-5 on immature human CD34(+) cells: role in transendothelial/stromal migration and engraftment of NOD/SCID mice. Blood 95:3289–3296 10. Avigdor A, Goichberg P, Shivtiel S, Dar A, Peled A, Samira S, Kollet O, Hershkoviz R, Alon R, Hardan I, Ben-Hur H, Naor D, Nagler A, Lapidot T (2004) CD44 and hyaluronic acid cooperate with SDF-1 in the trafficking of human CD34+ stem/progenitor cells to bone marrow. Blood 103:2981–2989 11. Petit I, Szyper-Kravitz M, Nagler A, Lahav M, Peled A, Habler L, Ponomaryov T, Taichman RS, Arenzana-Seisdedos F, Fujii N, Sandbank J, Zipori D, Lapidot T (2002) G-CSF induces stem cell mobilization by decreasing bone marrow SDF-1 and up-regulating CXCR4. Nat Immunol 3:687–694 12. Semerad CL, Christopher MJ, Liu F, Short B, Simmons PJ, Winkler I, Levesque JP, Chappel J, Ross FP, Link DC (2005) G-CSF potently inhibits osteoblast activity and CXCL12 mRNA expression in the bone marrow. Blood 106:3020–3027 13. Levesque JP, Hendy J, Winkler IG, Takamatsu Y, Simmons PJ (2003) Granulocyte colonystimulating factor induces the release in the bone marrow of proteases that cleave c-KIT receptor (CD117) from the surface of hematopoietic progenitor cells. Exp Hematol 31:109–117 14. Hattori K, Heissig B, Tashiro K, Honjo T, Tateno M, Shieh JH, Hackett NR, Quitoriano MS, Crystal RG, Rafii S, Moore MA (2001) Plasma elevation of stromal cell-derived factor-1 induces mobilization of mature and immature hematopoietic progenitor and stem cells. Blood 97:3354–3360 15. Broxmeyer HE, Orschell CM, Clapp DW, Hangoc G, Cooper S, Plett PA, Liles WC, Li X, Graham-Evans B, Campbell TB, Calandra G, Bridger G, Dale DC, Srour EF (2005) Rapid mobilization of murine and human hematopoietic stem and progenitor cells with AMD3100, a CXCR4 antagonist. J Exp Med 201: 1307–1318

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16. Pusic I, DiPersio JF (2010) Update on clinical experience with AMD3100, an SDF-1/ CXCL12-CXCR4 inhibitor, in mobilization of hematopoietic stem and progenitor cells. Curr Opin Hematol 17:319–326 17. Gazitt Y, Freytes CO, Akay C, Badel K, Calandra G (2007) Improved mobilization of peripheral blood CD34+ cells and dendritic cells by AMD3100 plus granulocyte-colony-stimulating factor in non-Hodgkin’s lymphoma patients. Stem Cells Dev 16:657–666 18. Kollet O, Dar A, Shivtiel S, Kalinkovich A, Lapid K, Sztainberg Y, Tesio M, Samstein RM, Goichberg P, Spiegel A, Elson A, Lapidot T (2006) Osteoclasts degrade endosteal components and promote mobilization of hematopoietic progenitor cells. Nat Med 12: 657–664 19. Kollet O, Dar A, Lapidot T (2007) The multiple roles of osteoclasts in host defense: bone remodeling and hematopoietic stem cell mobilization. Annu Rev Immunol 25:51–69 20. Spiegel A, Shivtiel S, Kalinkovich A, Ludin A, Netzer N, Goichberg P, Azaria Y, Resnick I, Hardan I, Ben-Hur H, Nagler A, Rubinstein M, Lapidot T (2007) Catecholaminergic neurotransmitters regulate migration and repopulation of immature human CD34+ cells through Wnt signaling. Nat Immunol 8:1123–1131 21. Levesque JP, Hendy J, Takamatsu Y, Williams B, Winkler IG, Simmons PJ (2002) Mobilization by either cyclophosphamide or granulocyte colony-stimulating factor transforms the bone marrow into a highly proteolytic environment. Exp Hematol 30:440–449 22. Levesque JP, Takamatsu Y, Nilsson SK, Haylock DN, Simmons PJ (2001) Vascular cell adhesion molecule-1 (CD106) is cleaved by neutrophil proteases in the bone marrow following hematopoietic progenitor cell mobilization by granulocyte colony-stimulating factor. Blood 98:1289–1297 23. Tesio, M., Golan, K., Corso, S., Giordano, S., Schajnovitz, A., Vagima, Y., Shivtiel, S., Kalinkovich, A., Caione, L., Gammaitoni, L., Laurenti, E., Buss, E. C., Shezen, E., Itkin, T., Kollet, O., Petit, I., Trumpp, A., Christensen, J., Aglietta, M., Piacibello, W., Lapidot, T. (2010) Enhanced c-Met activity promotes G-CSF induced mobilization of hematopoietic progenitor cells via ROS signaling. Blood 24. Levesque JP, Winkler IG (2008) Mobilization of hematopoietic stem cells: state of the art. Curr Opin Organ Transplant 13:53–58 25. Kopp HG, Avecilla ST, Hooper AT, Rafii S (2005) The bone marrow vascular niche: home of HSC differentiation and mobilization. Physiology (Bethesda) 20:349–356

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26. Christopher MJ, Liu F, Hilton MJ, Long F, Link DC (2009) Suppression of CXCL12 production by bone marrow osteoblasts is a common and critical pathway for cytokine-induced mobilization. Blood 114:1331–1339 27. Mendez-Ferrer S, Michurina TV, Ferraro F, Mazloom AR, Macarthur BD, Lira SA, Scadden DT, Ma’ayan A, Enikolopov GN, Frenette PS (2010) Mesenchymal and haematopoietic stem cells form a unique bone marrow niche. Nature 466:829–834 28. Pitchford SC, Furze RC, Jones CP, Wengner AM, Rankin SM (2009) Differential mobilization of subsets of progenitor cells from the bone marrow. Cell Stem Cell 4:62–72 29. Aicher A, Kollet O, Heeschen C, Liebner S, Urbich C, Ihling C, Orlandi A, Lapidot T, Zeiher AM, Dimmeler S (2008) The Wnt antagonist Dickkopf-1 mobilizes vasculogenic progenitor cells via activation of the bone marrow endosteal stem cell niche. Circ Res 103:796–803 30. Dimmeler S (2010) Regulation of bone marrow-derived vascular progenitor cell mobilization and maintenance. Arterioscler Thromb Vasc Biol 30:1088–1093 31. Brouard N, Driessen R, Short B, Simmons PJ (2010) G-CSF increases mesenchymal precursor cell numbers in the bone marrow via an indirect mechanism involving osteoclast-mediated bone resorption. Stem Cell Res 5:65–75 32. Purton LE, Scadden DT (2007) Limiting factors in murine hematopoietic stem cell assays. Cell Stem Cell 1:263–270 33. Spangrude GJ, Heimfeld S, Weissman IL (1988) Purification and characterization of mouse hematopoietic stem cells. Science 241: 58–62 34. Ikuta K, Weissman IL (1992) Evidence that hematopoietic stem cells express mouse c-kit but do not depend on steel factor for their generation. Proc Natl Acad Sci USA 89: 1502–1506 35. Yang L, Bryder D, Adolfsson J, Nygren J, Mansson R, Sigvardsson M, Jacobsen SE (2005) Identification of Lin(−)Sca1(+)kit(+)CD34(+) Flt3− short-term hematopoietic stem cells capable of rapidly reconstituting and rescuing myeloablated transplant recipients. Blood 105:2717–2723 36. Kiel MJ, Yilmaz OH, Iwashita T, Yilmaz OH, Terhorst C, Morrison SJ (2005) SLAM family receptors distinguish hematopoietic stem and progenitor cells and reveal endothelial niches for stem cells. Cell 121:1109–1121 37. Vagima Y, Avigdor A, Goichberg P, Shivtiel S, Tesio M, Kalinkovich A, Golan K, Dar A, Kollet O, Petit I, Perl O, Rosenthal E, Resnick I,

Hardan I, Gellman YN, Naor D, Nagler A, Lapidot T (2009) MT1-MMP and RECK are involved in human CD34+ progenitor cell retention, egress, and mobilization. J Clin Invest 119:492–503 38. Jalili A, Shirvaikar N, Marquez-Curtis LA, Turner AR, Janowska-Wieczorek A (2010) The HGF/c-Met axis synergizes with G-CSF in the mobilization of hematopoietic stem/progenitor cells. Stem Cells Dev 19:1143–1151 39. Yin T, Li L (2006) The stem cell niches in bone. J Clin Invest 116:1195–1201 40. Roberts AW, Metcalf D (1995) Noncycling state of peripheral blood progenitor cells mobilized by granulocyte colony-stimulating factor and other cytokines. Blood 86:1600–1605 41. Wright DE, Cheshier SH, Wagers AJ, Randall TD, Christensen JL, Weissman IL (2001) Cyclophosphamide/granulocyte colonystimulating factor causes selective mobilization of bone marrow hematopoietic stem cells into the blood after M phase of the cell cycle. Blood 97:2278–2285 42. Uchida N, He D, Friera AM, Reitsma M, Sasaki D, Chen B, Tsukamoto A (1997) The unexpected G0/G1 cell cycle status of mobilized hematopoietic stem cells from peripheral blood. Blood 89:465–472 43. Witko-Sarsat V, Canteloup S, Durant S, Desdouets C, Chabernaud R, Lemarchand P, Descamps-Latscha B (2002) Cleavage of p21waf1 by proteinase-3, a myeloid-specific serine protease, potentiates cell proliferation. J Biol Chem 277:47338–47347 44. Voermans C, Kooi ML, Rodenhuis S, van der Lelie H, van der Schoot CE, Gerritsen WR (2001) In vitro migratory capacity of CD34+ cells is related to hematopoietic recovery after autologous stem cell transplantation. Blood 97:799–804 45. Janowska-Wieczorek, A., Matsuzaki, A., L, A. M. (2000) The Hematopoietic Microenvironment: Matrix Metalloproteinases in the Hematopoietic Microenvironment. Hematology 4:515–527. 46. Everts V, Korper W, Jansen DC, Steinfort J, Lammerse I, Heera S, Docherty AJ, Beertsen W (1999) Functional heterogeneity of osteoclasts: matrix metalloproteinases participate in osteoclastic resorption of calvarial bone but not in resorption of long bone. FASEB J 13:1219–1230 47. Christopherson KW 2nd, Cooper S, Broxmeyer HE (2003) Cell surface peptidase CD26/ DPPIV mediates G-CSF mobilization of mouse progenitor cells. Blood 101:4680–4686 48. Levesque JP, Liu F, Simmons PJ, Betsuyaku T, Senior RM, Pham C, Link DC (2004)

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Characterization of hematopoietic progenitor mobilization in protease-deficient mice. Blood 104:65–72 49. Shirvaikar, N., Marquez-Curtis, L. A., Shaw, A. R., Turner, A. R., Janowska-Wieczorek, A. (2010) MT1-MMP association with membrane lipid rafts facilitates G-CSF-induced hematopoietic stem/progenitor cell mobilization. Exp Hematol. 50. Kent DG, Copley MR, Benz C, Wohrer S, Dykstra BJ, Ma E, Cheyne J, Zhao Y, Bowie MB, Zhao Y, Gasparetto M, Delaney A, Smith C, Marra M, Eaves CJ (2009) Prospective isolation and molecular characterization of hematopoietic stem cells with durable self-renewal potential. Blood 113:6342–6350 51. Lapidot T, Dar A, Kollet O (2005) How do stem cells find their way home? Blood 106: 1901–1910 52. Wright DE, Wagers AJ, Gulati AP, Johnson FL, Weissman IL (2001) Physiological migration of hematopoietic stem and progenitor cells. Science 294:1933–1936 53. Abkowitz JL, Robinson AE, Kale S, Long MW, Chen J (2003) Mobilization of hematopoietic stem cells during homeostasis and after cytokine exposure. Blood 102:1249–1253 54. Chen J, Larochelle A, Fricker S, Bridger G, Dunbar CE, Abkowitz JL (2006) Mobilization as a preparative regimen for hematopoietic stem cell transplantation. Blood 107:3764–3771 55. Hiramatsu H, Nishikomori R, Heike T, Ito M, Kobayashi K, Katamura K, Nakahata T (2003) Complete reconstitution of human lymphocytes from cord blood CD34+ cells using the NOD/SCID/gammacnull mice model. Blood 102:873–880 56. Leung W, Ramirez M, Civin CI (1999) Quantity and quality of engrafting cells in cord blood and autologous mobilized peripheral blood. Biol Blood Marrow Transplant 5:69–76 57. Pitchford SC, Hahnel MJ, Jones CP, Rankin SM (2010) Troubleshooting: quantification of mobilization of progenitor cell subsets from bone marrow in vivo. J Pharmacol Toxicol Methods 61:113–121 58. Teitelbaum SL (2010) Stem cells and osteoporosis therapy. Cell Stem Cell 7:553–554 59. Winkler IG, Sims NA, Pettit AR, Barbier V, Nowlan B, Helwani F, Poulton IJ, van Rooijen N, Alexander KA, Raggatt LJ, Levesque JP (2010) Bone marrow macrophages maintain hematopoietic stem cell (HSC) niches and their depletion mobilizes HSCs. Blood 116: 4815–4828 60. Lymperi, S., Ersek, A., Ferraro, F., Dazzi, F., Horwood, N. J. (2010) Inhibition of osteoclast

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function reduces hematopoietic stem cell numbers in vivo. Blood. 61. Levesque JP, Helwani FM, Winkler IG (2010) The endosteal ‘osteoblastic’ niche and its role in hematopoietic stem cell homing and mobilization. Leukemia 24:1979–1992 62. Jung, Y., Shiozawa, Y., Wang, J., Patel, L. R., Havens, A. M., Song, J., Krebsbach, P. H., Roodman, G. D., Taichman, R. S. (2010) Annexin-2 is a regulator of stromal cell-derived factor-1/CXCL12 function in the hematopoietic stem cell endosteal niche. Exp Hematol. 63. Kalinkovich A, Spiegel A, Shivtiel S, Kollet O, Jordaney N, Piacibello W, Lapidot T (2009) Blood-forming stem cells are nervous: direct and indirect regulation of immature human CD34+ cells by the nervous system. Brain Behav Immun 23:1059–1065 64. Mendez-Ferrer S, Lucas D, Battista M, Frenette PS (2008) Haematopoietic stem cell release is regulated by circadian oscillations. Nature 452:442–447 65. Lapidot, T., Kollet, O. (2010) The brain-boneblood triad: traffic lights for stem-cell homing and mobilization. Hematology 30th edition., 1–6. 66. Dar, A., Schajnovitz, A., Lapid, K., Kalinkovich, A., Itkin, T., Ludin, A., Kao, M., M., Battista, M., Tesio, M., Kollet, O., Netzer Cohen, N., Margalita, R., Buss, E., Baleux, F., Oishi, S., Fujii, N., Larochelle, A., Dunbar, C., Broxmeyer, H., Frenette, P., Lapidot, T. (2011) Rapid mobilization of hematopoietic progenitors by AMD3100 and catecholamines is mediated by CXCR4-dependent SDF-1 release from bone marrow stromal cells. Leukemia (in press) 67. Jiang, S., Alberich-Jorda, M., Zagozdzon, R., Parmar, K., Fu, Y., Mauch, P., Banu, N., Makriyannis, A., Tenen, D. G., Avraham, S., Groopman, J. E., Avraham, H. K. (2010) Cannabinoid receptor 2 and its agonists mediate hematopoiesis and hematopoietic stem and progenitor cell mobilization. Blood 68. Hoggatt J, Pelus LM (2010) Eicosanoid regulation of hematopoiesis and hematopoietic stem and progenitor trafficking. Leukemia 24: 1993–2002 69. Kawamori Y, Katayama Y, Asada N, Minagawa K, Sato M, Okamura A, Shimoyama M, Nakagawa K, Okano T, Tanimoto M, Kato S, Matsui T (2010) Role for vitamin D receptor in the neuronal control of the hematopoietic stem cell niche. Blood 116:5528–5535 70. Lucas D, Battista M, Shi PA, Isola L, Frenette PS (2008) Mobilized hematopoietic stem cell yield depends on species-specific circadian timing. Cell Stem Cell 3:364–366

Chapter 3 Hematopoietic Stem Cell Mobilization with G-CSF Chitra Hosing Abstract Cytokine mobilized peripheral blood stem cells are the preferred source of stem cells in autologous stem cell transplantation and have virtually replaced bone marrow as the stem cell source. In recent years, a dramatic increase has been reported in the use of peripheral blood stem cells for allogeneic transplantation as well. The reason for this rise is that peripheral blood stem cell transplants when compared to bone marrow transplants are associated with a more rapid recovery of granulocytes and platelets after transplantation and a lower regimen-related and transplant-related mortality. Peripheral blood stem cells can be easily harvested on an outpatient basis without the need for general anesthesia. In most cases peripheral blood stem cells are collected after G-CSF administration. In this chapter we describe peripheral blood stem cell mobilization in autologous transplant patients and in allogeneic donors using G-CSF. Key words: Hematopoietic stem cell mobilization, G-CSF, Autologous stem cell transplantation, Allogeneic stem cell transplantation

1. Introduction High-dose chemotherapy followed by autologous or allogeneic stem cell transplantation is used in the management of a variety of hematological and nonhematological malignancies. Stem cells were first described in the peripheral blood of mice back in 1962 (1) and in humans in 1971 (2). The subsequent development of apheresis instruments made it possible to collect peripheral blood stem cells (PBSC) (3, 4). Goldman and colleagues and Korbling et al. were among the first to demonstrate that PBSC collected from patients could reestablish normal marrow hematopoiesis after high-dose chemotherapy (5, 6). Use of PBSC for allogeneic stem cell transplantation was first reported in 1989 when PBSC were collected

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_3, © Springer Science+Business Media, LLC 2012

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38

C. Hosing

from an “unstimulated” donor who required ten apheresis procedures, to collect adequate cells. The recipient experienced trilineage engraftment (7). In 1993, Dreger et al. reported successful engraftment using allogeneic PBSC collected after mobilization with granulocyte-colony stimulating factor (G-CSF) in a patient who had engraftment failure after bone marrow (BM) transplantation from the same donor (8). Since mid-1990s PBSCs mobilized with cytokines have virtually replaced BM as the source of stem cells for autologous transplantation and are been increasingly used in allogeneic transplantation. The advantages of PBSC rather than BM are ease of collection and rapid engraftment. PBSC can be harvested without the need of general anesthesia and the discomfort of multiple BM aspirations. Furthermore, some studies have shown that PBSC restore immune functions more rapidly than BM (9, 10). Other studies have reported a lower incidence of documented infections, fewer febrile days, a lower number of red cell and platelet transfusion, lower demand for antibiotics and intensive care requirements resulting in reduced costs of PBSC transplants compared with BM (11). The other advantages of G-CSF mobilization are the safe outpatient self-application and the fixed-day apheresis. Filgrastim (granulocyte colony-stimulating factor (G-CSF)) and sargramostim (granulocyte macrophage colony-stimulating factor (GM-CSF)) are currently the only FDA-approved colonystimulating factors for stem cell mobilization (12, 13). In a study of 1,306 normal donors 99% of donors were mobilized with G-CSF (14). Filgrastim is a granulocyte-colony stimulating factor analog used to stimulate the proliferation and differentiation of granulocytes. It is produced by recombinant DNA technology. The gene for human GCSF is inserted into E. coli and the G-CSF produced closely resembles naturally produced G-CSF in humans. G-CSF was originally used to treat neutropenia and is now the cytokine of choice for increasing the number of hematopoietic stem cells in the blood before collection by leukapheresis for use in allogeneic and autologous stem cell transplantation. G-CSF stimulates the expansion and activation of myeloid and granulocyte precursors within the bone marrow, resulting in PBSC mobilization after a few days. Pegylated filgrastim is a covalent conjugate of G-CSF and monomethoxypolyethylene glycol, with a half-life of about 33 h (15). As a result of this conjugation there is decrease in the renal elimination of pegylated filgrastim resulting in adequate levels of G-CSF for approximately 2 weeks. Pegylated filgrastim is currently approved for the treatment of chemotherapy-induced neutropenia, but has been also utilized for stem cell mobilization because of its convenience (16).

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Hematopoietic Stem Cell Mobilization with G-CSF

39

2. Materials 1. Recombinant human G-CSF (Neupogen®, Amgen). 2. Pegylated filgrastim/pegfilgrastim (Neulasta®, Amgen).

3. Methods Grigg et al. evaluated the kinetics of mobilization by G-CSF in normal volunteers. G-CSF was injected subcutaneously at a dose of 3, 5, or 10 μg/kg/day. A subset of volunteers from each dose cohort underwent leukapheresis on study day 6 (after 5 days of G-CSF). Granulocyte-macrophage colony-forming cell (GM-CFC) numbers in the blood were maximal after 5 days of G-CSF, a broader peak was evident for CD34+ stem cells between days 4 and 6. The 95 % confidence intervals for mean number of PBSC per milliliter of blood in the three dose cohorts overlapped on each study day. However, on the peak day, CD34+ stem cells were significantly higher in the 10 μg/kg/day cohort than in a pool of the other two cohorts. Leukapheresis products obtained at the 10 μg/kg/day dose level contained a median GM-CFC number of 93 × 104/kg (range 50–172 × 104/kg). Collections from volunteers receiving lower doses of G-CSF contained a median GM-CFC number of 36 × 104/kg (range 5–204 × 104/kg). All leukapheresis products obtained at the 10 μg/kg/day dose level were potentially sufficient for allogeneic transplantation purposes. Thus, G-CSF 10 μg/kg/day for 5 days with a single leukapheresis on the following day is a highly effective regimen for PBSC mobilization and collection in normal donors (17). Molineux et al. studied the effects of single daily doses of pegfilgrastim (a PEGylated form of the recombinant human G-CSF) at 30, 60, 100, and 300 μg/kg (18). Successful CD34+ stem cell mobilization was observed at all doses. At the highest dose of 300 μg/kg a peak number of CD34+ cells/μl were seen at day 4. The numbers approached normal by day 12 or 13. In all other dose groups the peak was at the same time, but returned to normal range by day 9. In the same study mobilization of stem cells as measured by GM-CFC/ml was also noted with the maximum response seen at the 100 μg/kg dose level. There was no additional benefit to increasing the dose to 300 μg/kg (18–20). 3.1. Stem Cell Mobilization with G-CSF

1. G-CSF is generally given at a dose of 10 μg/kg/day subcutaneously and continued until the completion of apheresis. The half-life of G-CSF is only 3–4 h and therefore daily subcutaneous injections are required (11, 21–23) (see Note 1).

40

C. Hosing

2. The dose range for G-CSF that has been used varies from 10 μg/kg/day up to as high as 32 μg/kg/day, administered subcutaneously (24–29). 3. The optimal dose of G-CSF needed to collect enough CD34+ stem cells with minimal toxicity to the donors has not been established. 4. For G-CSF doses of up to 10 μg/kg/day, a dose–response relationship exists between the dose of G-CSF and the mobilization of CD34+ progenitor cells (30, 31). 5. G-CSF can be administered as a single daily injection or as twice-a-day injection (17, 32). 6. The total white blood cell count rises within 4–6 h but a substantial increase in circulating CD34+ stem cells does not occur until the third day after the first G-CSF injection (32). 7. Stem cell collection is usually started on the fourth or fifth day after the G-CSF injections begin. Most centers start collection when the circulating peripheral blood CD34 count is 10/μl (Fig. 1) (25–28). 8. The optimal stem cell dose for transplantation is not established, but most centers collect between 4 and 6 × 106 CD34+ cells/kg of the recipient’s body weight. The minimal acceptable dose is 2 × 106 CD34+ cells/kg of the recipient’s body weight (33). 9. G-CSF induces myeloid expansion, activation, and degranulation leading to release of neutrophil proteases in the marrow. The proteases cleave and inactivate some adhesive connections

Fig. 1. Efficacy of CD34+ mobilization and PBSC yield. Printed with permission from Holig et al. 2009 © American Society of Hematology.

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Hematopoietic Stem Cell Mobilization with G-CSF

41

like CXCR4/SDF-1α, VCAM-1/VLA-4 and facilitate stem cell mobilization (34). The down regulation of SDF-1α during G-CSF administration is beside the proteolytic activity caused also by the decrease of CXCL 12 transcription in stromal cells (35). G-CSF with chemotherapy mobilization is further associated with down regulation of expression of serine-proteinase inhibitors (α1antitrypsin, etc.), which under normal conditions block activity of serine proteases released by BM neutrophils. Together with this fact the down regulation facilitates an accumulation of proteases within the marrow microenvironment (36). 10. Almost all patients report some bone pain after growth factor administration. In most cases, the pain can be relieved by acetaminophen, and very rarely narcotics are required. The side effects are dose related and resolve within a few days of G-CSF discontinuation (31, 37, 38), see Table 1. 11. As summarized in Table 2, uncommon but severe side effects requiring discontinuation of G-CSF have been reported in 1–3% of donors (31, 38). G-CSF administration has been reported to precipitate serious and even life-threatening sickle cell crises in donors with hemoglobin SS, hemoglobin S ± beta

Table 1 Commonly reported symptoms associated with G-CSF administration Reference

Grigg (17) Anderlini (38) Bishop (26) Stroncek (51) Stroncek (51)

Dose of G-CSF (μg/kg/day) 10

12

5

5

10

41

19

21

–

53

76

83

32/11

68/0

44

74

67

0

27

–

–

No. of patients

15

341

Bone pain

87

84

Myalgias/arthralgias

27

Headache

33

Fever/flu-like symptoms

7

– 54

Chills/rigors

–

–

22

5

14

Body aches

–

–

–

–

–

Fatigue

47

31

–

37

43

Nausea/vomiting

–

13

–

16

24

Insomnia

–

–

–

16

24

Paresthesia

–

–

–

16

38

Diarrhea

–

–

–

11

5

Rash

–

–

–

11

5

42

C. Hosing

Table 2 Unusual and/or major adverse events reported during (or shortly after) G-CSF administration in normal donors Reference

Side effect

Nuamah et al. (52), Becker et al. (53)

Splenic rupture

de Azevedo and Tabak (54), Arimura et al. (55)

Capillary leak syndrome

Vij et al. (56)

Unstable angina

Bensinger et al. (57)

Myocardial infarctiona

Parkkali et al. (58), Huhn et al. (59)

Iritis, episcleritis

Storek et al. (60)

Flare-up of rheumatoid arthritis; ankylosing spondylitis

Spitzer et al. (61)

Acute gouty arthritis

Pei et al. (62)

Intracranial hemorrhage

Adkins (63)

Anaphylactoid reaction

Abboud et al. (39), Adler et al. (40), Grigg (41)

Sickle cell crisis in patients with hemoglobin SS, hemoglobin SC, or hemoglobin S ± β thalassemia

a

Donor had a prior history of coronary artery disease and myocardial infarction

thalassemia, or hemoglobin SC (39–41). This complication has not been reported in donors with the sickle cell trait (42) (see Note 2). 12. There is an increase in the white blood cell count after G-CSF administration, mostly due to the increase in the absolute neutrophil counts. The total white blood cell count may be as high as 70 or 80 × 109/l after 5 days of administration (43–45) (see Note 3). 13. There is a slight, but significant, decrease of platelet counts and hemoglobin levels after G-CSF administration, which persists for approximately 30 days after PBSC donation. 14. Transient elevations in the levels of lactate dehydrogenase, alkaline phosphatase, and alanine aminotransferase are seen after 4–5 days of G-CSF administration (31, 46). Levels of serum potassium, magnesium, and blood urea nitrogen decline minimally (31). Holig et al. found that leukocyte counts 4 weeks after PBSC collection were significantly lower than the white blood count on day 0 (P < .001). After 6 months, 1 year, and 5 years after PBSC donation, the white blood counts were higher than at 4 weeks but never completely returned to baseline values. Changes in absolute neutrophil counts resembled those in white blood count.

3

Hematopoietic Stem Cell Mobilization with G-CSF

43

Lymphocyte counts were significantly diminished up to 1 year after PBSC donation and were slightly elevated after 2–5 years. Platelet counts reached pretreatment values 6 months after apheresis and remained stable thereafter (47). Pulsipher et al. reviewed the NMDP follow-up data regarding the incidence of development of malignancies in this cohort of 2,408 donors. Annual attempts at follow-up were made for all donors (median follow-up, 49 months; range, 2 days to 99 months). No cases of acute myelogenous leukemia or myelodysplasia were reported. Twenty-five nonhematologic cancers of various types occurred along with one case of chronic lymphocytic leukemia. Comparison of the incidence of these cancers to expected rates according to the SEER database showed no evidence of increased cancer risk in the donor cohort (47). Others have reported similar results (14, 45, 48, 49). 3.2. Stem Cell Mobilization with Pegfilgrastim

In a prospective, phase II study of pegfilgrastim, administered as a single injection to mobilize autologous PBSC in patients with multiple myeloma, 19 patients received 12 mg pegfilgrastim. A median of 8.4 (range 4.1–15.8) × 106 CD34+ cells/kg could be collected. Sustained hematological recovery occurred in all the patients who underwent high-dose chemotherapy followed by autologous PBSC transplantation with pegfilgrastim-mobilized cells (20). Use of single-dose pegfilgrastim for the mobilization of allogeneic peripheral blood stem cells in healthy family and unrelated donors was described by Kroschinsky et al. (50). In their study 25 related or unrelated healthy donors received a single-dose of 12 mg pegfilgrastim for mobilization of allogeneic peripheral blood progenitor cells. In 80% of donors only a single apheresis procedure was necessary to reach the target progenitor cell dose. 1. Pegfilgrastim is also effective for mobilization of PBSC for collection by apheresis and is given as a single subcutaneous injection of 6 or 12 mg. 2. The efficacy and toxicity profile of pegfilgrastim is similar to that described with G-CSF treatment (20). 3. Bone pain, headaches and transient elevations of liver enzymes were the main adverse events (50).

3.3. Overcoming Inefficient Stem Cell Mobilization

There are no established mobilization strategies for poorly mobilizing patients. Some approaches that have been used include: 1. Increasing the dose of G-CSF to 32 μg/kg/day in two divided doses. 2. Adding plerixafor, GM-CSF, or other chemokines if available. 3. If failure to mobilize with cytokines alone, then remobilize with chemotherapy priming and cytokines.

44

C. Hosing

4. If any evidence for viral infection during mobilization, then remobilize after infection has resolved (viral infection may exert a myelosuppressive effect on the marrow). 5. Bone marrow harvest or in some instances G-CSF primed marrow can be used.

4. Notes 1. Most physicians will round the dose of G-CSF to the nearest vial size. G-CSF is available in the US in prefilled syringes of 300 and 480 μg. 2. G-CSF administration has been reported to precipitate lifethreatening sickle cell crises in donors with hemoglobin SS, hemoglobin S ± beta thalassemia, or hemoglobin SC. Therefore, G-CSF should be administered with great caution (if at all) to normal donors with any demonstrable hemoglobin S. 3. Although leukostasis has never been reported in donors, most physicians decrease the G-CSF dose when the WBC is higher than 70 or 75 × 109/l. References 1. Goodman JW, Hodgson GS (1962) Evidence for stem cells in the peripheral blood of mice. Blood 19:702–714 2. McCredie KB, Hersh EM, Freireich EJ (1971) Cells capable of colony formation in the peripheral blood of man. Science 171:293–294 3. Weiner RS, Richman CM, Yankee RA (1977) Semicontinuous flow centrifugation for the pheresis of immunocompetent cells and stem cells. Blood 49:391–397 4. Hillyer CD, Tiegerman KO, Berkman EM (1991) Increase in circulating colony-forming units-granulocyte-macrophage during largevolume leukapheresis: evaluation of a new cell separator. Transfusion 31:327–332 5. Korbling M, Dorken B, Ho AD, Pezzutto A, Hunstein W, Fliedner TM (1986) Autologous transplantation of blood-derived hemopoietic stem cells after myeloablative therapy in a patient with Burkitt’s lymphoma. Blood 67:529–532 6. Goldman JM (1979) Autografting cryopreserved buffy coat cells for chronic granulocytic leukaemia in transformation. Exp Hematol 7(Suppl 5):389–397 7. Kessinger A, Smith DM, Strandjord SE, Landmark JD, Dooley DC, Law P, Coccia PF, Warkentin PI, Weisenburger DD, Armitage JO

8.

9.

10.

11.

(1989) Allogeneic transplantation of bloodderived, T cell-depleted hemopoietic stem cells after myeloablative treatment in a patient with acute lymphoblastic leukemia. Bone Marrow Transplant 4:643–646 Dreger P, Suttorp M, Haferlach T, Loffler H, Schmitz N, Schroyens W (1993) Allogeneic granulocyte colony-stimulating factor-mobilized peripheral blood progenitor cells for treatment of engraftment failure after bone marrow transplantation. Blood 81:1404–1407 Storek J, Dawson MA, Storer B, Stevens-Ayers T, Maloney DG, Marr KA, Witherspoon RP, Bensinger W, Flowers ME, Martin P, Storb R, Appelbaum FR, Boeckh M (2001) Immune reconstitution after allogeneic marrow transplantation compared with blood stem cell transplantation. Blood 97:3380–3389 Bensinger WI, Martin PJ, Storer B, Clift R, Forman SJ, Negrin R, Kashyap A, Flowers ME, Lilleby K, Chauncey TR, Storb R, Appelbaum FR (2001) Transplantation of bone marrow as compared with peripheral-blood cells from HLA-identical relatives in patients with hematologic cancers. N Engl J Med 344:175–181 Schmitz N, Linch DC, Dreger P, Goldstone AH, Boogaerts MA, Ferrant A, Demuynck

3

12. 13. 14.

15. 16.

17.

18.

19.

20.

Hematopoietic Stem Cell Mobilization with G-CSF

HM, Link H, Zander A, Barge A (1996) Randomised trial of filgrastim-mobilised peripheral blood progenitor cell transplantation versus autologous bone-marrow transplantation in lymphoma patients. Lancet 347:353–357 Product information. Leukine (sargrastim). W.B.H. and Pharmaceuticals, A, Seattle Product information. Neupagen (filgrastim). Amgen Inc., Thousand Oaks, CA Anderlini P, Rizzo JD, Nugent ML, Schmitz N, Champlin RE, Horowitz MM (2001) Peripheral blood stem cell donation: an analysis from the International Bone Marrow Transplant Registry (IBMTR) and European Group for Blood and Marrow Transplant (EBMT) databases. Bone Marrow Transplant 27:689–692 Curran MP, Goa KL (2002) Pegfilgrastim. Drugs 62:1207–1213, discussion 1214–1205 Isidori A, Tani M, Bonifazi F, Zinzani P, Curti A, Motta MR, Rizzi S, Giudice V, Farese O, Rovito M, Alinari L, Conte R, Baccarani M, Lemoli RM (2005) Phase II study of a single pegfilgrastim injection as an adjunct to chemotherapy to mobilize stem cells into the peripheral blood of pretreated lymphoma patients. Haematologica 90:225–231 Grigg AP, Roberts AW, Raunow H, Houghton S, Layton JE, Boyd AW, McGrath KM, Maher D (1995) Optimizing dose and scheduling of filgrastim (granulocyte colony-stimulating factor) for mobilization and collection of peripheral blood progenitor cells in normal volunteers. Blood 86:4437–4445 Molineux G, Kinstler O, Briddell B, Hartley C, McElroy P, Kerzic P, Sutherland W, Stoney G, Kern B, Fletcher FA, Cohen A, Korach E, Ulich T, McNiece I, Lockbaum P, MillerMessana MA, Gardner S, Hunt T, Schwab G (1999) A new form of filgrastim with sustained duration in vivo and enhanced ability to mobilize PBPC in both mice and humans. Exp Hematol 27:1724–1734 Johnston E, Crawford J, Blackwell S, Bjurstrom T, Lockbaum P, Roskos L, Yang BB, Gardner S, Miller-Messana MA, Shoemaker D, Garst J, Schwab G (2000) Randomized, dose-escalation study of SD/01 compared with daily filgrastim in patients receiving chemotherapy. J Clin Oncol 18:2522–2528 Hosing C, Qazilbash MH, Kebriaei P, Giralt S, Davis MS, Popat U, Anderlini P, Shpall EJ, McMannis J, Korbling M, Champlin RE (2006) Fixed-dose single agent pegfilgrastim for peripheral blood progenitor cell mobilisation in patients with multiple myeloma. Br J Haematol 133:533–537

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21. Nademanee A, Sniecinski I, Schmidt GM, Dagis AC, O’Donnell MR, Snyder DS, Parker PM, Stein AS, Smith EP, Molina A et al (1994) High-dose therapy followed by autologous peripheral-blood stem-cell transplantation for patients with Hodgkin’s disease and nonHodgkin’s lymphoma using unprimed and granulocyte colony-stimulating factor-mobilized peripheral-blood stem cells. J Clin Oncol 12:2176–2186 22. Gazitt Y, Freytes CO, Callander N, Tsai TW, Alsina M, Anderson J, Holle L, Cruz J, Devore P, McGrath M, West G, Alvarez R, Montgomery W (1999) Successful PBSC mobilization with highdose G-CSF for patients failing a first round of mobilization. J Hematother 8:173–183 23. Bensinger WI, Price TH, Dale DC, Appelbaum FR, Clift R, Lilleby K, Williams B, Storb R, Thomas ED, Buckner CD (1993) The effects of daily recombinant human granulocyte colony-stimulating factor administration on normal granulocyte donors undergoing leukapheresis. Blood 81:1883–1888 24. Anderlini P, Przepiorka D, Champlin R, Korbling M (1996) Peripheral blood stem cell apheresis in normal donors: the neglected side. Blood 88:3663–3664 25. Waller CF, Bertz H, Wenger MK, Fetscher S, Hardung M, Engelhardt M, Behringer D, Lange W, Mertelsmann R, Finke J (1996) Mobilization of peripheral blood progenitor cells for allogeneic transplantation: efficacy and toxicity of a high-dose rhG-CSF regimen. Bone Marrow Transplant 18:279–283 26. Bishop MR, Tarantolo SR, Jackson JD, Anderson JR, Schmit-Pokorny K, Zacharias D, Pavletic ZS, Pirruccello SJ, Vose JM, Bierman PJ, Warkentin PI, Armitage JO, Kessinger A (1997) Allogeneic-blood stem-cell collection following mobilization with low-dose granulocyte colony-stimulating factor. J Clin Oncol 15:1601–1607 27. Sato N, Sawada K, Takahashi TA, Mogi Y, Asano S, Koike T, Sekiguchi S (1994) A time course study for optimal harvest of peripheral blood progenitor cells by granulocyte colonystimulating factor in healthy volunteers. Exp Hematol 22:973–978 28. Bensinger WI, Weaver CH, Appelbaum FR, Rowley S, Demirer T, Sanders J, Storb R, Buckner CD (1995) Transplantation of allogeneic peripheral blood stem cells mobilized by recombinant human granulocyte colony-stimulating factor. Blood 85:1655–1658 29. Kroger N, Zeller W, Fehse N, Hassan HT, Kruger W, Gutensohn K, Lolliger C, Zander AR (1998) Mobilizing peripheral blood stem cells with high-dose G-CSF alone is as effective

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30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

C. Hosing as with Dexa-BEAM plus G-CSF in lymphoma patients. Br J Haematol 102:1101–1106 Hoglund M, Smedmyr B, Simonsson B, Totterman T, Bengtsson M (1996) Dosedependent mobilisation of haematopoietic progenitor cells in healthy volunteers receiving glycosylated rHuG-CSF. Bone Marrow Transplant 18:19–27 Stroncek DF, Clay ME, Petzoldt ML, Smith J, Jaszcz W, Oldham FB, McCullough J (1996) Treatment of normal individuals with granulocyte-colony-stimulating factor: donor experiences and the effects on peripheral blood CD34+ cell counts and on the collection of peripheral blood stem cells. Transfusion 36:601–610 Link H, Arseniev L, Bahre O, Berenson RJ, Battmer K, Kadar JG, Jacobs R, Casper J, Kuhl J, Schubert J, Diedrich H, Poliwoda H (1995) Combined transplantation of allogeneic bone marrow and CD34+ blood cells. Blood 86:2500–2508 Wuchter P, Ran D, Bruckner T, Schmitt T, Witzens-Harig M, Neben K, Goldschmidt H, Ho AD (2010) Poor mobilization of hematopoietic stem cells-definitions, incidence, risk factors, and impact on outcome of autologous transplantation. Biol Blood Marrow Transplant 16:490–499 Winkler IG, Levesque JP (2006) Mechanisms of hematopoietic stem cell mobilization: when innate immunity assails the cells that make blood and bone. Exp Hematol 34:996–1009 Semerad CL, Christopher MJ, Liu F, Short B, Simmons PJ, Winkler I, Levesque JP, Chappel J, Ross FP, Link DC (2005) G-CSF potently inhibits osteoblast activity and CXCL12 mRNA expression in the bone marrow. Blood 106:3020–3027 Winkler IG, Hendy J, Coughlin P, Horvath A, Levesque JP (2005) Serine protease inhibitors serpina1 and serpina3 are down-regulated in bone marrow during hematopoietic progenitor mobilization. J Exp Med 201:1077–1088 Anderlini P, Przepiorka D, Champlin R, Korbling M (1996) Biologic and clinical effects of granulocyte colony-stimulating factor in normal individuals. Blood 88:2819–2825 Anderlini P, Donato M, Chan KW, Huh YO, Gee AP, Lauppe MJ, Champlin RE, Korbling M (1999) Allogeneic blood progenitor cell collection in normal donors after mobilization with filgrastim: the M.D. Anderson Cancer Center experience. Transfusion 39:555–560 Abboud M, Laver J, Blau CA (1998) Granulocytosis causing sickle-cell crisis. Lancet 351:959 Adler BK, Salzman DE, Carabasi MH, Vaughan WP, Reddy VV, Prchal JT (2001) Fatal sickle

41.

42.

43.

44.

45.

46.

47.

48.

49.

cell crisis after granulocyte colony-stimulating factor administration. Blood 97:3313–3314 Grigg AP (2001) Granulocyte colony-stimulating factor-induced sickle cell crisis and multiorgan dysfunction in a patient with compound heterozygous sickle cell/beta + thalassemia. Blood 97:3998–3999 Kang EM, Areman EM, David-Ocampo V, Fitzhugh C, Link ME, Read EJ, Leitman SF, Rodgers GP, Tisdale JF (2002) Mobilization, collection, and processing of peripheral blood stem cells in individuals with sickle cell trait. Blood 99:850–855 Anderlini P, Korbling M, Dale D, Gratwohl A, Schmitz N, Stroncek D, Howe C, Leitman S, Horowitz M, Gluckman E, Rowley S, Przepiorka D, Champlin R (1997) Allogeneic blood stem cell transplantation: considerations for donors. Blood 90:903–908 Stroncek DF, Clay ME, Herr G, Smith J, Ilstrup S, McCullough J (1997) Blood counts in healthy donors 1 year after the collection of granulocyte-colony-stimulating factor-mobilized progenitor cells and the results of a second mobilization and collection. Transfusion 37:304–308 Pulsipher MA, Chitphakdithai P, Miller JP, Logan BR, King RJ, Rizzo JD, Leitman SF, Anderlini P, Haagenson MD, Kurian S, Klein JP, Horowitz MM, Confer DL (2009) Adverse events among 2408 unrelated donors of peripheral blood stem cells: results of a prospective trial from the National Marrow Donor Program. Blood 113:3604–3611 Anderlini P, Przepiorka D, Seong D, Miller P, Sundberg J, Lichtiger B, Norfleet F, Chan KW, Champlin R, Korbling M (1996) Clinical toxicity and laboratory effects of granulocyte-colony-stimulating factor (filgrastim) mobilization and blood stem cell apheresis from normal donors, and analysis of charges for the procedures. Transfusion 36:590–595 Holig K, Kramer M, Kroschinsky F, Bornhauser M, Mengling T, Schmidt AH, Rutt C, Ehninger G (2009) Safety and efficacy of hematopoietic stem cell collection from mobilized peripheral blood in unrelated volunteers: 12 years of single-center experience in 3928 donors. Blood 114:3757–3763 Miflin G, Charley C, Stainer C, Anderson S, Hunter A, Russell N (1996) Stem cell mobilization in normal donors for allogeneic transplantation: analysis of safety and factors affecting efficacy. Br J Haematol 95:345–348 Cavallaro AM, Lilleby K, Majolino I, Storb R, Appelbaum FR, Rowley SD, Bensinger WI (2000) Three to six year follow-up of normal donors who received recombinant human

3

50.

51.

52.

53.

54.

55.

56.

Hematopoietic Stem Cell Mobilization with G-CSF

granulocyte colony-stimulating factor. Bone Marrow Transplant 25:85–89 Kroschinsky F, Holig K, Poppe-Thiede K, Zimmer K, Ordemann R, Blechschmidt M, Oelschlaegel U, Bornhauser M, Rall G, Rutt C, Ehninger G (2005) Single-dose pegfilgrastim for the mobilization of allogeneic CD34+ peripheral blood progenitor cells in healthy family and unrelated donors. Haematologica 90:1665–1671 Stroncek DF, Clay ME, Jaszcz W, Lennon S, Smith J, McCullough J (1999) Collection of two peripheral blood stem cell concentrates from healthy donors. Transfus Med 9:37–50 Nuamah NM, Goker H, Kilic YA, Dagmoura H, Cakmak A (2006) Spontaneous splenic rupture in a healthy allogeneic donor of peripheral-blood stem cell following the administration of granulocyte colony-stimulating factor (g-csf). A case report and review of the literature. Haematologica 91:ECR08 Becker PS, Wagle M, Matous S, Swanson RS, Pihan G, Lowry PA, Stewart FM, Heard SO (1997) Spontaneous splenic rupture following administration of granulocyte colony-stimulating factor (G-CSF): occurrence in an allogeneic donor of peripheral blood stem cells. Biol Blood Marrow Transplant 3:45–49 de Azevedo AM, Goldberg Tabak D (2001) Life-threatening capillary leak syndrome after G-CSF mobilization and collection of peripheral blood progenitor cells for allogeneic transplantation. Bone Marrow Transplant 28:311–312 Arimura K, Inoue H, Kukita T, Matsushita K, Akimot M, Kawamata N, Yamaguchi A, Kawada H, Ozak A, Arima N, Te C (2005) Acute lung injury in a healthy donor during mobilization of peripheral blood stem cells using granulocyte-colony stimulating factor alone. Haematologica 90:ECR10 Vij R, Adkins DR, Brown RA, Khoury H, DiPersio JF, Goodnough T (1999) Unstable

57.

58.

59.

60.

61.

62.

63.

47

angina in a peripheral blood stem and progenitor cell donor given granulocyte-colony-stimulating factor. Transfusion 39:542–543 Bensinger WI, Buckner CD, Rowley S, Storb R, Appelbaum FR (1996) Treatment of normal donors with recombinant growth factors for transplantation of allogeneic blood stem cells. Bone Marrow Transplant 17(Suppl 2):S19–S21 Parkkali T, Volin L, Siren MK, Ruutu T (1996) Acute iritis induced by granulocyte colonystimulating factor used for mobilization in a volunteer unrelated peripheral blood progenitor cell donor. Bone Marrow Transplant 17:433–434 Huhn RD, Yurkow EJ, Tushinski R, Clarke L, Sturgill MG, Hoffman R, Sheay W, Cody R, Philipp C, Resta D, George M (1996) Recombinant human interleukin-3 (rhIL-3) enhances the mobilization of peripheral blood progenitor cells by recombinant human granulocyte colony-stimulating factor (rhG-CSF) in normal volunteers. Exp Hematol 24:839–847 Storek J, Glaspy JA, Grody WW, Susi E, Slater ED (1993) Adult-onset cyclic neutropenia responsive to cyclosporine therapy in a patient with ankylosing spondylitis. Am J Hematol 43:139–143 Spitzer T, McAfee S, Poliquin C, Colby C (1998) Acute gouty arthritis following recombinant human granulocyte colonystimulating factor therapy in an allogeneic blood stem cell donor. Bone Marrow Transplant 21:966–967 Pei RZ, Ma JX, Zhang PS, Liu XH, Cao JJ, Du XH (2008) Intracranial hemorrhage caused by cerebrovascular malformation after donation of rhG-CSF-primed allogeneic PBSC. Bone Marrow Transplant 42:61–62 Adkins DR (1998) Anaphylactoid reaction in a normal donor given granulocyte colonystimulating factor. J Clin Oncol 16:812–813

Chapter 4 Hematopoietic Stem Cell Mobilization with Agents Other than G-CSF Jonathan Hoggatt and Louis M. Pelus Abstract Hematopoietic stem and progenitor mobilization has revolutionized the field of hematopoietic transplantation. Currently, hematopoietic grafts acquired from the peripheral blood of patients or donors treated with granulocyte-colony stimulating factor (G-CSF) are the preferred source for transplantation. G-CSF mobilization regimens, however, are associated with known morbidities and a significant number of normal donors and patient populations fail to mobilize sufficient numbers of hematopoietic stem and progenitor cells for transplantation, necessitating the need for non-G-CSF mobilization strategies. Mechanistic studies evaluating hematopoietic bone marrow niche interactions have uncovered novel agents with the capacity for hematopoietic mobilization. This chapter provides a comprehensive overview of mobilizing agents, other than G-CSF, and experimental procedures and technical aspects important to evaluate and define their hematopoietic mobilizing activities alone and in combination. Key words: Mobilization, Hematopoietic stem cells, AMD3100, GRO beta, VLA-4 inhibitor, Fucoidan, BIO5192, CXCR4, SDF-1, G-CSF

1. Introduction Allogeneic hematopoietic stem cell transplantation (HCT) is a curative option for many patients with hematological malignancies. The source of stem cells used for transplant can have a significant impact on patient outcome. In spite of a higher incidence of chronic graft-versus-host disease (GVHD) observed with mobilized peripheral blood hematopoietic stem cell grafts (1–3), studies comparing granulocyte-colony stimulating factor (G-CSF)-mobilized peripheral blood stem cells (PBSC) to bone marrow have shown that PBSC are associated with more rapid engraftment, reduction in infectious complications, and in patients with advanced

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_4, © Springer Science+Business Media, LLC 2012

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malignancies, lower regimen-related mortality (4–6). In many centers PBSC are now the preferred hematopoietic stem cell source used for human leukocyte antigen (HLA)-identical sibling as well as matched related and unrelated donor transplantation (7, 8). G-CSF, granulocyte macrophage-colony stimulating factor (GM-CSF), and more recently plerixafor (AMD3100), for patients who fail to mobilize with a G- or GM-CSF, are the only FDA approved agents for mobilizing autologous PBSC, and G-CSF is the preferred mobilizing agent in the allogeneic setting. G-CSF, however, is associated with morbidity in the form of bone pain that may result in absence from work and disruption of lifestyle for the donor during the mobilization process. Furthermore, G-CSF has also been associated with serious, albeit rare, toxicity, including splenic rupture, in normal donors (9–12). Methods of mobilizing PBSC that avoid the use of growth factors such as G-CSF are, therefore, of great interest.

2. The Hematopoietic Niche Control of hematopoietic proliferation and differentiation is highly complex, and homeostatic balance is likely maintained by both intrinsic and genetic cues within individual cells and extrinsic cues from the supportive microenvironment in which hematopoietic stem cells (HSC) reside. HSC reside in very defined and limited microenvironments, or “niches” (13), and signals within these niches direct HSC maintenance. In mammals, the primary HSC niche is contained within the bone marrow, which is comprised of stromal cells and an extracellular matrix of collagens, fibronectin, and proteoglycans (14). Recent studies have shown that osteoblasts within the endosteal bone marrow niche are a significant regulatory component of hematopoiesis (15–18). Within the niche, HSCs are thought to be “tethered” to osteoblasts, other stromal cells, and the extracellular matrix through a variety of adhesion molecule interactions, many of which are likely redundant systems. Early studies exploring the role of osteoblasts in maintaining HSCs suggested that N-cadherin interactions mediated the positive effects on HSCs (16); however, more recent studies have contradicted these findings (19, 20). Numerous other adhesion molecules have been implicated as contributing to HSC and HPC tethering, including, but not limited to, the integrins α4β1—very late antigen-4 (VLA-4) (21–26), α5β1—very late antigen-5 (VLA5) (22, 23, 25, 27), α4β7—lymphocyte Peyer’s patch adhesion molecule-1 (LPAM-1) (28), the alpha 6 integrins (Laminins) (29, 30), CD44 (22, 31), E-selectins (32–34), the angiopoietin receptor tyrosine kinase with immunoglobulin-like and EGF-like

4

Hematopoietic Stem Cell Mobilization with Agents Other than G-CSF

51

domains-2 (Tie-2) (18), osteopontin (OPN) (35, 36), endolyn (CD164) (37), and the calcium-sensing receptor (CaR) (38). The most explored niche interaction, and perhaps the most important in regulating HSC and HPC trafficking to and from the marrow niche, is the interaction between the CXC chemokine receptor 4 (CXCR4) and its ligand stromal cell-derived factor-1α (SDF-1α). SDF-1α is produced by osteoblasts (39), and has also been found on endothelial cells and within bone itself (40, 41). HSC and hematopoietic progenitor cells (HPC) express CXCR4 and are chemo-attracted to and retained within the bone marrow by SDF-1α (42–44). Under steady state conditions, HSC and HPC normally reside within the bone marrow niches, while the mature cells ultimately exit the marrow and enter the peripheral blood. However, considerable evidence over the last several decades demonstrates that HSC and HPC also traffic to the peripheral blood (45–50), and this steady state trafficking leaves open niche spaces that can be repopulated by transplanted HSC (51). Based on observations that increased HPC were found in patients after chemotherapy (52, 53), we now know that this natural egress of HSC and HPC into the periphery can be enhanced, allowing for “mobilization” of these cells to the peripheral blood (47, 48). Mobilized adult HSC and HPC are widely used for autologous and allogeneic transplantation and have improved patient outcomes compared to bone marrow. Mobilization can be achieved through administration of chemotherapy (52–54), or hematopoietic growth factors, chemokines, or small molecule inhibitors or antibodies against chemokine receptors and integrins. Procedures to monitor hematopoietic mobilization induced by G-CSF have been described in other chapters in this book. Many of the methods and procedures to monitor hematopoietic stem and progenitor cells are not unique to specific mobilizing agents and can be applied to mobilization experiments exploring novel agents. This remainder of this chapter will describe current compounds and procedures for non-G-CSF mediated PBSC mobilization, with a focus on unique aspects and procedures relevant to these agents and rapid mobilizers.

3. Mobilization by Agents Other Than G-CSF 3.1. Agents That Disrupt CXCR4/ SDF-1a

Many agents capable of mobilizing HSC and HPC act through mechanisms that disrupt the CXCR4/SDF-1α axis. Most notably, the CXCR4 antagonist AMD3100 (Plerixafor; Mozobil™) mobilizes HSC and HPC (55–60) and received FDA approval in December 2008 for use in combination with G-CSF for patients with Non-Hodgkin’s lymphoma and multiple myeloma. In addition to AMD3100, the CXCR4 antagonists T140 (61) and T134

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(62) are both capable of mobilization. Hematopoietic mobilization has also been reported after administration of CXCR4 partial agonists, including (met)-SDF-1β (63), CTCE-0214 (64), and CTCE-0021 (60), through desensitization and reduced surface expression of the CXCR4 receptor. A number of polymeric compounds have been reported to mobilize HSC and HPC including Betafectin (65, 66), sulfated polysaccharides (Fucoidan) (67–69), sulfated colominic acid (70), and the smaller glycosaminoglycan (GAG) mimetics (71), which appear to alter plasma SDF-1α levels (69–71), enhance matrix metalloproteinase-9 (MMP-9) production (65, 68, 71), increase CXCR4 receptor function on HPC (70), and perhaps affect selectin and other adhesion molecules through undefined mechanisms. The complement system, particularly the C3a peptide fragment of the third complement system, has been reported to increase sensitivity of HSC and HPC to SDF-1α (72), which acts to counteract normal mobilization responses that reduce marrow SDF-1α. An inhibitor of the C3a receptor, SB 290517, when used in combination with G-CSF reduces this increased sensitivity to SDF-1α and results in an enhancement in mobilization, marked by a reduced requirement for G-CSF (73). 3.2. Other Mobilization Agents

As previously described, HSC and HPC are tethered within the bone marrow through adhesion interactions within the niche. The mobilization agents described thus far mechanistically function by disrupting one of these interactions, the CXCR4/SDF-1α axis; however, disruption of other HSC/niche interactions presents further targets for hematopoietic mobilization strategies. Targeting the interaction between VLA-4 and VCAM-1 with either antibodies against VLA-4 (24, 74), antibodies against VCAM-1 (75, 76), or a small molecule inhibitor of VLA-4 (BIO5192) (77), results in hematopoietic mobilization. Recently, signaling through the Ephephrin A3 axis was shown to increase adhesion to fibronectin and VCAM-1, and disruption of this signaling axis in vivo with a soluble EphA3-Fc fusion protein results in hematopoietic mobilization (78). Defibrotide, an adenosine receptor agonist, has been reported to reduce expression of the adhesion molecules P-selectin (79) and intercellular adhesion molecule-1 (ICAM-1) (80), and in vivo administration along with G-CSF enhances mobilization (81). The CXCR2 agonist GROβΔ4 has been shown to mobilize HSC and HPC, with peak mobilization occurring 15 min post administration (82–84), due primarily to a rapid increase in MMP-9 activity. GROβΔ4 can synergistically increase mobilization by G-CSF, reduce the requirement for G-CSF, and mobilizes HSC with enhanced engraftment and long-term repopulating abilities. In contrast to CXCR4, CXCR2 is not expressed on HSC and HPC, rather mobilization is mediated via protease release from

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neutrophils (84, 85), demonstrating that mobilization agents can target nonhematopoietic cells to illicit mobilization responses. While the agents discussed thus far have targeted receptors or ligands at the cell surface, particularly the CXCR4/SDF-1α pathway, mobilization strategies have also targeted intracellular signaling pathways downstream of CXCR4 and other receptors. Pertussis toxin (Ptx) is an inhibitor of the Gαi G-protein, which is coupled to the CXCR4 receptor, and in vivo administration results in abrogation of response to SDF-1α, and mobilization (86). Similarly, the Rho GTPase, Rac1, is a mediator of the downstream signaling pathways of CXCR4 and β integrins, and administration of a small molecule inhibitor of Rac1, NSC23766, has been reported to mobilize HSC and HPC (87).

4. Mobilization Agent Administration and Blood Collection 4.1. Dosing and Kinetics of Administration 4.2. Evaluating Combinations of Agents

While mobilization by G-CSF typically requires 4–6 days of administration to achieve optimal mobilization, a common characteristic of non-G-CSF mobilization agents is that they are considerably more rapid. Table 1 represents the optimal dose and route of administration for agents reported to mobilize hematopoietic progenitor cells, and the time post administration that has been reported to result in peak mobilization. The rapid kinetics of response and variation in mobilization mechanisms make exploration of combination treatment regimens highly attractive to evaluate for potential synergy in response. However, when exploring combination treatment, several different dosing regimens should be attempted to fully explore possible synergistic activity. As an example, we explored the combination of AMD3100 with GROβΔ4. The peak mobilization with AMD3100 is at 60 min post administration, while the peak for GROβΔ4 is 15 min. Therefore, we hypothesized that if GROβΔ4 was given 45 min post administration of AMD3100, and blood was collected 15 min later, allowing for blood collection at the peak time for each agent on its own, that the maximum mobilization of the combination would be achieved. However, this dosing regimen did not show any synergy and at best resulted in only additive mobilization of HPC (Fig. 1). However, if both AMD3100 and GROβΔ4 are given at the same time, and blood is collected 15 min post administration, a significant synergy in mobilization was observed, that persisted for >2 h. Thus, optimum kinetics of a mobilization agent on its own may not be the optimal kinetics when used in combination with other agents. Similarly, some compounds, based on molecular mechanism of action, may be theorized to enhance mobilization; however, the compounds may only work in combination with

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Table 1 Mobilization agent dosing and kinetics of administration Mobilization agent

Regimen

Route

Bleeding time (post last dose)

References

AMD3100

5 mg/kg

SC

1h

(55, 56)

T140

5 mg/kg

SC

2h

(61)

T134

10 mg/kg

SC

1h

(62)

(met)-SDF-1β

300 μg/mouse

IV

48 h

(63)

CTCE-0214

75 μg/mouse

IV

4h

(64)

CTCE-0021

25 mg/kg

SC

1h

(60)

Betafectin (PGG-β glucan) Fucoidan

2 mg/kg 9.6 mg/kg 100 mg/kg × 3 days 25 mg/kg × 6 doses in 48 h

IV IV IV IP

30 min 24 h 3h 2h

(66) (65) (68, 69) (67)

Sulfated colominic acid

100 mg/kg

IV

30 min

(70)

GAG mimetics

50 mg/kg

IP

3h

(71)

SB290157*

500 ng/mouse × 3 days

IP

6h

(73)

Anti-CD49d(VLA-4)

2 mg/kg × 3 days

IV

~24 h

(24, 74)

Anti-VCAM-1

5 mg/kg × 2 days 2 mg/kg × 3 days

IV IV

6h ~24 h

(75) (76)

BIO5192

1 mg/kg

IV

1h

(77)

300 μg/mouse

IP

30 min

(78)

Defibrotide

15 mg/mouse × 5 days

IP

2h

(81)

GROβΔ4

2.5 mg/kg

SC

15 min

(82, 83)

Pertussis toxin

100 ng i.v.

IV

96 h

(86)

NSC23766

2.5 mg/kg

IP

6h

(87)

EphA3-Fc a

SC subcutaneous, IV intravenous, IP intraperitoneal a Mobilization response only seen when used in combination with a G-CSF regimen

another mobilizing agent (as is the case with Defibrotide and SB290157). In some circumstances, it may be advantageous to perform mobilization assays in combination with G-CSF or another agent, even if an experimental agent failed to mobilize on its own. 4.3. Peripheral Blood Collection

Peripheral blood should be collected at the peak of mobilization. There are two important parameters to consider when designing and performing mobilization experiments with rapid mobilizing agents: (1) it has been reported that circadian rhythms can affect

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Fig. 1. Evaluation of AMD3100 and GROβΔ4 combination treatment. Mice were treated with AMD3100 (5 mg/kg, SC), GROβΔ4 (2.5 mg/kg, SC), or the combination of the two agents, either staggered (as shown), or simultaneously, and peripheral blood was assayed for CFU-GM content at the indicated time points post treatment.

hematopoietic trafficking and mobilization (88, 89). In our laboratory, we typically time all of our mobilization experiments so that bleeding begins at about 10:00 a.m., which in our facility is ~4 h after initiation of light. Variations in bleeding time, if not controlled, can make interpretation of results from experiment to experiment difficult. (2) Many of the agents described, particularly GROβΔ4, exhibit rapid peaks (15 min) in peripheral HSC and HPC that return to baseline within an hour post administration. Therefore, in experiments with many mice, it is important to stagger the dosing of the mobilization agent and the acquisition of peripheral blood, such that blood collection can be performed at the peak mobilization time. In the case of GROβΔ4 we typically treat a cage of 3–5 mice approximately every 10 min, allowing for enough time in-between cages for blood collection and injections in the next set of mice. In addition, a permanent marker can be used to mark the tail of each mouse in a cage to keep track of the order of injections. Our laboratory normally collects peripheral blood using a cardiac puncture technique; however, other techniques may be suitable (i.e., retroorbital). After collection of blood, a complete blood count is performed using either a veterinary cell counter (Hemavet 950FS, Drew Scientific; or similar

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instrument), or white blood cell count determined manually with a hemocytometer. For analysis of HSC and HPC mobilization, we isolate low density mononuclear cells (LDMC) before additional assays using the Lympholyte®-Mammal (Cedarlane Labs, Burlington, NC, #CL5120) gradient separation technique.

5. Determination of HPC and HSC Mobilization 5.1. Progenitor Cell Analysis: Colony Assays

5.2. Competitive Transplantation Assays

Numerous in vitro colony-forming cell assays are available to identify populations of HPC with distinct lineage-restricted differentiation patterns and can be characterized by the type of colonies they form in semi-solid agar, methylcellulose or plasma clot. HPC can be identified as colony-forming unit-granulocyte (CFU-G), colonyforming unit-monocyte/macrophage (CFU-M), colony forming unit-granulocyte/macrophage (CFU-GM), burst-forming uniterythroid (BFU-E) or the colony-forming unit-erythroid (CFUE). Additionally, megakaryocyte progenitor cells (CFU-Mk or CFU-Meg), and progenitors with multipotential have been described, the most common one assayed today referred to as a colony-forming unit granulocyte/erythrocyte/monocyte/megakaryocyte (CFU-GEMM) (90–94). Typically our laboratory utilizes the semi-solid agar CFU-GM assay to screen mobilization agents and regimens as we have previously described (83, 95). In some experiments, multiple colony types, including CFU-GM, BFU-E and CFU-GEMM are enumerated in 1% methylcellulose containing erythropoietin (EPO), GM-CSF and stem cell factor (SCF) (82, 96). The reader is referred to Chapter 3 of this book for further details on progenitor cell assays. Initially, the colony forming unit-spleen (CFU-S) assay (97) was believed to measure HSC, and is still used by many investigators today as a surrogate HSC assay. Other surrogate assays commonly used to imply HSC function include the cobblestone area-forming cell (CAFC) assay (98–100) and the long-term culture-initiating cell (LTC-IC) assay (101–103). While these assays may certainly be indicative of more immature populations of cells than detected in CFU assays, they are not definitive assays for HSC function (104, 105). The only true measure of HSC function is the ability to fully repopulate a lethally irradiated host. By this definition, the “presence” of HSC could be determined just by monitoring survival of lethally irradiated transplant recipients. If the mice live (longer than 16 weeks), with reconstitution of all blood lineages, then by definition, the graft is considered to have contained HSC. However, this strict “survival” method does not allow for the ability to quantify HSC number or function, and limits the ability to

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compare HSC grafts, and therefore the HSC mobilizing capacity of a given mobilizer or combination of agents. To address this problem, various types of long-term repopulation assays, which assess long-term repopulating cells (LTRC), an HSC synonym, with comparison against a “competitor” graft were developed. The standard competitive HSC repopulation assay was first described by Harrison (106) followed by description of a calculation for competing repopulating units (RU) (107) that are one measure used to enumerate HSC. In this assay, a donor HSC graft is admixed with a competing bone marrow graft from a congenic, wild-type mouse, and the mixture is transplanted into a lethally irradiated recipient. Markers distinct for the donor graft and the competitor graft are then used to distinguish blood production from each source of cells, allowing for a comparison of the repopulating ability of each. The standard method of employing this technique today uses the C57Bl/6 (CD45.2) mouse and the B6.SJLPtrcAPep3B/BoyJ (BOYJ) (CD45.1) mouse. These congenic strains of mice only differ at the CD45 antigen, and can be distinguished with specific monoclonal antibodies, allowing for assessment of chimerism in recipient animals (108). A variation of this assay is the limiting-dilution competitive repopulation assay, in which a series of dilutions of the donor, or “test” graft, is compared to a standard number of competing cells (normally 2 × 105 whole bone marrow cells). A minimum threshold of peripheral blood cell (or bone marrow) reconstitution is set (~2–5 %) and the number of mice that do not reconstitute with the test graft is determined and the frequency of competitive repopulating units (CRU), or HSC, contained within the test graft determined by Poisson statistics (109–111). It has recently been suggested by Drs. Purton and Scadden that a nomenclature distinction between RU and CRU should be made to describe the above transplantation assays (112); however, to date, CRU is still commonly used in both instances. 5.3. Transplantation Assays with Mobilized Blood

Our laboratory utilizes these competitive transplantation assays to validate the presence of LTRC in a mobilized peripheral blood product and assess the relative quantity and function of HSC contained in the graft. We use LDMC from a Lympholyte®-Mammal separation and compare these cells in ratios to 2 × 105 whole bone marrow competitor cells. It should be noted that LDMC from mobilized mice, depending on the mobilization regimen used, are considerably less competitive than bone marrow. Therefore, higher ratios of cells should be used, particularly in limiting dilution analysis. Smaller pilot studies evaluating the relative competitiveness of LDMC from mobilized donors is often advantageous to establish appropriate donor–competitor ratios. Typically, mobilized LDMC– competitor ratios of 1:1 to 5:1 are evaluated.

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5.4. Serial Transplantation

Competitive transplantation assays are routinely analyzed 12–16 weeks post transplant, and if multilineage peripheral blood reconstitution is seen at this time point, it is assumed that HSC were transplanted. However, it is becoming increasingly clear that HSCs are a heterogeneous cell population with varying capacities for self-renewal, and as a consequence, varying capacities for extended repopulation. Early studies analyzing CFU-S after serial transplantation hinted at a reduction in self-renewal ability following multiple transplants (113), and serial transplantation was used by others to assess the potential of “younger” HSC (114–116). It was found that in normal mice, the ability of HSC to self-renew is lost after four or five serial transplantations (115). Recently, experimental evidence indicates the presence of three classes of HSC that differ in the ability to self-renew and the capacity for multipotent differentiation into all blood lineages: short-term HSC (ST-HSC) capable of full reconstitution for up to 16 weeks, intermediateterm HSC (IT-HSC) capable of full reconstitution for up to 32 weeks, and long-term HSC (LT-HSC) capable of reconstitution for longer than 32 weeks and/or through serial transplantation (117). In light of these various potentials for self-renewal, the most stringent test of HSC potential, specifically the LT-HSC, is serial transplantation from primary recipients into secondary recipients, or beyond.

5.5. Flow Cytometric Methods

So far, the discussion on stem and progenitor identity has focused on experimental assays to determine HSC and HPC that are all direct or indirect measures of the functional ability of these cells; whether the ability to form lineage specific colonies in media, or repopulation of lethally irradiated recipients. In addition to these functional assays, immunophenotypic analysis is commonly used to determine the number or frequency of HSC and HPC, and used as a means to “sort” specific populations for further experimentation. Immunophenotypic analysis utilizes antigen specific antibodies coupled with fluorescent labels and fluorescence-activated cell sorting (FACS) that is able to rapidly enumerate and/or collect specific cell populations. Early work on immunophenotyping hematopoietic cell populations demonstrated that mature B cells and their immediate precursors could be defined by a specific antibody (118), which has lead to a set of lineage markers (Lin) to define mature blood cells including erythrocytes, granulocytes, macrophages, T-cells, B-cells, natural killer (NK) cells and megakaryocytes, and lineage negative cells that are enriched for earlier stem and progenitor populations (119). Later, it was demonstrated that repopulating cells could be further defined by the absence of lineage markers (Linneg) with expression of stem cell antigen-1 (Sca1) and low expression of Thy1.1 (120). An additional marker for the stem cell factor (SCF) receptor (c-kit) (121–123) was later added to further define the murine HSC population. These cells

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will be referred to as Sca-1+ c-kit+ Linneg (SKL) cells. Although enriched for HSC function, SKL cells are still heterogeneous. Additional markers have recently been identified to further enrich for HSCs, including CD34 (124), and fms-related tyrosine kinase-3 (Flt3) (125, 126), which allow for the characterization of LT-HSC (CD34− Flt3− SKL), ST-HSC (CD34+ Flt3− SKL) and multipotent progenitors (MPPs) (CD34+ Flt3+ SKL) (125). While CD34 may be an appropriate marker to distinguish short-term and long-term HSC at steady state, G-CSF mobilized HSC express CD34, and revert to CD34− when back at steady state (127), perhaps reducing the reliability of CD34 when evaluating other mobilization regimens. Several additional markers have now been identified that further refine HSC identity, including Endoglin (CD105) (128, 129), Tie2 (CD202) (18), endothelial protein C receptor (CD201) (130), CD49b (117), and notably the signaling lymphocyte activation molecule (SLAM) family of receptors CD150, CD48, and CD244 (131, 132). Defining an HSC population as CD150+ CD48− SKL, which is highly enriched for LT-HSC (133), has been routinely used by our laboratory to evaluate a wide array of mobilization agents with success. However, interpretation of phenotypic analysis for HSC content in mobilized grafts should always be cautious in the absence of transplantation data to verify HSC function. For detailed methods on flow cytometry techniques to identify HSC, the reader is referred to (134–136).

6. The Need for Transplantation Assays

Without HSC activity in a mobilized graft, the graft will fail to fully reconstitute a myeloablated host long-term, ultimately leading to graft failure and mortality. In most cases, in vitro progenitor assays or flow cytometric analysis are appropriate for broad characterization and optimization of mobilization regimens and correlate with HSC mobilization. However, we have observed two specific instances where this has not held true. We evaluated the ability of 2 × 106 peripheral blood LDMC from mice mobilized with G-CSF, GROβΔ4, an alternate CXCR2 ligand GROγ, and the CXCR4 partial agonist CTCE-0021 to rescue a lethally irradiated recipient, all of which were able to significantly mobilize CFU-GM to peripheral blood compared to vehicle control (Table 2). However, even though both CTCE-0021 and GROγ mobilized equivalent amounts of CFU-GM compared to GROβΔ4, they failed to mobilize sufficient numbers of HSC to rescue lethally irradiated recipients, demonstrating the importance of transplantation assays to validate HSC content in mobilized grafts. Thus, one cannot rely solely on in vitro progenitor cell assays or flow cytometry to predict HSC mobilization. While mobilized products containing HPC but

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Table 2 Mobilization of CFU-GM versus Survival in Lethally Irradiated Host Mobilizing agent

CFU-GM/ml Blood

Survival

Vehicle G-CSF GROβΔ4 GROg

34±2 1749±711 386±48 421±72

0/5 5/5 5/5 1/5

CTCE-0021

386±161

0/5

Mice were treated with G-CSF (50µg/mouse, bid, SC, 4 days), GROβΔ4 (2.5 mg/kg, SC), GROg (2.5 mg/kg, SC), or CTCE-0021 (25 mg/kg, SC), and LDMC and femur flushes collected. Whole bone marrow from femur flushes were plated in semi-solid agar for CFU-GM colony assays, and 2x106 LDMC were transplanted into 5 lethally irradiated (1100 cGy, split dose) mice. Shown are the number of mice, out of 5, which survived >16 weeks post-transplant

not HSC may have clinical utility in some cases, validation of HSC content is imperative if the graft is intended for hematopoietic reconstitution in a myeloablated host.

7. Conclusion While G-CSF mobilized hematopoietic grafts have revolutionized hematopoietic cell transplantation, there still remains a need for alternatives and improvements. The procoagulant effects of G-CSF increase the risk of myocardial infarction and cerebral ischemia in high-risk individuals (137, 138). G-CSF is contraindicated in patients with Sickle Cell Disease, owing to its potential to precipitate sickle crisis (139, 140), which has a negative impact on the potential utility of using G-CSF mobilized blood for adult HSC gene therapy for these patients. Poor mobilization in response to G-CSF occurs in 25 % of patients, particularly those with lymphoma and multiple myeloma (141) and 15 % of normal donors (142), requiring extended aphereses (143). In addition, the incidence of chronic GVHD is higher (1–3) for G-CSF-mobilized PBSC than bone marrow. Hence, there continues to be a need to search for additional safe and effective mobilizing agents to expand the use of hematopoietic grafts and PBSC transplantation. Multiple agents already identified and others to be identified in the future may provide these alternatives. Their potential use, however, depends on exacting characterization of their effects and function. The procedures we have described are a guide we have found useful to evaluate HPC and HSC mobilization.

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Chapter 5 Hematopoietic Stem Cell Mobilization: A Clinical Protocol Gina Pesek and Michele Cottler-Fox Abstract Autologous hematopoietic stem cell transplantation is the standard treatment for a wide variety of malignancies. At present, most hematopoietic progenitor/stem cell (HPC) collections are collected from the peripheral blood via leukapheresis following chemotherapy and/or growth factor-mediated mobilization. Most mobilization regimens consist of chemotherapy followed by one or more growth factors such as G-CSF, GM-CSF, or plerixafor. Occasionally a subset of patients will prove unable to mobilize effectively and will not collect at least 2.0 × 106 CD34+ cells/kg, the number of HPC currently considered to be appropriate for transplant in order to achieve timely engraftment and recovery of hematopoiesis. When this occurs it may be necessary to either remobilize, possibly with a different method, or to do a marrow harvest. Recent research has explored the benefits of using HPC outside of the oncology arena, notably in the area of cardiac regeneration following infarction, making the subject of mobilization potentially important to many areas of medicine. Key words: Mobilization, HPC, Apheresis, Growth factors, Clinical practice

1. Introduction Harvest of hematopoietic progenitor cells is primarily accomplished today via leukapheresis due to increased cell yields and improved patient comfort over bone marrow harvest, although it is possible this may change as other progenitor cell populations more prevalent in marrow than in blood (endothelial progenitor cells and mesenchymal stromal cells) become clinically important. Mobilization of HPC from marrow into blood is influenced by many factors but may best be predicted in the myeloma population by age, prior treatment, and platelet count (1, 2). Mobilization can be accomplished with a variety of agents;

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_5, © Springer Science+Business Media, LLC 2012

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however, chemotherapy used in conjunction with growth factors has been shown to be more effective than either alone (3). Granulocyte colony-stimulating factor (G-CSF), granulocytemacrophage colony-stimulating factor (GM-CSF), and plerixafor are currently the agents most commonly used in the USA. Other mobilizing growth factors include erythropoietin (mobilizes CD34+ endothelial progenitors), GROb/CXCL2 (under investigation) and stem cell factor (not approved for use in the USA). The three clinical protocols used most commonly at our institution, chemotherapy plus G-CSF, G-CSF alone or with GM-CSF, and G-CSF plus plerixafor with or without preceding chemotherapy, are detailed within this chapter.

2. Materials 2.1. Administration of Growth Factors

1. Syringe. 2. Sub-q needle. 3. Alcohol swabs. 4. Nonsterile gloves. 5. Band-Aid. 6. Sharps container. 7. Ambulatory pump tubing with 0.22 μm inline filter.

2.2. Mobilization with G-CSF With or Without GM-CSF

1. G-CSF dosed at 5 mcg/kg body weight BID (see Note 1).

2.3. Mobilization with VDT-PACE Chemotherapy Plus G-CSF

1. G-CSF dosed at 5 mcg/kg body weight BID.

2. GM-CSF dosed at 250 mcg QD.

2. Cisplatin (P), Cyclophosphamide (C), and Etoposide (E). 1 L prepared in Normal Saline (NS). 3. Dexamethasone (D) and Thalidomide (T)—outpatient oral prescriptions. 4. Doxorubicin (A) Prepared in NS. 5. Bortezomib (V).

2.4. Mobilization with G-CSF Plus Plerixafor

1. G-CSF dosed at 5 mcg/kg body weight BID. 2. Plerixafor dosed at 240 mcg/kg/day, not to exceed 40 mg/ day (see Note 2).

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3. Methods The choice of mobilization method is dependent on multiple factors, including whether the donor is autologous or allogeneic, the disease state, and results of previous attempts at mobilization. For example, a patient who has failed to mobilize with chemotherapy plus G-CSF may obtain adequate HPC with the addition of plerixafor to the regimen. A discussion of mobilization would not be complete without addressing the question “Is the regimen working?” Beginning collection on a fixed day after administration of chemotherapy and/or growth factors is possible, but optimal timing of collection can be predicted based on enumeration of circulating CD34+ cells in the donor’s peripheral blood (4). Peripheral blood CD34+ cells may be monitored daily by flow cytometry using one of two commercially available single platform systems, ProCount (Becton-Dickinson, Mt. View, CA) Note (3) or StemKit (Beckman-Coulter, Fullerton, CA), or by using the ISHAGE method and an automated cell counter (a dual platform system) which evaluates cells based on surface expression of CD34. The single platform tests allow direct comparison of results between institutions, a feature not always possible with standard flow cytometry (5). While some centers use this result as the primary criterion for when to start collection (usually set at between 5 and 20 CD34+ cells/μL), other institutions use the result as part of a predictive formula which gives an estimation of expected number of CD34+ cells which may be collected: Blood volume processed (L) × (CD34+ cells/μL × machine collection efficiency)/Patient weight (kg). Currently a common goal of apheresis is to provide a graft for transplant of 2–4 × 106 CD34+ cells/kg in a single apheresis (6, 7). Another method of enumeration now in use is the Sysmex automated cell counter (Kobe, Japan), which uses an HPC window (the Immature Information channel or IMI) based on cell size, density, and lysis resistance to predict optimal collection times (8). Studies comparing the HPC counts of the Sysmex and flow cytometry for CD34 expression have shown predominantly favorable results using the Sysmex system (8, 9), although the Sysmex HPC number does not work in the predictive formula above ( Cottler-Fox, personal observation). Finally, since the above methods are based on cell surface marker expression, and since it is recognized that not all HPC express surface CD34 (10), the intracellular enzymatic activity of HPC has become a target of interest. A recently developed commercially available system (Aldagen, Durham, NC) employs staining with Aldecount® reagent to assess intracytosolic aldehyde dehydrogenase (ALDH) activity, present in both CD34+ and CD34− cells (11).

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3.1. Administration of Growth Factors

1. Verify order on the patient medication record, including dosage, frequency, and route of administration. 2. Verify the patient’s identity by asking name and date of birth. 3. Explain procedure and potential side effects to patient (see Note 4). 4. Gather materials and medication needed. 5. Inspect medication, ensuring that there is no cloudiness or discoloration. 6. Wash hands. 7. Aseptically draw up medication according to dosage ordered. Change to Sub-q needle. 8. Choose injection site. 9. Put on gloves. 10. Cleanse site with alcohol and allow to dry. 11. Remove needle cap ensuring that needle remains sterile. 12. Administer growth factor. 13. Grasp skin around injection site with nondominant hand, forming a 1 in. fat fold. 14. Position needle with bevel up. 15. Tell the patient that he/she will feel a prick. 16. Insert needle quickly in one motion at 45 or 90° angle and depress plunger. 17. Remove needle gently and quickly at same angle used for insertion. 18. Dispose of needle in sharps container. 19. If oozing, place Band-Aid over injection site. 20. Remove the gloves and wash hands.

3.2. Mobilization Using G-CSF With or Without GM-CSF (see Note 5)

Autologous HPC Collection: 1. Informed consent is obtained by the physician (or designee) prior to the administration of growth factor (GF). 2. Patients undergoing autologous HPC mobilization have a thorough physical, laboratory, radiologic, and serologic screening prior to receiving GF for mobilization. Donor qualification and infectious disease testing results must be documented according to the institutional SOP and per AABB/FACT/JACIE standards (FACT Standard C6 Donor, Evaluation, and Management; AABB Standard 5.10 Donor evaluation). 3. Pregnancy test must be done on all female donors who are of childbearing potential. Contraception and/or barrier protection should be used while patient is undergoing this process.

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4. The initial dose of G-CSF for individuals who have not undergone prior attempts at HPC mobilization is 5 mcg/kg BID (see Note 1). Patients who have mobilized poorly previously or patients who are at significant risk for poor mobilization may receive G-CSF at a higher dose (approximately 8 mcg/ kg/day) at the discretion of the physician (see Note 1). 5. A decision may be made at this time about use of GM-CSF in addition to G-CSF. Our institutional protocol uses 250 mcg of GM-CSF QD × 3 days prior to initiation of G-CSF. 6. G-CSF is supplied by the manufacturer in 300 and 480 mcg vials. The dose of G-CSF is rounded to the nearest vial size. 7. The initial dose of GF is typically administered in the outpatient clinic area. Outpatient nursing staff educate the donor in selfadministration of the growth factor if insurance allows this. 8. Donors undergo vascular access device placement if their veins are not adequate to support collection. Our apheresis protocol is for large volume leukapheresis, which means that the vein or catheter must tolerate a flow rate of 125–150 mL/ min. (see Note 6). On the 5th day of G-CSF administration, a complete blood count, blood chemistry panel, ionized calcium and HPC blood CD34 quantification are obtained on autologous donors. For best results using the Terumo (formerly Cobe) Spectra apheresis device, hematocrit should be at least 27% (personal observation, M. Cottler-Fox). Lab work will be reviewed on day 5 of G-CSF and daily thereafter to determine start date of HPC collection based upon predictive formula. HPC mobilization may be interrupted at the discretion of the physician if the yield is poor. If mobilization with G-CSF alone is poor, the autologous donor is assessed for collection with alternate growth factors or chemotherapy. 3.3. Mobilization with VDT-PACE Chemotherapy Plus G-CSF 3.3.1. Autologous HPC Collection

1. Informed consent is obtained and insurance screening is performed prior to the administration of VTD-PACE for HPC mobilization. 2. Patients undergoing autologous HPC mobilization undergo a thorough physical, laboratory, radiologic, and serologic screening prior to receiving GF for mobilization. Donor qualification and infectious disease testing results must be documented according to the institutional SOP and per AABB/FACT/ JACIE standards (FACT Standard C6 Donor, Evaluation, and Management; AABB Standard 5.10 Donor evaluation). 3. Pregnancy test must be done on all female donors who are of childbearing potential. Contraception and/or barrier protection should be used while patient is undergoing this process.

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3.3.2. Eligibility Criteria

1. Patients with Multiple Myeloma ((MM) responsive to chemotherapy, primary refractory disease, or patients with recurrent disease), selected patients with plasmacytomas, Waldenstrom macroglobulinemia, light-chain amyloid (AL), and non-Hodgkin lymphoma (NHL) that meet published criteria for treatment of MM and/or HPC mobilization. 2. Capacity to provide informed consent. 3. Physiologic age >18 years and <75 years. 4. Zubrod <2, unless performance is solely due to symptoms of MM-related bone disease. 5. No active infections or sepsis that would preclude the use of this regimen. 6. No history of anaphylaxis or severe allergic reaction to any of the components of therapy. 7. No other medical or psychosocial problems, which in the opinion of the primary physician would place the patient at unacceptably high risk from this treatment regimen.

3.3.3. Exclusion Criteria

1. Pregnancy. 2. Concurrent, uncontrolled life-threatening bacterial or invasive fungal infection.

3.3.4. Mobilization with G-CSF

1. Patients receive appropriate antiemetics 30–60 min prior to chemotherapy dosing on each day and given daily until 2 days after the last dose of chemotherapy. 2. Based upon the patient’s allergy history and the discretion of the primary physician, patients undergoing VDT-PACE therapy may receive prophylactic antibiotic, antiviral, and antifungal therapy post chemotherapy until neutrophil engraftment (WBC >1,000 mm3). 3. G-CSF typically starts 6 days after chemotherapy and continues until mobilization and collection is completed. 4. The chemotherapy doses listed below in steps 8–12 are based upon calculated body weight (CBW) unless subject weighs less than 60 kg. If less than 60 kg, then actual weight will be used to calculate chemotherapy doses. In addition, body surface area (BSA) based on CBW is capped at 2 m2. 5. On days 1–4, patients receive dexamethasone 40 mg by mouth 30 min before chemotherapy. 6. On days 1–4, patients receive thalidomide 100 or 200 mg by mouth. (Dose may be determined by primary physician). Before starting chemotherapy, the nurse needs to ensure that patient has filled the thalidomide prescription.

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7. On days 1, 4, 8, and 11 patients receive bortezomib 1 mg/m2, which is given IVP 8. On days 1–4 patients receive cyclophosphamide 400 mg/m2, etoposide 40 mg/m2, and cisplatin 10 mg/m2 as intravenous continuous infusion over 24h. The daily cyclophosphamide, etoposide and cisplatin are mixed in a liter of normal saline for injection to be infused via ambulatory pump. Pump tubing is attached to the intravenous solution and partially primed by the Pharmacy. 9. On days 1–4 patients receives doxorubicin 10 mg/m2 intravenously administered as a continuous infusion over 24h via ambulatory pump. Pump tubing is attached to the intravenous solution and partially primed by the Pharmacy. 10. If the patient has a high tumor burden they may need tumor lysis prophylaxis, hydration fluids and other medications. 11. When the WBC count becomes ≥2.0 × 103/mm3 a complete blood count, blood chemistry panel, ionized calcium and HPC blood CD34 quantification are obtained (see Note 3). Lab work is reviewed daily thereafter by apheresis to determine start date of HPC collection based upon the predictive formula. 3.4. Mobilization with G-CSF Plus Plerixafor

1. Informed consent is obtained by the physician prior to administration of G-CSF and plerixafor. 2. Pregnancy test is performed on all patients who are of childbearing potential. Contraception and/or barrier protection should be used while patient is undergoing this process. 3. The expected side effects of G-CSF and plerixafor are discussed with the patient prior to administering initial dose of these agents. 4. The dose of G-CSF is 5 mcg/kg twice daily, starting on day 1, and continued until apheresis is completed. G-CSF will be administered subcutaneously in two divided daily doses as per Subheading 3.1 above. G-CSF is supplied by the manufacturer in 300 and 480 mcg vials. The dose of G-CSF will be rounded to the nearest vial size. At the discretion of the physician, G-CSF dose may be increased up to a maximum dose of 8 mcg/kg BID. 5. The dose of plerixafor is 240 mcg/kg/day (not to exceed 40 mg/day), which starts on day 4 of G-CSF at 5 p.m. (see Note 2). It is administered for a maximum of four daily doses. Plerixafor will be administered subcutaneously, as per Subheading 3.1 above. If the estimated e-GFR is ≤50 mL/ min, the dose of plerixafor is 160 mcg/kg/day (not to exceed

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27 mg/day), starting on day 4 of G-CSF and given for a maximum of four doses. Plerixafor is available as a single-use vial containing 1.2 mL (20 mg/mL) solution. Plerixafor doses >1.2 mL are administered as two subcutaneous injections. 6. Patients undergo vascular device placement on day 4 of G-CSF. 7. On Day 5 of G-CSF administration, patient reports for HPC collection. A complete blood count, blood chemistry panel, ionized calcium and HPC blood CD34 quantification are obtained on the patient. 8. Mobilization of HPC may be interrupted at the discretion of the physician if HPC mobilization is poor.

4. Notes 1. Standards may vary among institutions. 2. Dosing of plerixafor during clinical trials occurred with a 10h interval between dose and collection (10 p.m. dose for 8 a.m. collection); however, we have shown improved collection with a 15h dosing interval (12) (5 p.m. dose for 8 a.m. collection). 3. We use the ProCount test (Becton Dickinson, San Jose, California) and the test is not validated for a WBC less than 2.0. 4. The most commonly reported side effect with colony stimulating factors is bone pain, and other adverse effects include headache, fever, myalgia and nausea. Serious but rare events include splenic rupture, acute respiratory distress syndrome, exacerbation of autoimmune conditions and sickle cell crisis (13, 14). The most commonly reported adverse effects of plerixafor include diarrhea, nausea/vomiting, fatigue, headache, dizziness, arthralgia, and injection site reactions (15). 5. We do not use GM-CSF in amyloid patients as we have seen significant adverse reactions to the drug in this population. 6. Choice of central venous catheter is based on the flow rate needed for the procedure. Institutions not performing large volume leukapheresis may use slower inlet flow rates than we use and choose their catheters accordingly.

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References 1. Tricot G, Jagannath S, Vesole D et al (1995) Peripheral blood stem cell transplants for multiple myeloma: identification of favorable variables for rapid engraftment in 225 patients. Blood 85:588–596 2. Morris CL, Siegel E, Barlogie B et al (2003) Mobilization of CD34+ cells in elderly patients (>/= 70 years) with multiple myeloma: influence of age, prior therapy, platelet count and mobilization regimen. Br J Haematol 120:413–423 3. Bensinger W, Appelbaum F, Rowley S et al (1995) Factors that influence collection and engraftment of autologous peripheral-blood stem cells. J Clin Oncol 13:2547–2555 4. Rosenbaum ER, O’Connell B, Cottler-Fox M (2012) Validation of a formula for predicting daily CD34(+) cell collection by leukapheresis. Cytotherapy 14(4):461–466 5. Rivadeneyra-Espinoza L, Perez-Romano B, Gonzalez-Flores A et al (2006) Instrumentand protocol-dependent variation in the enumeration of CD34+ cells by flow cytometry. Transfusion 46:530–536 6. Bender JG, To LB, Williams S et al (1992) Defining a therapeutic dose of peripheral blood stem cells. J Hematother 1:329–341 7. Weaver CH, Hazelton B, Birch R et al (1995) An analysis of engraftment kinetics as a function of the CD34 content of peripheral blood progenitor cell collections in 692 patients after the administration of myeloablative chemotherapy. Blood 86:3961–3969

8. Suh C, Kim S, Kim SH et al (2004) Initiation of peripheral blood progenitor cell harvest based on peripheral blood hematopoietic progenitor cell counts enumerated by the Sysmex SE9000. Transfusion 44:1762–1768 9. Park KU, Kim SH, Suh C et al (2001) Correlation of hematopoietic progenitor cell count determined by the SE-automated hematology analyzer with CD34(+) cell count by flow cytometry in leukapheresis products. Am J Hematol 67:42–47 10. Dao MA, Arevelo J, Nolta JA (2003) Reversibility of CD34 expression on human hematopoietic stem cells that retain the capacity for secondary reconstitution. Blood 101:112–118 11. Hess DA, Wirthlin L, Craft TP et al (2006) Selection based on CD133 and high aldehyde dehydrogenase activity isolates long-term reconstituting human hematopoietic stem cells. Blood 107:2162–2169 12. Rosenbaum ER, Nakagawa M, Pesek G et al (2009) A 15 hour extended dosing-collection interval for Plerixafor is at least as effective as the standard 10 hour interval. Blood 114:2152 13. Product information. Leukine (sargramostim). Seattle, WA: Bayer Healthcare Pharmaceuticals, April 2008 14. Product information. Neupogen (filgrastim). Thousand Oaks, CA: Amgen Inc., 1991–1996. 15. Product information. Mozobil (plerixafor). Cambridge, MA: Genzyme Corporation, December 2008

Chapter 6 Monitoring Blood for CD34+ Cells to Determine Timing of Hematopoietic Progenitor Cells Apheresis M. Louette Vaughn and Edmund K. Waller Abstract Hematopoietic Progenitor Cell (HPC) Apheresis generally results in a mononuclear cell product that is highly enriched for hematopoietic stem and progenitor cells when performed on autologous patients in whom autologous stem cell transplant is planned who have been mobilized with cytotoxic chemotherapy and exogenous hematopoietic growth factors (cytokines) and possibly CXCR4 antagonists. Alternatively, patients scheduled for autologous transplants may be mobilized with cytokines only or a combination of cytokines and CXCR4 antagonists. Allogeneic Donors, either matched related donors (MRD) or matched unrelated donors (MUD), are typically mobilized with cytokines only. The HPC Apheresis product, enriched for hematopoietic progenitor cells collected from the patient/donor’s peripheral blood via an apheresis system, is used for restoring hematopoiesis in the patient/recipient who has received myeloablative therapy. Timing of the collection of an HPC Apheresis product from allogeneic donors is based on the schedule of the recipient’s myeloablative regime. However, the optimal timing of collection on HPC Apheresis product from a patient scheduled for an autologous stem cell transplant can be complex. Key words: Peripheral blood progenitor cells (PBPC), Cytokines, Granulocyte colony stimulating factor (G-CSF), CD34+ cells, Plerixarfor (Mozobil™), COBE® Spectra Apheresis System™, Sysmex XE-2100L® automated hematology analyzer

1. Introduction Granulocyte colony stimulating factor (G-CSF) mobilized peripheral blood stem cells (PBSC) have become the major source of hematopoietic stem cells for autologous transplant (1). The most commonly used method of stem cell mobilization involves daily or twice injections of G-CSF with or without chemotherapy. (2). In 2009 plerixafor (Mozobil™), an antagonist of the α-chemokine receptor CXCR4, was approved by Food and Drug Administration (FDA) for use in combination with G-CSF to mobilize PBPC in Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_6, © Springer Science+Business Media, LLC 2012

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lymphoma and myeloma patients anticipating autologous transplant (3). Several clinical trials have demonstrated that plerixafor (Mozobil™) can dramatically increase peripheral blood CD34+ cell counts, increasing the efficiency of stem cell collection in lymphoma and myeloma patients (4–6). Successful collection of the HPC graft by apheresis involves coordinating the administration and timing of the CXCR4 antagonist and (in the case of autologous transplant recipients) cytotoxic chemotherapy with the apheresis team and treating physician. Autologous donors respond differently to mobilization regimes based on diagnosis, age, and amount and type of prior treatment (chemotherapy and/or radiation) (7). 1.1. Chemotherapy and Cytokine Mobilization Initial Follow-Up

Determining the timing of HPC Apheresis after chemotherapy and G-CSF mobilization can be complex. Generally the Autologous donor is scheduled for outpatient lab tests, including a CBC to be drawn 7–10 days after the administration of cytotoxic chemotherapy that is used for mobilization in conjunction with once daily G-CSF injections. Follow-up Standard of Care Labs include Complete Blood Count (CBC), Automated Differential, Magnesium, and basic Biochemical Profile with a turnaround time of approximately 60 min. After the labs are resulted the Autologous donor is seen and evaluated by a mid-level Practitioner. At this point in time, it is not unusual for the Autologous donor’s absolute WBC count to be <1,000/μL. At this institution when the absolute WBC count exceeds 1,000/μL, a peripheral blood sample is sent for CD34+ cell determination with turnaround time of 2–3 h. Based on lab results the Autologous donor may also require transfusion or electrolyte replacement (see Note 1).

1.2. Transfer of Care After Chemotherapy and Cytokine Mobilization to Hemapheresis Center

After this initial follow-up outpatient visit in the BMT clinic, the donor will be instructed when to begin twice daily G-CSF injections and when to report to the Hemapheresis Center for labs and assessment for timing for initiation of HPC Apheresis. Generally this visit occurs on or about Day 12–14 after chemo mobilization. Standard of care labs—CBC, Automated Differential, Magnesium, and basic Biochemical Profile—are drawn with turnaround time of approximately 60 min. The Hemapheresis Nurse Clinician performs donor assessment to include vital signs and temperature and symptoms of chest pain, shortness of breath, lightheadedness, headache, and bleeding. Based on lab results and donor assessment, the donor may also require transfusion or electrolyte replacement. At this institution when the absolute WBC count exceeds 1,000/μL, a peripheral blood sample is sent for CD34+ cell determination with turnaround time usually 2–3 h (see Note 2).

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1.3. Timing of HPC Apheresis Based on CD34+ Cell Count, Hematocrit, and Platelet Count

When peripheral blood CD34+ cell count reaches 10–20 cells/μL, Hematocrit is ³27.0%, and Platelet Count is ³35,000, HPC Apheresis is initiated on the COBE ®Spectra Apheresis System™. Generally absolute WBC count is ³10,000–20,000/μL at this time.

1.4. Use of Plerixarfor (Mozobil™) After Chemotherapy and Cytokine Mobilization

If peripheral blood CD34+ cell count is £10 cells/μL and absolute WBC count is ³10,000–20,000/μL, the patient is considered to be a poor mobilizer, and plerixarfor (Mozobil™) may be administered 12–14 h prior to initiation of HPC Apheresis (the following morning) at a dose of 0.16–0.24 mg/kg (not to exceed 27–40 mg/ day) (8). Plerixafor (Mozobil™) injection is indicated for use in combination with granulocyte-colony stimulating factor (G-CSF) to mobilize hematopoietic stem cells to the bloodstream for collection and subsequent autologous transplantation in patients with non-Hodgkin’s lymphoma and multiple myeloma. HPC Apheresis is initiated the following morning 12–14 h after Plerixafor (Mozobil™) injection.

1.5. Autologous Donor Cytokine Mobilization Only

G-CSF is administered at 7.5 mcg/kg twice a day (BID) for 5 days and HPC Apheresis is initiated on Day #5 of G-CSF administration. On Day #4 of G-CSF administration, the donor is scheduled for lab draws and a peripheral blood sample is send for CD34+ cell determination. Turnaround time is usually 2–3 h. If peripheral blood CD34+ cell count is ³10–20 cells/μL, HPC Apheresis is initiated on the morning of Day #5 of G-CSF administration. If peripheral blood CD34+ cell count is £10 cells/μL, plerixarfor (Mozobil™) may be administered 12–14 h prior to initiation of HPC Apheresis at 0.16–0.24 mg/kg (not to exceed 27–40 mg/ day) on the evening of Day #4. HPC Apheresis is initiated on the morning of Day #5 of G-CSF administration.

1.6. Allogeneic Donors

Allogeneic Donors, either matched related donors (MRD) or matched unrelated donors (MUD), are mobilized with cytokines only. Allogeneic donors may respond differently to cytokine mobilization based on age. Generally at this institution MRD Allogeneic donors (age ³60 years) are mobilized and cells are collected and stored prior to recipient beginning chemotherapy conditioning regime. Generally if the allogeneic donors age is <60 years, cells are collected in the morning and given fresh to the recipient in the afternoon. G-CSF is administered at 5.0–7.5 mcg/kg twice a day (BID) for 5 days and HPC Apheresis is initiated on Day #5 of G-CSF administration.

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2. Materials 2.1. Autologous Donor Chemotherapy and Cytokine Mobilization

1. Vacutainer Adapter. 2. Vacutainer Holder. 3. Alcohol Swabs. 4. Lavender Top Vacutainer Tubes. 5. Green Top Vacutainer Tubes. 6. Thermometer. 7. Stethoscope. 8. Sphygmomanometer. 9. Sysmex XE2100L Automated Cell Analyzer. 10. Plerixafor (Mozobil™) injection, optional.

2.2. Allogeneic Cytokine Mobilization

1. Vacutainer Adapter. 2. Vacutainer Holder. 3. Alcohol Swabs. 4. Lavender Top Vacutainer Tubes. 5. Green Top Vacutainer Tubes. 6. Thermometer. 7. Stethoscope. 8. Sphygmomanometer. 9. Sysmex XE2100L Automated Cell Analyzer.

3. Methods 3.1. Sysmex XE-2100L™ Automated Cell Analyzer

This facility’s Hemapheresis Center has a Sysmex XE-2100L™ automated hematology analyzer to perform blood tests for 23 standard blood parameters and for four research blood parameters, which includes immature granulocyte%, immature granulocyte count, HPC% and HPC Count(#). The XE-2100L performs hematology analyses utilizing the RF/DC detection method, HydroDynamic Focusing (DC Detection), Flow Cytometry Method (using a semiconductor laser), and SLS-Hemoglobin Method. The RF/DC detection method is used to measure the total cell size and total cellular density to determine HPC% and HPC# (9). This facility has completed extensive correlation studies between HPC# and CD34+ cell results utilizing the same blood sample for both tests to determine the HPC# that correlates to 10–20 CD34+ cells/μL.

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When an Autologous Donor presents to the Hemapheresis Center for labs and assessment after either chemotherapy and G-CSF mobilization or G-CSF mobilization only, prior to sending the donor’s peripheral blood sample for CD34+ cell determination, the sample is analyzed on the Sysmex XE-2100L™. In 3 min the results of the HPC# is known so a determination can be made to initiate HPC Apheresis now or wait the 2–3 h for the CD34+ cell results. Use of the HPC# to determine timing of HPC Apheresis has greatly increased patient satisfaction and decreased wait times for donors.

4. Notes 1. When combined Magnesium Sulfate and KCL IV Infusions are given only 10 mEq/h of KCL can be administered. If Hematocrit <27.0%, transfuse with red blood cells (RBCs). Generally each unit of RBCs boosts the Hematocrit by 3.0%. Autologous donors are instructed to give their G-CSF injection approximately 12 h apart (at 6:00 a.m. and 6:00 p.m.) rotating the injection sites in the upper arm, thighs, or abdomen. For K+ <4.0: IV Infusion of KCL 40 mEq in 100 cm3 D5W IV over 4 h. For Mg2+ <1.5: IV Infusion of Magnesium Sulfate 4 g IV in 100 mL D5W over 2 h. For Platelet Count <20,000: transfuse with Platelet Apheresis products. 2. If Autologous donor’s temperature is ³100.5 °F, blood cultures are drawn for analysis. References 1. Moog R (2006) Mobilization and harvesting of peripheral blood stem cells. Curr Stem Cell Res Ther 1:189–201 2. Pusic I, DiPersio JF (2008) The use of growth factors in hematopoietic stem cell transplantation. Curr Pharm Des 14:1950–1961 3. Traynor K (2009) Plerixafor approved for autologous hematopoietic stem-cell transplantation. Am J Health Syst Pharm 66:112 4. Cashen A, Lopez S, Gao F et al (2008) A phase II study of plerixafor (AMD3100) plus G-CSF for autologous hematopoietic progenitor cell mobilization in patients with Hodgkin lymphoma. Biol Blood Marrow Transplant 14:1253–1261 5. DiPersio JF, Micallef IN, Stiff PJ et al (2009) Phase III prospective randomized double-blind placebo-controlled trial of plerixafor plus granulocyte colony-stimulating factor com-

pared with placebo plus granulocyte colonystimulating factor for autologous stem-cell mobilization and transplantation for patients with non-Hodgkin’s lymphoma. J Clin Oncol 27:4767–4773 6. DiPersio JF, Stadtmauer EA, Nademanee A et al (2009) Plerixafor and G-CSF versus placebo and G-CSF to mobilize hematopoietic stem cells for autologous stem cell transplantation in patients with multiple myeloma. Blood 113:5720–5726 7. McLeod BC, Price TH, Weinstein R (2003) Apheresis: principles and practice, 2nd edn. AABB Press, Bethesda, MA, pp 508–510 8. Mozobil™ (plerixafor injection) Prescribing Information. Genzyme Corporation 9. Operators Manual for SYSMEX XE-2100L. Sysmex America Corporation. Mundelein, Illinois, USA

Chapter 7 Hematopoietic Progenitor Cell Collection S. Darlene Marlow and Myra House Abstract Hematopoietic progenitor cells can be mobilized from the bone marrow microenvironment into the peripheral blood following treatment of patients with myeloid cytokines (GCSF, GMCSF, IL3), a CXCR4 antagonist (Plerixafor) and/or following a hematopoietic recovery from cytotoxic chemotherapy. The hematopoietic stem and progenitor cells are contained within the mononuclear cell fraction of peripheral blood and can be collected by apheresis in which the cellular constituents of the blood are separated on the basis of their buoyant density. Modern apheresis allows processing of five or more blood volumes (24 L or more) over a 4–5-h period to efficiently remove and separate more than 70 % of the CD34 positive cell progenitors present to blood. Management of a patient undergoing apheresis requires careful attention to venous access, calcium placement to counteract the effects of the citrate uses anticoagulant and hemodynamic monitoring. The principles of setting up the COBE spectra and its operation are reviewed. Management of common toxicities including hypocalcemia, allergic reactions, and vasovagal reactions are described in the next chapter. Key words: Cytotoxic chemotherapy, Hypocalcemia, Apheresis

1. Introduction The administration of myeloid cytokine or the CXCR antagonists plerixafor to donors causes leukocytosis and mobilization of CD34 positive cells from the bone marrow into the peripheral circulation. For grafting purposes, hematopoietic progenitor cells can be collected from the peripheral blood by apheresis. In this procedure, the leukocytes are separated from red cells and plasma following centrifugation in an extracorporeal system, usually maintained by continuous recirculation of blood through the

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apheresis device. Management of the apheresis procedure requires careful attention to flow rate of blood from the patient into the device to prevent hypotension, administration of anticoagulant into the extracorporeal loop to prevent thrombosis of the product within the device or its tubing, and monitoring of total extracorporeal blood volume and administered fluids to prevent volume overload. Programming the total volume of blood to be processed through apheresis, allows the collection of a target number of leukocytes that can be safely performed in an outpatient setting. Specific settings on the COBE Spectra apheresis machine allow collection of different leukocyte subsets with enrichment of hematopoietic progenitors using the mononuclear cell preprogrammed setting on the machine (1–4). Management of toxicities of the apheresis procedure includes reversal of hypocalcemia, hypotension, and allergic reactions. Once a patient has been mobilized with growth factor, and determined to be ready for a Hematopoietic Stem Cell Collection, the COBE Spectra apheresis instrument will be set up and primed using the steps outlined in this chapter.

2. Materials 1. Single-stage filler. 2. WBC disposable set. 3. Anticoagulent Citrate Dextrose Formula A (ACD-A). 4. 0.9 % NaCl for injection (1,000 mL). 5. Calcium Gluconate per physician orders. 6. 10–20 cc Syringes. 7. Sampling site coupler. 8. Pressure sensitive security seals. 9. COBE Spectra WBC Colorgram. 10. Central Venous Catheter supplies. 11. Paperwork (i.e., MD Orders, Apheresis Flow Sheet, Product Labeling Log, etc.). 12. 17–19 gauge needles, if using venous access. 13. Venipuncture site preparation supplies. 14. Preprocedure lab supplies. 15. IV Infusion set.

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3. Methods 3.1. Equipment and Disposable Tubing Setup and Instillation

1. Plug instrument into emergency power outlet. 2. Turn power switch ON. 3. Install the Standard single-stage filler (see Note 1). 4. Remove the disposable package cover and record lot number on appropriate paperwork. 5. Remove the inlet coil and hang on left side on the machine. 6. Place the access saline line over the top of the machine. 7. Remove the return coil and hang on the left side of the machine. 8. Place the return saline spike over the top of the machine. 9. Remove the bags and place them in the correct position on the IV pole. Refer to the Operators Manual for correct bag position. 10. Remove the return pump cartridge and snap it into position between the plasma and return pump. 11. Remove the access pump cartridge and snap it into position between the AC and inlet pumps. 12. Place the AC line over the top of the machine. 13. Press the CONTINUE key to load the tubing into the pump housings. Verify that all four pumps are loaded. 14. Place the lines in the collect/replace and plasma valves. 15. Place the return pressure sensor in the return pressure housing. 16. Place the RBC line into the RBC valve. Ensure that the line is completely inserted into the RBC detector. 17. Position the return and inlet air chambers in the air detectors with the air chamber filters below the air detector housings. 18. Place the waste lines into the waste valve assembly. 19. Place the line in the centrifuge pressure sensor using a “flossing” action. 20. Place the access pressure sensor in the access pressure sensor housing. 21. Position the return line in the return valve so that the line runs horizontally through the center of the valve.

3.2. Installing Channel in Centrifuge

1. Press the UNLOCK COVER key. 2. Open the centrifuge by sliding the centrifuge cover back and lowering the door.

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3. Gently extend the tubing leading to the centrifuge loop to full length to ensure that the tubing is not twisted. 4. Fold the centrifuge channel in half and load by inserting the channel through the loading port and pulling through from the top. 5. Position the channel in the correct orientation above the filler slots before placing the centrifuge collar into the collar holder. 6. Load the centrifuge collar into the centrifuge collar holder, closing the cover the over the collar. 7. Lower the filler latch into the locked position. 8. Press the channel into position. 9. Press the tubes into the appropriate slots in the filler. 10. Place the lower bearing in the lower bearing holder, making sure that the hard plastic bearing is securely seated in the holder. 11. Place the upper bearing in the upper bearing holder, making sure that the hard plastic bearing is securely seated in the holder. 12. Place the upper collar into the upper holder. 13. Place the multilumen tubing in the exit slot on the right side of the system using a “flossing” action. 14. Rotate the centrifuge clockwise several times to ensure that the tubing does not twist and stays in place. 15. Close the centrifuge cover. 3.3. Priming the Disposable Tubing Set

1. Check the needle luer connections to make sure that they are secure. 2. Press the “3” key to select WBC when prompted to select set type, selecting “1” for MNC. 3. Close the white pinch clamps on the access and return lines. Close the roller clamps on both saline lines. The spike leading to the AC container is orange. 4. Press the CONTINUE key. 5. Connect the AC line with the orange spike to the AC container and place in the AC level detector. 6. Connect the access and return saline lines to the saline container. Press the CONTINUE key. 7. Open the roller clamps on the two saline lines. Press the CONTINUE key to prime the disposable set.

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8. When prompted, prime the access and return connections by opening the white pinch clamps near the luer connections (see Note 3). 9. Close the access saline line, clamp the access line and press the CONTINUE key to test the AC ratio (see Note 2). 3.4. Alarm Tests

1. Press the CONTINUE key to clear the warning from the screen. 2. Press the YES key to perform the alarm tests. 3. Ensure that the access saline line is closed and the access and return lines are clamped. Press the CONTINUE key. The machine will test the access pressure sensor low alarm and the return chamber alarm. 4. At the prompt, open the roller clamp on the access saline line. Press the CONTINUE key. The machine will test the return pressure high alarm and the air in return chamber alarm. 5. At the prompt, verify that the return line valve is closed. Press the CONTINUE key to go on to the next test. 6. Press the UNLOCK COVER key and open the centrifuge door to perform the fluid leak detector alarm test. 7. Touch the fluid leak detector with your fingers to initiate the alarm. Verify if the machine indicates that fluid is detected in the centrifuge. 8. Close the cover and door. Press the CONTINUE key. 9. Close the roller clamp on the access saline. 10. Place the sampling site coupler into the collect bag, using aseptic technique.

3.5. Enter Patient Data

1. Enter the patient’s sex. Press ENTER key. 2. Enter the patient’s height. Press ENTER key. 3. Enter the patient’s weight. Press ENTER key. 4. Respond to the Total Blood Volume by pressing the YES key. 5. Inject ACD-A to coupler, using aseptic technique, typically at 10 % of collect volume.

3.6. Connecting the Patient

1. Take patients vital signs, record on apheresis flow sheet. 2. Access the patient. Collect labs as ordered. 3. Enter the hematocrit as whole number, press ENTER to accept. 4. Approve the WBC values by pressing the YES key. Pressing NO prompts the operator to change a value. Change values according to standard procedure or orders.

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3.7. Run Mode

1. Press the CONTINUE key to initiate the Run Mode. 2. When prompted, close the return saline roller clamp, press CLEAR. 3. The system will go into “Quick Start,” to set up the RBC/ Plasma Interface. 4. Once “Quick Start” is complete, monitor the collect line using the Colorgram, (Approximately 1–2 % Hct). Use the plasma pump to establish the interface making adjustments up or down in increments of 0.3 mL to get to the optimum interface color (see Notes 4 and 5). 5. Start Calcium Gluconate infusion, per standard procedure or physician order. 6. Monitor Vital Signs every 15–30 min and record on apheresis run sheet. 7. Continue collection per procedure. Typically, total blood processed is 10–24 L or 240–245 min.

3.8. Rinseback

1. Once collection values are reached, the machine will alarm, prompting the operator to CONTINUE or RINSEBACK. If RINSEBACK is selected, press “RINSEBACK”. 2. Clamp and seal collect bag, using aseptic technique. Remove product. 3. Clamp access line, open access saline roller clamp. Continue rinseback until instrument screen flashes “RINSEBACK COMPLETE”. 4. Close pinch clamp on return line and return saline roller clamp. 5. Record final values on apheresis flow sheet. 6. Disconnect patient using aseptic technique. 7. Press CONTINUE to unload pumps.

4. Notes 1. Use aseptic technique throughout the procedure. 2. Once fluid enters the tubing do not disturb any of the sensors. 3. If you are using a blood warmer, install according to manufacturer’s instruction. 4. Changing one value may affect other values. 5. Make changes gradually to the plasma pump flow rate; the separation will take a minute to respond.

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References 1. AABB. Technical manual, Current edn. AABB, Bethesda, Maryland 2. AABB. Apheresis principles and practice, Current edn. AABB Press, Bethesda, Maryland 3. Foundation for Accreditation of Cellular Therapy (FACT). Standards for cellular therapy

product collection, processing and administration, Current edn. JACIE International, Omaha, Nebraska 4. Gambro BCT, Inc. (2003) Cobe spectra cell therapy guide. Gambro BCT, Inc., Lakewood, Colorado

Chapter 8 Managing Apheresis Complications During the Hematopoietic Stem Cell Collection S. Darlene Marlow and Myra House Abstract Adverse reactions may occur during the Hematopoietic Progenitor Cells via Apheresis (HPC-A) collection, although they are rare. They can be associated with physical, psychological, or environmental factors. The most common complications are hypocalcemia related to citrate toxicity, allergic reactions, and vasovagal reactions. Recognizing and treating the symptom early can reduce the severity of most of these reactions. Procedural steps are outlined in this chapter for the proper management of apheresis complications. Key words: Apheresis, Hematopoietic Stem Cell, Adverse reactions

1. Introduction The collection of an engrafting dose of hematopoietic progenitor cells can be efficiently achieved by apheresis (1). In the apheresis maneuver small volumes of the donor’s blood are subjected to centrifugation in a sterile chamber with separation of leukocytes from plasma and red cells. Efficient apheresis includes continuous recirculation of the patient’s blood into the chamber with return of plasma and red cells and extraction of the leukocyte-enriched product (2). Toxicities of the apheresis procedure result from the need for an anticoagulant that must be added to the patient blood to prevent coagulation in the tubing or the centrifugation chamber of the apheresis unit, the potential for pyrogens that could be reintroduced to the patient, and the development of iatrogenic thrombocytopenia due to retention of platelets in the apheresis product (3). Apheresis complications can be safely managed by following the steps outlined in this chapter.

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_8, © Springer Science+Business Media, LLC 2012

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2. Materials 1. Blood Pressure monitoring system. 2. Oxygen Saturation monitoring system. 3. Thermometer. 4. Emergency Cart. 5. IV (intravenous) Infusion Pump. 6. Benadryl IV. 7. Tums. 8. Antinausea medication. 9. Calcium Gluconate 2 g IV (intravenous). 10. Normal Saline (1 L). 11. Warm Blankets. 12. IV Tubing. 13. Paper Bags.

3. Methods 3.1. Hypocalcemia/ Citrate Toxicity (see Note 1)

If the patient complains of tingling, numbness around the mouth and finger tips (perioral and circumoral paresthesias), a feeling of vibrations, epigastric discomfort, mild muscle cramps, or epigastric discomfort, proceed as follows: 1. Slow down the Inlet Flow rate. 2. Infuse IV calcium, (typically calcium gluconate 1–2 g/250 ml Normal Saline). 3. Give oral calcium (i.e., tums, milk, yogurt). 4. Massage legs. 5. Offer warm blankets. For severe muscle cramps, tetany, pressure or pain in chest, hypotension, nausea/vomiting, and generalized numbness, proceed as follows: 6. Pause the procedure. 7. Notify Medical Director. 8. Administer a saline bolus 9. Increase IV calcium flow rate.

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10. Administer an antinausea medication. 11. If the procedure is to be continued, adjust inlet flow to a slower rate. 3.2. Allergic Reaction (see Note 2)

If a patient experiences burning eyes, periorbital edema, hives, urticaria, wheezing, SOB, hypotension, and tachycardia, proceed as follows: 1. Discontinue the procedure. 2. Notify the Medical Director. 3. Administer Benadryl IVP (intravenous push). 4. If symptoms are not relieved immediately, give epinephrine (subcutaneous) and/or Solumedrol IV. 5. If the procedure is to be continued, reload the apheresis machine with a new kit that has been double primed.

3.3. Vasovagal/ Syncope Reaction (see Note 3)

If a patient experiences diaphoresis, pallor, dizziness, weakness, loss of consciousness, hypotension, nausea/vomiting, or hyperventilation, proceed as follows: 1. Pause the procedure. 2. Administer a 100 ml bolus of Normal Saline. 3. KVO (keep vein open) slow intravenous solution drip to maintain vein access. 4. Loosen constrictive clothing. 5. Elevate the feet. 6. Apply cold compresses to the forehead. 7. Have patient rebreathe into a paper bag for hyperventilating. 8. Calm and reassure the patient. 9. If the symptoms do not progress further, restart the collection and closely monitor the patient.

4. Notes 1. Extra precaution should be taken with patients in liver or renal failure because they can metabolize citrate poorly. 2. The most common cause of an allergic reaction during the HPC-A collection is the ethylene oxide gas used in the manufacturing of the apheresis kit. 3. Syncope during a HPC-A collection is most often caused by nervousness, the sight of blood, or it may happen for unexplained reasons.

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References 1. AABB. Apheresis principles and practice, Current edn. AABB Press, Bethesda, Maryland 2. Code of federal regulations, Parts 200–299 and 600–799, Current edn. U.S. Government Printing Office, Washington, D.C

3. Technical manual. Bethesda, Maryland

AABB,

Current

edn.

Chapter 9 Hematopoietic Progenitor Cell Apheresis Processing Eleanor S. Hamilton and Edmund K. Waller Abstract Modern apheresis generally results in a mononuclear cell product that is highly enriched for hematopoietic stem and progenitor cells when performed on patients who have mobilized with myeloid growth factors, CXCR4 antagonists or upon recovery from cytotoxic chemotherapy. The duration of apheresis during the days in which elevated numbers of CD34 positive cells are present in the blood depends upon the efficiency of the apheresis maneuver, the numbers of the CD34 target cell population in the blood, and the weight of the intended transplant recipient. The ability to freeze a hematopoietic progenitor cell (HPC) graft and subsequently thaw it while retaining viability of the hematopoietic stem and progenitor cell population has permitted routine collection of autologous hematopoietic progenitor cell grafts for treatment of patients with cancer using myeloablative doses of chemotherapy and radiation with stem cell support. Key words: Apheresis, Processing, Concentration, Cryopreservation, Thawing, Red cell reduction, Plasma reduction

1. Introduction Quality control procedures to assure that an adequate cell dose has been collected include sterility tests of the apheresis product, measurement of the numbers of CD34 positive cells in the apheresis product as well as their viability using dye exclusion methods. The HPC Apheresis (HPC-A) products contain not only hematopoietic stem and progenitor cells, but also variable amounts of red blood cells, lymphocytes, other white blood cells, and plasma. While these additional cell types are not problematic for autologous transplantation, they may cause transfusion related morbidities in allogeneic recipients. For example, ABO incompatible red cells or plasma in the apheresis product may cause posttransplant hemolysis (1). Additional tests to be performed on patients with

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_9, © Springer Science+Business Media, LLC 2012

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cancer undergoing autologous hematopoietic progenitor cell transplant include measurements of minimal residual disease, measurements of cancer cells that may be contaminated in the apheresis product using flow cytometry as well as detection of cytogenetic abnormalities. If the grafts are to be cryopreserved, they should be volume reduced to decrease the amount of cryoprotectant needed as well as the space needed for storage (2). Successful cryopreservation requires careful monitoring of the rate of freezing. Quality control procedures to address each of these important steps are addressed. To ensure consistent quality of cellular products processing, minimal standards must be set by each institution and monitored regularly (3). 1.1. Product Concentration/Plasma Reduction

A large volume apheresis typically produces a suspension of mononuclear cells enriched for hematopoietic stem and progenitor cells in a solution of plasma mixed with anticoagulant. The volume of the apheresis product following a 24 L apheresis procedure may exceed 300 mL and volume reduction is required to concentrate cells prior to cryopreservation and/or reinfusion of the cell product. The principles of cell concentration using low speed centrifugation as well as washing the cell product are reviewed using the Cobe 2991 cell washer (4). Procedures for calculating the net recovery of the CD34 positive cells following cell concentration or cell washing are noted as well as the minimal and optimal number of CD34 positive cells per kilogram sufficient to ensure reproducible hematopoietic engraftment following infusion into a patient treated with myeloablative doses of chemotherapy and radiation.

1.2. Product Cryopreservation

Successful cryopreservation of a viable hematopoietic cell progenitor cell population requires careful attention to the concentration of DMSO which is added to the hematopoietic stem or progenitor cells prior to phasing the concentration of other protein in the freezing solution, typically albumin, as well as control of the rate of freezing (5). Good clinical practice procedures for the stem cell lot require labeling of the product in the manner that labels are not dislodged during the freezing or thawing procedure.

1.3. Cell Thawing

Thawing is done quickly in a water bath solution with procedures to recover the thawed suspension of hematopoietic progenitor cells should the bag containing these cells break or rupture during the thawing procedure. The presence of DMSO with the hematopoietic progenitor cells leads to decreased viability of these cells over time (6). The timing of thawing and stem cell reinfusion to prevent loss of viable stem cells was reviewed.

1.4. Red Cell Reduction of HPC and Apheresis

Typically the apheresis product contains less than 10% of the red cell content of peripheral blood on a volume basis, with measured hematocrits usually in the range of 3–4%. In cases of allogeneic hematopoietic progenitor cell transplant in which the donor and

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recipient have ABO or Rh incompatibility, most of the donor red cells in the apheresis product must be removed prior to hematopoietic stem cell infusion in order to prevent a transfusion reaction. The procedure for removing the red blood cells from the apheresis product is described using differential sedimentation following lowspeed centrifugation as well as the maximal volume of incompatible red blood cells that can be typically reinfused into an ABO mismatched recipient without precipitating a severe transfusion reaction (7). Upon completion of processing the maximum volume of ABO incompatible red cells that can be safely infused is 30 mL.

2. Materials 2.1. Quality Assurance

1. Small sampling syringe 3–10 mL with luer lock end. 2. Sterile red luer lock cap. 3. Alcohol swabs, Isopropyl Alcohol, 70% v/v. 4. BacT/Alert FAN aerobic and anaerobic culture bottles.

2.2. Product Concentration/Plasma Reduction

1. Cobe 2991 Blood Cell Processing Set. 2. Small sampling syringe 3–10 mL with luer lock end. 3. Single Coupler plasma transfer set (Charter Med). 4. Sterile red luer lock cap. 5. 1,000 mL Plasmalyte A (Baxter Healthcare). 6. 60 mL syringes with luer lock end. 7. 250 mL of 5% human albumin with vented primary IV tubing (Baxter). 8. 3-way stopcock with male luer lock adapter (Mallinckrodt).

2.3. Product Cryopreservation

1. Controlled Systems).

rate

freezer

CBS-2100

(Custom

Biogenic

2. Origen 500 mL Freezing bags (Cryostore CS500, Origen Biomedical). 3. CryoServe—DMSO, 50 mL bottles (Bioniche Pharma). 4. 250 mL 5% albumin with vented luer lock IV tubing. 5. 1,000 mL Plasmalyte A (Baxter). 6. Single Coupler plasma transfer set (Charter Med). 7. Origen single couplers SA-10, DMSO resistant (Origen Biomedical). 8. 3-way stopcock with male luer lock adapter (Mallinckrodt). 9. 60 mL syringes with luer lock end. 10. 18 gauge blunt fill needles.

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11. Sampling site coupler (Fenwal). 12. Alcohol swabs, Isopropyl Alcohol, 70% v/v. 13. Hemostats. 14. 1 mL polypropylene low-temperature freezing vials. 2.4. Thawing

1. Three to four bottles (1,000 mL) of 0.9% Irrigation Sodium Chloride, sterile (Baxter). 2. 70% isopropyl alcohol. 3. Water bath. 4. Calibrated thermometer. 5. Portable insulated Nalgene storage bucket. 6. Cryogloves. 7. Clean towel.

2.5. Red Cell Reduction of HPC and Apheresis

1. Two 300 mL transfer bags with couplers (Fenwal). 2. Two sampling site couplers (Fenwal). 3. Single Coupler plasma transfer set (Charter Med). 4. Alcohol swabs, Isopropyl Alcohol, 70% v/v. 5. 18 gauge blunt fill needles. 6. Two 20 cc syringes. 7. Small sampling syringe 3–10 mL with luer lock end.

3. Methods 3.1. Quality Assurance

1. A nucleated cell count is performed on a hematology analyzer. A well-mixed 1 mL sample is aseptically taken from both the collection and the concentrated cell bag. White blood cell counts are obtained for nucleated cells (see Note 1). 2. Two sterility cultures are performed. A 1 mL sample is aseptically removed from the collection bag and the second sample is removed prior to cryopreservation. The top of the BacT/ Alert FAN bottle is cleaned with alcohol and the sample is injected into the bottle. The bottles are labeled with the patient information label and numbered “pre” and “post” respectively. All reports are expected to be negative (see Note 2). 3. A 1 mL sample from the processed transplant dose is removed for flow cytometry analysis. The samples should be are performed daily and reported on the day of each collection. The CD34% result is used by the physician to evaluate the need for further collections. The minimal CD34+ dose for transplant is

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2 × 106/kg, the optimal dose is >5 × 106/kg unless stated otherwise for a particular protocol. 4. A test for sample viability is performed with every product. The minimal accepted viability is >75% (see Note 3). 5. For an Autologous patient, a 1 mL sample of the product is sent to the laboratory which has been analyzing the patient’s blood and marrow samples for disease state related information. For example, lymphoma and myeloma patient samples may be monitored by the Flow Cytometry lab, whereas a patient with leukemia may be monitored by the Cytogenetics lab. 6. For all samples requiring cryopreservation, a copy of the freezing curve obtained from the step down freezer printer will be reviewed by the Medical Director (see Note 4). 3.2. Product Concentration/Plasma Reduction

1. The collection bag and if applicable, autologous plasma are properly identified and obtained from the Apheresis Department. 2. Tare the scale with empty Apheresis bag and weigh the product bag. Divide the weight in grams by 1.058 to determine the cell volume. 3. In a laminar flow hood, aseptically insert a single coupler into the cell product bag and obtain 1 mL samples for both total nucleated cell count and pre processing sterility. 4. Load the processing Cobe 2991 round bag into the centrifuge. Position the two white alignment blocks around the center stem of the cell-processing bag. Rotate the bowl cover counterclockwise until the locking plunger falls into place. Close the front and rear sliding cover. Lower the seal weight. Latch the centrifuge cover by rotating the knob counterclockwise (see Note 5). 5. The 2991 cell washer tubing is loaded by using the TUBE LOAD and the Valve selector knob. Load the tubing kit by placing the red line into Valve 1, the blue line on top of the COLLECT valve, the green line into Valve 2, the yellow line on top of Valve 3 and the purple line into the SOV valve and hang the waste bag on the hanger bars on the left side of the machine. 6. Place Press STOP/RESET and place valve selector on Valve 1. Place hemostats on the blue and yellow lines, attach a 1,000 mL bag of Plasmalyte A to the green line. 7. In the MANUAL MODE, set the cell washer settings as follows: Centrifuge speed 3,000; Super Out rate 450; Minimum agitate time 70; Super out volume 580. 8. Attach the cell product bag to the spike on the red line, press BLOOD IN and allow the entire cell product to drain into the processing bag. Press AIR OUT to evacuate air from the bag.

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9. After evacuating the air, a small amount of product will remain in the product bag. Using hemostats to control the flow of cells down the line, load the rest of the product by pressing BLOOD IN and stopping the cells before they exit the red line. Press STOP/RESET. 10. Rinse the product bag by placing a hemostat on the clear tubing above the rotating seal, remove the product bag and place on top the centrifuge cover. Press PRE-DILUTE and allow a small volume (50–60 mL) of the Plasmalyte A to flow into the empty cell product bag. Press STOP-RESET and mix the diluent with the remaining cells in the bag. 11. Remove the hemostats from the clear tubing above the rotating seal and hang the product bag on the left hanger bar. 12. Press BLOOD IN and allow the cell product to drain into the bag using a pair of hemostats to control the flow from the cell product line into the centrifuge bag and when the cells have reached the end of the red line, immediately hemostat the red line; (do not allow air to enter the processing bag). Press STOP/RESET. 13. To completely fill the processing bag, change the valve selector to Valve 2 and press TUBE LOAD. When the Plasmalyte A stops flowing, press STOP/RESET. 14. Press START/SPIN to start the centrifuge. Spin for 10 min. 15. Remove the 5% albumin from its container and vent the bottle with the IV tubing. Attach a 3-way stopcock to the other end of the tubing. Attach a sterile 60 mL syringe to a port of the stopcock. The albumin will be used to rinse the processing bag, dilute the product, and as part of the cryoprotectant solution. 16. At the end of 10 min press SUPER OUT to expel the supernatant portion from the processing bag. (With the SUPER OUT Volume set at 580, it is recommended to still observe the centrifuge contents when expelling the waste from the processing bag. This is to make sure that there is no cell loss during SUPER OUT). There will be approximately 40–50 mL of cells left in the bag. Press STOP/RESET. 17. Seal the clear line above the rotating seal. Remove the bag from the centrifuge and place into the laminar flow hood (see Note 6). 18. Remove the freezing bag from the packaging and visually inspect the container for tears and/or leaks. 19. Attach a single coupler to the processing bag, resuspend and remove the concentrated peripheral blood stem cell product with a sterile 60 mL syringe. Attach the 60 mL syringe to the labeled freezing bag and transfer the cells (see Note 7).

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Table 1 Chart for 10% DMSO freezing solution preparation (ingredients indicated in microliters) Plasmalyte A

30

60

90

120

150

180

240

5% Albumin

10

20

30

40

50

60

80

DMSO

10

20

30

40

50

60

80

Total

50

100

150

200

250

300

400

20. Remove 10–30 mL of 5% albumin with a syringe and attach it to the single coupler on the processing bag. Rinse the bag well and withdraw the rinsed cell suspension. Add the rinsed cell suspension to the concentrated product in the freezing bag. Repeat so that the centrifuge bag is rinsed twice. 21. Dilute the cell volume to at least 100 mL with the albumin. For transplant products, the cells are placed into a minimum of two freezing bags (see Note 8). 22. Mix the cells in the freezing bag and remove two 1-mL samples for both total nucleated cell count and Flow Cytometry analysis (Viability and CD34%). 23. Remove half of the cells from the freezing bag and place them into another appropriately labeled freezing bag. 3.3. Product Cryopreservation

1. In a laminar flow hood, prepare an empty freezing bag to be used as a blank to monitor the temperature during the freezing procedure: 2. With a 60 mL syringe, withdraw 50 mL of 5% albumin. Attach the 60 mL syringe to an empty freezing bag and transfer the albumin into the bag. Close the clamp on the tubing and remove the syringe (see Note 9). 3. Label the bag with the word “Blank,” heat-seal the tubing with the sealer, cut the tubing at the seal and remove. Place a sampling site coupler one of the ports. 4. Place the blank and the labeled freezing bags containing the concentrated HPC-A cells into a bucket filled with ice. 5. Determine the amount of freezing solution to be prepared. The amount of freezing solution needed is determined by the sum of the HPC-A cells and the volume of the blank. The freezing solution contains: 60% Plasmalyte A; 20% human albumin 5%; 20% DMSO (Table 1). 6. Aseptically attach a 600 mL transfer bag with a coupler into a bag of 1,000 mL Plasmalyte A and close tubing to bag with a hemostat.

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7. Hang the bag of Plasmalyte A on the right hanger of the Cobe 2991, place the transfer bag on the scale and press the tare button. 8. Remove the hemostat and allow the desired volume of Plasmalyte A into the transfer pack based upon the above chart. 9. When the desired volume is reached, quickly replace the hemostat. Heat-seal the line to the transfer bag (see Note 10). 10. Under the laminar flow hood, close the slide clamp of a single coupler from Gambro® and insert into the transfer bag. 11. Using a 60 mL syringe, aspirate the necessary amount of 5% albumin, connect the syringe to the coupler on the transfer bag, open the slide clamp, and transfer the albumin or plasma into the transfer bag. Close the slide clamp on the transfer bag. 12. Label the bag as “freezing solution.” And place the transfer bag with the Plasmalyte A and albumin into the bucket filled with ice and let the transfer bag cool at least 10 min. 13. While the freezing solution is cooling label the metal freezing holders using a water resistant lab marker. Label each holder with the required identifying information (see Note 11). 14. Place a clean towel on the counter next to the metal freezing canisters. 15. Turn on the control rate freezer to prep the chamber (see Note 12). 16. Using a 60 mL syringe, withdraw the appropriate amount of dimethyl sulfoxide (DMSO) and connect the syringe to the coupler on the freezing solution (Plasmalyte A and albumin). Keep the solution on ice. 17. While gently rocking the tan bucket, express 5–10 mL of DMSO into the transfer bag containing the freezing solution. Wait several minutes and repeat the process until all the DMSO has been added. 18. Allow the freezing solution to chill at least 5 min. 19. Remove the freezing solution from the ice and mix well and attach a sterile 60 mL syringe to the coupler on the freezing bag, release the clamp and withdraw 50 mL of freezing solution for the blank. Close the clamp and attach another 60 mL syringe to the bag. Keep the solution on ice. 20. Attach an 18 gauge blunt fill needle to the syringe containing 50 mL of freezing solution and slowly add 5–10 mL at a time of the freezing solution to the blank. Leave the syringe on the coupler. Mix by shaking the bucket of ice. Let the blank sit in the ice. 21. Remove the transfer bag containing the remaining freezing solution from the ice and mix well. Release the clamp and

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withdraw the amount of freezing solution needed for one bag of HPC-A. Example: If you have 60 mL of cells then you will need to add 60 mL of freezing solution. 22. Aseptically luer lock the syringe to the bag of HPC-A cells. Leave the bags in the ice. 23. Begin adding the freezing solution 5–10 mL at a time. Shake the bucket of ice to mix the solution with the HPC-A cells. Let cool. Leave the syringe attached to the tubing. The solution should be added a small amount at a time over a period of 10 min. 24. Repeat steps 11–13 until all the bags of cells are attached to syringes of freezing solution. You will need a separate syringe for each freezing bag. 25. At the point when almost all the freezing solution is added to the blank, begin the freezing program. The Initial cooling stage takes only about a minute to complete. The Idle light will come on and the program will not continue until CONTINUE is pressed. 26. With the syringe still attached to the bag, remove the bag and draw off the excess air that is in the blank. 27. Dry off the water from the outside of the bag with the clean towel and put the bag into the special metal holder for the blank. 28. Carefully insert the probe through the sampling site coupler and into the blank. Use care not to puncture the bag. Make sure that the probe is inserted into the liquid. 29. Place the blank in the middle slot of the freezer rack and close the door. The freezer will continue to cool. 30. Remove the first bag of cells. With the syringe still attached to the tubing of the bag, begin to express the air. At this time remove a 1 mL sample for sterility. Once the samples have been removed heat-seal the tubing. 31. Dry the bag with the clean towel, cut and remove the tubing at the seal. Carefully place each bag into the corresponding labeled canister and place the canisters and vial into the control rate freezer. 32. The chamber will still be in a prep stage, press CONTINUE and the freezing process will begin. There are five steps in this program (Table 2). 33. When the freezing is complete, an alarm will sound. Click “Stop” to end Click EXIT. 34. Fill a cooler with approximately 2–3 of liquid nitrogen. Remove the metal canisters and vial from the freezer. Place them into the cooler of liquid nitrogen.

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Table 2 Control rate freezing program

Cooling rate

Target temperature

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35. Transport them to the liquid nitrogen freezers and put them in the liquid nitrogen storage freezer location. 36. All cellular products containing a biohazard label should be stored in the top slots of the freezer (vapor phase only) to prevent possible cross contamination with other products (see Note 13). 3.4. Thawing

1. Schedule the time to thaw the cells with the patient’s nurse. 2. Approximately 30 min before thawing, clean the inside of the water bath and thermometer with alcohol. Fill the bath with sterile saline (Use approximately three to four bottles of the saline). This allows enough time for the temperature to reach 37°C. 3. Fill the cooler with liquid nitrogen. This cooler will be used to transport the HPC-A canisters to the thawing location. 4. Wearing cryogloves and protective eyewear, slowly lift the freezer rack, holding the canisters of cell products to be infused, from the freezer. 5. Place the canisters into the cooler and transport the canisters, infusion sheet with infusion information, and cryogloves to the thawing room. 6. Utilizing the infusion form, the nurse and staff member will then identify the information on the canisters Together verify the patient name, product identification number, date of birth, and the cell type. 7. When the nurse is ready to infuse, remove the first bag of cells from the freezing canister, verify the patient information on the freezing label and immerse the bag into the saline bath, keeping the ports above the water line. Allow the cells to thaw with minimal to no manipulation. It should only take 2–3 min (see Notes 14 and 15).

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8. Once the cell product is thawed, remove the bag from the bath and take the bag to the patient’s room. 9. Prior to infusion of the cells, the nurse and staff member will confirm the name and hospital number with the patient’s armband. Each bag that is brought for infusion will be verified by this procedure. 10. Repeat step 9 for each remaining bag to be thawed. 11. Once the cells are thawed, they are to be infused within 30 min. 12. When the infusion is completed, empty the saline from the bath and clean the bath and thermometer with alcohol. 3.5. Red Cell Reduction of HPC, Apheresis

1. The collection bag and if applicable, autologous plasma are properly identified and obtained from the Apheresis Department. 2. Tare the scale with empty Apheresis bag and weigh the product bag. Divide the weight in grams by 1.058 to determine the cell volume. 3. In a laminar flow hood, aseptically insert a single coupler into the cell product bag and obtain 1 mL samples for both total nucleated cell count and pre processing sterility. 4. Calculate the volume of red cells in the product(s) prior to processing. Multiply the volume in milliliters times the hematocrit %. An acceptable volume of ABO incompatible red cells needs to be determined by each facility to determine if red cell reduction is necessary (see Note 16). 5. Under the laminar flow hood, place relatively equal amounts of apheresis product into two 300 mL transfer bags. Note: this will be very subjective, to maintain the integrity and sterility of the system. 6. Place each bag in a centrifuge bag with the ports facing downward. 7. Weigh and balance each bag in the centrifuge cylinders using a trip balance scale, adding balancing disks as needed. 8. Centrifuge at 200 × g for 10 min at 22°C. 9. Carefully remove bags from centrifuge and allow to hang from the hooks in the laminar flow hood for 20 min (see Note 17). 10. Carefully place a sample site coupler in the bag. 11. Using aseptic technique, carefully remove red cells from the bottom of the bag with a 20 cc syringe, until the buffy coat layer is disturbed. Do not remove the buffy coat. 12. Gently mix bag and remove 1 mL sample for white blood cells and hematocrit. Subtract the volume of the red cells removed

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from the total volume of the product and perform necessary calculations as in step 4 (see Note 18). 13. Remove samples for flow cytometry analysis and sterility and label for infusion to the recipient.

4. Notes 1. Commonly white blood cell counts are higher than the linearity of most hematology analyzers. Perform a cont on the neat sample and then make a 1:10 dilution with the analyzers diluent and repeat the count. Compare the red blood cell and hemoglobin numbers to verify the dilution is accurate. Multiply the diluted result by 10 to determine the nucleated cell count × 106/mL. 2. Other microbiological testing can be performed besides the BacT/Alert system as it is cost prohibitive for small laboratories. Whichever method is used must detect common contaminants, both bacterial and fungal and must be validated for sensitivity and specificity. 3. Viability can be performed in conjunction with flow cytometry for CD34+ enumeration. Viability by 7-Amino-actinomycin D (7-AAD) or Propridium Iodide (PI) is more accurate than light microscopy techniques such as trypan blue viability (8). 4. Freeze a 0.5–1.0 mL sample vial of each product that is frozen. In the event of mechanical failure of the controlled rate freezer, the cells may be thawed for viability to determine if the cells sustained damage. 5. Once a day, prior to loading the Cobe 2991, prime the machine and check that the hydraulic fluid line has no bubbles. To do this press START/SPIN and let run for 15 s before pressing SUPER OUT. The excess pressure light should come on within 15 s. 6. Always heat-seal products two times and cut the seal farthest away from the product. This prevents the product from potential contamination should a single seal fail. 7. Always use a new 60 cc syringe and only use it once to help prevent contamination. 8. The maximum amount of nucleated cells that should be put into a single freezing bag is 400 × 108 total cells. 9. If 5% human albumin is not available then dilute 50 mL of 25% human albumin with 200 mL of normal saline in a transfer bag to make a 5% solution.

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10. Alternatively you can insert a single coupler into the bag of Plasmalyte A and measure the appropriate amount using 60 cc syringes. 11. The metal freezing holders may be reused. Clean permanent marker off outside of holder with Isopropyl alcohol. 12. Note: Make sure there is enough liquid nitrogen in the tank and the valve is open prior to starting the procedure. DMSO is added last and only just before freezing. DMSO releases heat and requires to be completely chilled before adding to the cells. 13. If a controlled rate freezer is unavailable then the cells may be placed directly from ice buckets into a −80°C freezer for an hour and then placed into long term liquid nitrogen storage. 14. If a bag containing product is found to have a leak, seal off leak with a hemostat and draw product out through a syringe using a sample site coupler. 15. If the cells clump after thawing then 10% Anticoagulant Citrate Dextrose Solution A (ACD-A) can be added to the bag to dissolve the clumps. 16. Acceptable volume of incompatible red cells is £30 mL in a hematopoietic progenitor cell apheresis product postprocessing. 17. If a good delineation is not received from centrifugation, the product can hang for up to 60 min. 18. If the volume of red cells does not fall in the acceptable range, then the laboratory medical director will be notified for further instructions. References 1. Lasky LC (1995) Marrow and stem cell processing for transplantation. American Association of Blood Banks, Bethesda, MD 2. Hillyer C (2009) Transfusion medicine and hemostasis. Elsevier, Burlington, MA 3. American Association of Blood Banks (2008) Standards for cellular therapy product services. American Association of Blood Banks, Bethesda, MD 4. Caridian BCT (2008) COBE 2991 Cell Processor Operators Manual. Caridian BCT, Inc., Lakewood, CO

5. Areman E (1992) Bone marrow and stem cell processing: a manual of current techniques. F.A. Davis Co., Philadelphia, PA 6. Sacher RA (1992) Marrow and transplantation: practical and technical aspects of stem cell reconstitution. American Association of Blood Banks, Bethesda, MD 7. Buckhalter R (2003) Inverted spin method for removing RBC’s from buffy coat products. Cytotherapy 5:553–557 8. Snyder E (2004) Cellular therapy: a Physician’s handbook. American Association of Blood Banks, Bethesda, MD

Chapter 10 Toxicities of Mobilized Stem Cell Infusion Jonathan L. Kaufman Abstract Infusion of bone marrow or cytokine mobilized stem cells can be associated with severe and potentially life-threatening toxicities. The etiology of the toxicities can be related to volume of infusion, dimethylsulfoxide (DMSO) or bacterial or viral infection of the infused cell products. Serious toxicities including hypotension and hypertension, cardiac arrhythmias, and respiratory failure require the prompt attention by a trained physician. Non-life-threatening toxicities are common and include fever, chills, nausea and vomiting. Fever may be caused by an infection or cytokines released from leukocytes in the hematopoietic cell product. Other possible adverse events can be attributed to the reagents used in the processing procedure. Monitoring and management of toxicities and the roles of nurses, technicians, and physicians are outlined below. Key words: Peripheral blood stem cells (PBSC), Dimethylsulfoxide, Toxicities

1. Introduction Hematopoietic cell transplantation involves an infusion of hundreds of millions to billions of leukocytes into a transplant recipient who has been conditioned with chemotherapy and radiation. The hematopoietic cell graft may be collected from the patients themselves prior to conditioning and cryopreserved with DMSO, or maybe an allogeneic product from a volunteer donor that is administered fresh or following cryopreservation and thawing. Infusion of a large number of leukocytes that may be obtained from a donor who may be ABO incompatible with the recipient creates the potential for a variety of infusion reactions including classical hemolytic transfusion reactions as well as fever or hypotension related to cytokine in the infused product. The most important principle to reduce and avoid toxicities associated with

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_10, © Springer Science+Business Media, LLC 2012

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hematopoietic cell infusion is to ensure that the proper product is administered by careful verification of the donor and recipients identifying information. The infusion of a graft product that has been cryopreserved with DMSO and subsequently thawed commonly causes toxicity secondary to the infusion of relatively large volumes of DMSO, which can result from DMSO-induced histamine release or direct cardiovascular effects. Other possible adverse events can be attributed to the reagents used in the processing procedure or bacterial contamination. Proper management of the toxicities associated with stem cell infusion requires careful planning, informing the patient, and having on hand monitoring equipment and drugs (1).

2. Materials 1. 500 ml normal saline intravenous solution. 2. Y-type blood set with large standard blood filter. 3. Nonsterile gloves. 4. Oxygen flow meter with nasal prongs. 5. Suction canister, gauge, tubing, and Yankauer catheter. 6. Pulse oximeter. 7. Automatic blood pressure cuff. 8. Emergency cart with emergency medications and respiratory equipment. 9. Heparin flush. 10. Medications properly labeled at bedside: Benadryl (IV); Epinephrine 1:1,000, 1 mg; Hydrocortisone 500 mg (IV); Atropine 1 mg (IV). 11. Hard candy (optional).

3. Methods 3.1. General Instructions

1. The patient must have a multilumen central venous catheter in place prior to beginning the marrow or PBSC infusion. 2. A registered nurse is permitted to perform this procedure; however, a physician must be present in the patient care area during the infusion.

3.2. Patient Preparation

1. Explain the procedure to the patient.

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2. Review potential side effects of the infusion: garlic taste in mouth and odor for 24–36 h, flushing, urge to cough, chest tightness, nausea, and darkened urine. 3. Ask patient to void prior to the procedure. 4. Offer patient hard candy to suck on during infusion. 3.3. Preparation for Marrow/PBSC Infusion

1. Coordinate infusion time with Cellular Therapy Lab staff and inpatient Bone Marrow Transplant attending physician. (see Note 1). 2. Review history and physical on medical record. 3. Assess morning weight prior to infusion. 4. Perform brief physical exam including (1) auscultate heart for irregular heart rate or extra sounds, (2) auscultate lungs for wheezing, crackles, and diminished breath sounds, (3) assess respiratory rate and skin color; (4) assess mental status and anxiety level. 5. Attach finger probe for pulse oximetry and apply automatic blood pressure cuff. 6. Measure and record baseline vital signs, and pulse oximeter oxygen saturation (SpO2). 7. Obtain physician order to premedicate patient 15 min prior to infusion. Usual medications include: Benadryl 50 mg (IV push), Tylenol 650 mg (PO), and Lorazepam 1 mg (IV).

3.4. Infusion Procedure

1. Notify attending physician that the infusion is about to begin. (see Note 2). 2. If infusion is 1 h or less: Assess vital signs and SpO2 at 15 min × 4 and evaluate SpO2 up to 1 h after infusion is completed. 3. If infusion is greater than 1 h: Assess vital signs and SpO2 at 15 min × 4, at 30 min × 2, at 1 h × 4, and SpO2 up to 1 h after infusion is completed. 4. Verify with the Cellular Therapy Lab technologist that the following information is identical on each marrow/stem cell container and the patient’s armband: (1) Name; (2) Medical record number; (3) Date of Birth (verify verbally with patient); (4) Social security number (verify verbally with patient). 5. Sign the Stem Cell Infusion Form (RN and Stem Cell Processing technologist). 6. Attach a Y-type standard blood set to a bag of normal saline and prime the line with the saline solution. Infuse normal saline through the largest lumen of the central venous catheter at a KVO rate. 7. Don a pair of nonsterile exam gloves.

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8. Verify the patient’s name and medical record number on the thawed cell container and patient’s armband at the patient’s bed/chair side with the Cellular Therapy Lab technologist. Document verification on the patient’s medical record. 9. Attach the thawed cell container to the available coupler on the filter set. Open the roller clamp to allow the cells to enter the filter. 10. Adjust the rate of infusion to 10–20 ml/min with the roller clamp located below the filter. 11. When the cell container has been emptied of all contents, open the normal saline clamp and allow normal saline to completely flush the line to ensure that all cells are administered. Return the rate of the normal saline to KVO while awaiting the next cell container. 12. When the next thawed cell container arrives, remove the empty cell container from the coupler, and attach the next cell container. 13. After the last cell container has infused, thoroughly flush the line with normal saline and heparanize the catheter lumen. 14. Instruct patient to immediately report any hematuria, weakness, dizziness, dyspnea, or chills. 15. Per physician order, if hematuria is present post infusion, begin normal saline I.V. hydration. 3.5. Adverse Reaction Procedure

1. Adverse reactions are to be reported immediately to the transplant attending physician and the medical director of the Cellular Therapy Lab. 2. If the reaction occurs after the infusion is complete, the hematopoietic progenitor cell laboratory is notified. 3. The Stem Cell infusion and Adverse Reaction forms will be submitted to the medical director for review. 4. A copy of each form will be placed in the patient’s hospital and clinic chart and processing notebook. A copy of the documentation will be given to the laboratory manager for supervisory review. 5. If death results from the infusion, the director of compliance, center for biological evaluation and research shall be notified by telephone as soon as possible. A written report of the investigation shall be submitted to the director, office of compliance and center for biologics evaluation and research, within 7 days after the fatality. 6. If contamination is suspected to have caused the reaction, cultures will be submitted from the stem cell product and a blood culture will be obtained from the patient.

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The following information is included on the Infusion Record, a copy of which is placed on the patient’s chart: 1. Source of stem cells (marrow, peripheral blood, or umbilical cord). 2. Type of vascular access device used for administration. 3. Number of cell containers thawed. 4. Yield (cell count) (obtain this number from Stem Cell Processing staff). 5. Total volume infused. 6. Patient response to the infusion, including adverse reactions, any interventions required and outcomes.

4. Notes 1. Prior to Marrow/PBSC Infusion: Notify attending physician if patient has a history of asthma, COPD, CHF, renal insufficiency, electrolyte imbalance or hypercoagulable state (pulmonary embolus or deep vein thrombosis). Notify attending physician if morning weight is greater than dry weight. Document physical examination findings in the medical record. Notify attending physician of any abnormal findings prior to the infusion. To minimize infusion-related complications, provide emotional support to patient and significant other(s). 2. Infusion Procedure: Notify attending physician for heart rate <50 beats/min, systolic BP <90 mmHg, SpO2 <90 %, or if patient appears clinically unstable at any time. Decrease the infusion rate or stop briefly if the patient experiences flushing, tightness in the chest, nausea, or vomiting. Notify the physician if SpO2 falls below 90 %, or if the patient experiences severe tightness of the chest, wheezing, or hypotension. The number of cell containers will vary according to yield and patient body weight. The typical number of cell containers will be three to four. Reference 1. Zambelli A, Poggi G, Da Prada G, Pedrazzoli P, Cuomo A, Miotti D, Perotti C, Preti P, Robustelli della Cuna G (1998) Clinical toxicity

of cryopreserved circulating progenitor cells infusion. Anticancer Res 18:4705–4708

Chapter 11 Mobilization of Hematopoietic Stem Cells by Depleting Bone Marrow Macrophages Valérie Barbier, Ingrid G. Winkler, and Jean-Pierre Lévesque Abstract An important factor contributing to hematopoietic stem cell (HSC) mobilization is the ability of mobilizing cytokines and chemotherapy to disturb the cellular components of HSC niches, particularly osteoblasts and their progenitors, and to inhibit the production of HSC supportive cytokines and chemokines. Although the mechanisms by which niche cells are inhibited by mobilizing treatments is still incompletely understood, it has recently emerged that bone marrow macrophages play a critical role in maintaining osteoblasts, bone formation, and the expression of CXCL12, KIT ligand, and angiopoietin-1 necessary to HSC maintenance. In this chapter, we describe how to mobilize HSC into the blood in mice by depleting macrophages with clodronate-loaded liposomes and compare this mode of mobilization to mobilization induced by granulocyte colony-stimulating factor and cyclophosphamide. Detailed methods to analyze mobilization of phenotypic and functional reconstituting HSC are described with examples. Key words: Hematopoietic stem cells, Mobilization, Bone marrow, Macrophages, Stem cell niche, Depletion of macrophages, Clodronate, Liposomes

1. Introduction Until recently, it has been assumed that the mobilization of hematopoietic stem and progenitor cells (HSPC) in response to granulocyte colony-stimulating factor (G-CSF), cyclophosphamide, or the chemokines CXCL8 (interleukin-8) and CXCL2 (GROβ) was in part mediated by bone marrow granulocytes. This conclusion was drawn from the following observations: (1) Mice deficient for the G-CSF receptor (Csf3r) gene are neutropenic and do not mobilize in response to G-CSF, cyclophosphamide, or CXCL8 (interleukin-8) (1, 2); (2) mice pretreated with the

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_11, © Springer Science+Business Media, LLC 2012

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anti-Gr1 monoclonal antibody RB6-8C5 become neutropenic and mobilize poorly in response to G-CSF or CXCL2 (3); and (3) neutrophil proteases such as neutrophil elastase, cathepsin G, and MMP-9 accumulate in human and mouse bone marrow mobilized with G-CSF or cyclophosphamide (4, 5) with concomitant proteolytic cleavage of proteins essential to retain HSPC within the bone marrow such as vascular cell adhesion molecule (VCAM)-1 and CXCL12 (5–7). However, HSPC mobilization and downregulation of CXCL12 expression in the bone marrow can still be observed in mice deficient for neutrophil elastase, cathepsin-G and MMP-9 treated with G-CSF (8) suggesting alternative mechanisms are involved. Importantly the Gr-1 antibody RB6-8C5 typically used to deplete mouse granulocytes recognizes both Ly6-G and Ly6-C cell surface antigens, the latter being highly expressed by monocytes and macrophages. Thus, in vivo treatment with antiGr1 antibody is likely to deplete several populations of macrophages in addition to granulocytes. As a result, the exact contribution of granulocytes and their proteases remains controversial. Clearly, the downregulation of CXCL12, one of the main mechanisms by which HSPC are mobilized, involves protease-independent mechanisms such as the depletion or suppression of osteoblast lineage cells which produce CXCL12 in the bone marrow (9–13) in addition to CXCL12 inactivation by proteolytic cleavage. We have recently demonstrated in mice that bone marrow phagocytic macrophages play a critical role in maintaining osteoblasts, bone formation, and the expression of CXCL12, KIT ligand, and angiopoietin-1 necessary to HSC maintenance (14). Indeed, depletion of bone marrow macrophages in vivo either genetically using macrophage Fas-induced apoptosis (Mafia) transgenic mice, or pharmacologically using clodronate-loaded liposomes (1) depletes osteoblasts and inhibits bone formation, (2) reduces expression of CXCL12 and HSC-supportive cytokines at the endosteum, and (3) mobilizes HSC into the peripheral blood. All these events occur independently of G-CSF signaling and neutrophil proteases (14). In this chapter, we describe how to mobilize HSC in mice by depleting macrophages with clodronate-loaded liposomes and compare this mode of mobilization to mobilization induced by G-CSF and cyclophosphamide. Clodronate (dichloromethyl bisphosphonate) is a highly polar and negatively charged molecule that cannot cross lipid bilayers and cell plasma membranes. In aqueous solution, clodronate has very little toxicity. Once encapsulated into liposomes, it very efficiently kills all phagocytes including macrophages, activated dendritic cells and to a lower extent osteoclasts (15). Unlike G-CSF and other cytokines, liposomes are active across all mammalian species and could therefore be useful to further elucidate the role of macrophages in maintenance

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of HSC niches and osteoblasts in the bone marrow in both mice and primates, as well as their role in initiating mobilization of HSC and other progenitor populations (e.g., endothelial progenitor cells and mesenchymal stem cells). This chapter also describes common assays to detect mobilization of HSPC in blood such flow cytometry, colony-forming assays, and long-term competitive repopulation assays.

2. Materials 2.1. Animal Treatment

1. For granulocyte colony-stimulating factor (G-CSF)-induced HSC mobilization use Clinical grade recombinant human (rhu) G-CSF (Filgastim Amgen, Thousand Oaks, CA). The stock rhuG-CSF solution can be stored long-term at 4°C and is diluted in saline to 25 μg/mL on the day of use. 2. Clinical grade sterile injectable saline to dilute rhuG-CSF (see Note 1). 3. Clodronate-loaded liposomes and control phosphate buffered saline (PBS)-loaded liposomes. A detailed method to prepare these liposomes has been described earlier in this series (16) (see Note 2). 4. Clinical grade cyclophosphamide: Freshly prepared 20 mg/ mL stock in clinical grade saline can either be obtained from hospital pharmacy or purchased commercially. Store at 4°C (see Note 3). 5. Isoflurane inhalation anesthetic. 6. Insulin syringe with attached needle 27G1/2 for intravenous retroorbital injections (liposomes). 7. 1 mL tuberculin syringe mounted with 26G1/2 needles for subcutaneous (G-CSF) or intraperitoneal cyclophosphamide injections.

2.2. Tissue Harvest

1. Dulbecco’s Phosphate Buffered Saline (DPBS) without calcium or magnesium supplemented with 2% heat-inactivated newborn calf serum (NCS), sterile filtered. 2. 1 mL tuberculin syringe mounted with 23 G needles. 3. 5 mL polypropylene tubes. 4. 1.5 mL Eppendorf tubes. 5. Heparin for blood collection and for in vivo injection to inactivate thrombin clots. Dilute DBL heparin sodium from porcine mucous (Hospira, Lake Forest, IL) to 1 unit/μL in sterile injectable saline. Store 4°C.

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2.3. Cell Processing

1. Automated Hematology Analyzer KX-21N (Sysmex, Kobe, Japan) or manual counting of cells following (>1:20) dilution in white cell counting fluid using a brightfield Neubauer microscope counting chamber. 2. Tissue culture petri dishes 35 × 10 mm. 3. Microscope slides rough with frosted-glass ends. 4. 40 μm nylon cell strainers. 5. 10× red cell lysis buffer. 1.5 M NH4Cl, 100 mM NaHCO3, 10 mM EDTA pH 7.4. Sterile stock can be kept in fridge for many months. On the day of the experiment, dilute 1 part of 10× red cell lysis buffer with 9 parts of sterile water to make 1× red cell lysis buffer. 6. Refrigerated centrifuge to rotate 1–50 mL tubes and microplates at 370 × g.

2.4. Phenotypic Stains of Hematopoietic Stem Cells and Macrophages

1. 5 mL polypropylene tubes, Greiner cat#115262. 2. Stain tubes: ideally 1.2 mL microtibertube or other polypropylene staining tubes. 3. Fc Block: Purified rat anti-mouse Fcγ receptor II/III clone 2.4 G2 (block at 5 μg/mL) or batch-tested culture supernatant from Fc block hybridoma (block at 1:2 dilution). 4. MACS buffer: DPBS + 0.5% bovine serum albumin + 2 mM EDTA. 5. DPBS + 2% NCS. 6. Conjugated monoclonal antibodies specific for mouse antigens: CD3ε-biotin clone 145-2C11 (BD), 0.5 mg/mL. CD5-biotin clone 53-7.3 (BD), 0.5 mg/mL. CD45R (B220)-biotin clone RA3-6B2 (BD), 0.5 mg/mL. Gr1-biotin clone RB6-8C5 (BD), 0.5 mg/mL. F4/80-biotin clone BM8 (eBioscience), 0.2 mg/mL. CD41-biotin clone MWreg30 (eBioscience) cat#13-0411-85, 0.5 mg/mL. Ter119-biotin clone Ter119 (BD) 0.5 mg/mL. Sca-1-PECY7 clone D7 (BD), 0.2 mg/mL. Kit (CD117)-APC clone 2B8 (Biolegend) 0.2 mg/mL. CD48-FITC clone HM 48-1 (BD) 0.5 mg/mL. CD150-PE clone TC15-12 F 12.2 (Biolegend) 0.2 mg/mL. CD11b-V450 clone M1/70 (BD) 0.2 mg/mL. CD11b-PECY7 clone M1/70 (BD) 0.2 mg/mL. LY6G-PE clone 1A8 (BD) 0.2 mg/mL.

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F4/80-Alexa647 clone CI:A3-1 (Biolegend) 0.5 mg/mL. Streptavidin-Alexa700 (Invitrogen) 0.5 mg/mL. 7. 7-amino actinomycin D (7-AAD) for cell viability stain, 1 mg/ mL stock stored at 4°C. 8. 8–9 color flow cytometer equipped with 406 nm violet laser (with 450/50 filter for V450), 488 nm blue laser (with 530/40, 575/25, 710/30, and 787/43 filters for FITC, PE, 7-AAD, PECY7, respectively), and 643 nm red laser (with 665/20 and 750LP filters for APC/Alexa647 and APCCY7/ Alexa700 respectively). 9. FloJo software (Tree Star, Ashland, OR) or other for analysis of results. 2.5. Colony Assays

1. Methylcellulose Methocel MC (Fluka) or A4M powder Premium grade 4,000 cP (Dow Chemicals). 2. Iscove’s modified Dulbecco’s medium (IMDM) in liquid with Hepes, NaHCO3. 3. L-Glutamine (100×) stock at 200 mM. 4. Human insulin (Atrapid, NovoNordisk). 5. Recombinant mouse interleukin (IL)-3, mouse or human IL-6, mouse or rat KIT ligand (Peprotec, Israel), or conditioned medium of cell lines stably transfected with corresponding cDNA. 6. Penicillin 10,000 units/mL streptomycin 10,000 μg/mL (100×) stock. 7. Fluconazole solution for intravenous infusion 200 mg/100 mL (200×) stock (Diflucan, Pfizer). 8. Tissue culture petri dishes 35 × 10 mm. 9. 5 mL syringe. 10. Sterile cannulas (Unomedical, Mona Vale, NSW, Australia). 11. Humid incubator 37°C supplemented with 5%CO2. 12. Dissection microscope.

2.6. Transplantations

1. Mouse irradiator Gammacell 40E, Twin caesium 137 source (MDS Nordion, Canada). 2. Clinical grade injectable quality saline, sodium chloride injection BP, 0.9%, 50 × 10 mL sterile ampoules (Pfizer, West Ryde NSW, Australia). 3. 50 mL polypropylene tubes. 4. 14 mL round bottom polypropylene tubes with cap. 5. Ear puncher to identify individual mice.

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6. Antibiotics and fungicide to add to mouse drinking water for first 2 weeks post transplant: For antibiotics use either Bactrim oral suspension (cherry flavored children formulae) which is a combination of 8 mg/mL trimethoprim and 40 mg/mL sulfamthoxazole (use at 1:100 dilution) (Roche, Australia), or Ciprofloxacin 100 mg/50 mL (Bayer, Germany) dilute in drinking water to 125 μg/mL. For antifungal use Fluconazole solution 200 mg/100 mL (Pfizer) diluted 1:200 in drinking water. 2.7. Test Bleeds

1. Mouse restrainer. 2. Red lamp. 3. Surgical scalpel blades size 11. 4. Heparin: DBL heparin sodium (porcine mucous) 5,000 units in 0.2 mL (Hospira, Lake Forest, IL). 5. 10× red cell lysis buffer. 1.5 M NH4Cl, 100 mM NaHCO3, 10 mM EDTA pH 7.4. Sterile stock can be kept in fridge for many months. On the day of the experiment, dilute 1 part of 10× red cell lysis buffer in 9 parts of sterile water to make 1× red cell lysis buffer. 6. DPBS + 2% NCS. 7. Purified rat anti-mouse Fcγ receptor II/III clone 2.4 G2 (Fc Block) or hybridoma supernatant. 8. Stain tubes: 1.2 mL microtibertube. 9. Antibodies: CD45R (B220)-APCCY7 clone RA3-6B2 (BD) 0.2 mg/mL. CD11b-PECY7 clone M1/70 (BD) 0.2 mg/mL. CD45.1-PE clone A20 (BD) 0.2 mg/mL. CD45.2-APC clone 104 (BD) 0.2 mg/mL. CD3ε-FITC clone 145-2C11 (Biolegend) 0.5 mg/mL. 10. 7-AAD. 11. 8–9 color flow cytometer equipped with 405 nm violet laser, 488 nm blue laser, 635 nm red laser and filters as described above.

3. Methods We use the inbred mouse strain C57BL/6. This mouse strain expresses the Sca-1 (Ly6-A/E) antigen on HSPC and the CD45.2 alloantigen on all leukocytes. C57BL/6 mice have a relatively low

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mobilizing response relative to other inbred strains (such as BALB/c), but has the advantage that (1) most genetically modified mice including those used to identify signaling pathways involved in mobilization, are backcrossed into the C57BL/6 genetic background and (2) congenic CD45.1+ mice are available to track donor and competitive cells after transplantation. For competitive repopulation experiments, we use the C57BL/6 (mobilization treatment donor group) which expresses the CD45.2 alloantigen in combination with the congenic B6.SJL-PtprcaPep3b/BoyJ strain (CD45.1+) as recipients and as source of competing HSC (see Note 4). 3.1. Mobilization with rhuG-CSF

1. In the morning before injections, prepare a 25 μg/mL dilution of rhuG-CSF in saline in sterile flow cabinet for twice daily injections (injections may continue for up to 6 days). 2. Weigh mouse on scale. 3. Immobilize mouse by the neck scruff and inject subcutaneously (either in the scruff or in the belly) 100 μL of rhuG-CSF (25 μg/mL) per 20 g body weight. This corresponds to 125 μg per kg. Release mouse in the cage. 4. Repeat steps 2–4 twice daily for 2–6 days. The first day of injection is recorded as Day 0 and injections continue twice daily until euthanasia and tissue collection. Mobilization starts at day 2 of injections and peaks between days 4 and 6 of G-CSF administration.

3.2. Mobilization with Clodronate-Loaded Liposomes

1. To lightly anesthetize mice, place some gauze in a 50 mL tube and imbibe it with 50 μL of isoflurane. 2. Gently introduce the mouse in the tube, head first. Keep the mouse nose away from the gauze (isoflurane is an irritant). 3. When the mouse breathing slows down (after about 10 s), pull the mouse out of the tube and inject 200 μL liposome suspension (loaded with clodronate or with PBS as negative control) per 20 g body weight into the retroorbital sinus. 4. Replace the mouse in the cage. The mouse will wake up within a minute. 5. Repeat liposome administration every second day until euthanasia.

3.3. Mobilization with Cyclophosphamide

1. Weigh mouse on scale. 2. Immobilize mouse by the scruff and inject intraperitoneally 200 μL of cyclophosphamide (20 mg/mL) per 20 g body weight. This corresponds to 200 mg per kg. Release mouse in the cage.

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3.4. Tissue Harvest

At all times cells should be kept on ice and reagents and centrifuges at 4°C. 1. Before euthanasia record mouse weight. 2. To lightly anesthetize mice, put some gauze in a 50 mL tube and imbibe it with 50 μL of isoflurane. 3. Gently introduce the mouse in the tube, head first. Keep the mouse nose away from the gauze. 4. When the mouse breathing slows down (after about 10 s), retrieve the mouse from the tube and collect blood by cardiac puncture. To do this insert mounted 23 G needle into chest cavity and gently aspirate blood. 5. Collect up to 1 mL blood in a 5 mL polypropylene tube containing 15 units of heparin. Mix well. 6. Immediately euthanize mouse by cervical dislocation. 7. Remove skin to access and collect one femur using sterile scissors, tweezers, and scalpel. 8. Cut femoral epiphysis (ends) with scalpel, introduce 23 G needle mounted on 1 mL syringe loaded with 1 mL DPBS + 2%NCS into the femoral bone marrow cavity through the tibial plateau. Place bone with needle into 5 mL propylene tube and gently flush bone marrow out into the tube three to four times. Remove femur from needle with tweezers and insert needle this time in the epiphysis end of the femur and flush BM again three to four times (gently, do not foam). 9. Remove spleen from the peritoneal cage with tweezers and scalpel, record spleen weight. Transfer spleen to a propylene tube containing 1 mL DPBS + 2%NCS. Note that the size of the spleen will vary depending on the mobilizing agent and time-point. Spleen mobilized with rhuG-CSF will increase in size and weight, whereas spleen from a clodronate liposome mobilized mouse will decrease in size and weight. A ratio “g of spleen per body weight” can be calculated to show this association. 10. Discard the carcass following ethical procedures.

3.5. Cell Preparation

1. Blood: Take a 60 μL aliquot of whole blood and count blood leukocytes on Sysmex automated cell counter or by manual (microscopy) count of cells loaded on Neubauer counting chamber after 1:20 dilution in white cell counting fluid. Record leukocyte count per mL blood. Also transfer a portion of blood to new tube for colony assay (need 10 μL/dish in duplicate). An example of blood leukocyte concentration following mobilization treatment with rhu-GCSF, cyclophosphamide, or clodronate-loaded liposomes is given in Fig. 1.

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2. Spleen: Transfer each spleen into in 35 mm tissue culture petri dish and fill dish with 5 mL DPBS + 2%NCS. Mechanically dissociate spleen into single cell suspension by smearing between the frosted ends of two microscope slides. Dissociate the spleen progressively working from one end slowly to the opposite end. Transfer splenocyte suspension through 40 μm cell strainer nested in an open 50 mL tube. Wash slides and dish with another 5 mL DPBS + 2%NCS and transfer through the same strainer. Top up cell suspension volume to 10 mL with DPBS + 2%NCS . Take a 60 μL aliquot into Eppendorf tube and count leukocytes on automated Sysmex cell counter. Multiply splenocyte concentration/mL suspension by the volume (10 mL) of suspension to calculate the number of splenocytes per spleen. 3. BM: Dilute 20 μL whole bone marrow cell suspension with 80 μL DPBS + 2% NCS (1/5 dilution) into an Eppendorf tube. Count leukocytes on automated Sysmex cell counter. Multiply by 5 to obtain number of leukocytes per mL (or per femur). 3.6. Flow Cytometry Stains 3.6.1. Red Cell Lysis in Blood Samples

1. Add at least 3 mL of 1× Red Cell Lysis Buffer per mL of blood collected by cardiac puncture and incubate 6 min at room temperature (lyse 6 min if lysis buffer is at room temp, or lyse 10 min on ice if lysis buffer was at 4°C) all with gentle mixing (on a rotator). 2. Spin tubes at 370 × g for 5 min at 4°C. All the following steps are to be done on ice. 3. Aspirate supernatant (leave 500 μL as the cell pellet can rarely be seen at this stage). Mix well to resuspend cells then immediately add 4 mL of DPBS + 2%NCS and mix.

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4. Repeat wash (steps 2–3) but this time can remove more fluid from above cell pellet. 5. Resuspend the pellet in 1 mL final DPBS + 2%NCS. These blood leukocytes can now be used for antibody staining and flow cytometry. 3.6.2. Hematopoietic Stem Cell Staining

1. From the bone marrow flush (one femur in 1 mL DPBS + 2%NCS), lysed blood leukocyte or splenocyte suspension, transfer 5 × 106 cells into labeled stain tubes. 2. Fill up tubes with DPBS + 2%NCS 3. Spin at 370 × g for 5 min at 4°C. 4. Aspirate supernatant with cannula on vacuum line and leave 25 μL on cell pellet. Resuspend the cells by tapping the tubes with fingers or vigorous vortex (do not create foam). 5. Add 75 μL of antibody mix to each cell aliquot (final stain volume 100 μL for 5 × 106 cells). The antibody mix is made of lineage-biotin (CD3, CD5, B220, Gr1, CD11b, CD41, Ter119), Sca-1-PECY7, Kit-APC, CD48-FITC, CD150-PE (all at 1/300 dilution final) diluted in Fcblock hybridoma supernatant or in DPBS + 2%NCS + 2–5 μg/mL purified Fc Block antibody. 6. Mix then incubate on ice in the dark for 30 min with gentle rocking. 7. Wash stains with 1 mL of straight DPBS or MACS buffer. Repeats steps 3 and 4 (wash). 8. Add 75 μL of streptavidin-Alexa700 1/200 final in MACS buffer. 9. Repeat step 6 (mix and incubate). 10. Repeat steps 2–4 (wash). 11. Resuspend the cells in 300 μL of DPBS + 2%NCS (see Note 5). 12. For the single color controls (needed to set flow cytometer compensation values), add 0.2 μL of the designated antibody to 106 unstained control bone marrow cells in a final volume of 25 μL. For biotinylated antibodies stained with fluorochrome conjugated streptavidin, 0.2 μL of both biotinylated antibody and streptavidin can be added at the same time. After 30 min incubation on ice, wash the single color controls once as in steps 2–4 and proceed to step 11. 13. Analyze results with FloJo software. Report results as number of cells per femur, per mL of blood or per spleen using leukocytes counts recorded for each tissue and each mouse. An example of the gating strategy is shown in Fig. 2.

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Fig. 2. Mobilization of phenotypic HSPC in response to rhuG-CSF and clodronate-loaded liposomes. (a) Gating strategy to identify Lin− Sca1+ KIT+ (LSK) HSPC and LSK CD48− CD150+ HSC. Nucleated cells from a bone marrow flush from a saline treated mouse are gated according to forward scatter (FSC) and side scatter (SSC). Cell aggregates are gated out using signal pulse width, and viable cells are gated for their lack of 7-AAD staining. Typical gates of Lin- cells, LSK, and LSK CD48− CD150+ are then successively defined. (b) Time-course of LSK HSPC and LSK CD48− CD150+ HSC mobilization into the peripheral blood in response to rhu-GCSF. (c) Time-course of LSK HSPC mobilization into the peripheral blood in response to clodronate-loaded liposomes. Results are average ±SD of four mice per group. Significance were calculated with Student’s t test ***p £ 0.001, **0.001 < p £ 0.01 and *p £ 0.05.

3.6.3. Myeloid Stain on Bone Marrow Leukocytes

1. From the bone marrow flush (one femur in 1 mL DPBS + 2%NCS), transfer 106 cells (approximately 25 μL) into labeled stain tubes. 2. Fill up tubes with DPBS + 2%NCS. 3. Spin at 370 × g for 5 min at 4°C. 4. Aspirate the supernatant and leave 25 μL on cell pellet. Resuspend the cells by tapping the tubes with fingers. 5. Add 25 μL of 2× antibody mix to each cell aliquot. The 2× antibody mix is made of CD11b-PECY7, Ly6G-PE, F4/80Alexa647 for a final dilution of 1/300 of each in Fc block.

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6. Mix well. Incubate on ice in the dark for 30 min with gentle rocking. 7. For the single color controls, add 0.2 μL of the designated antibody to 106 unstained bone marrow cells (pooled cells from control mice) in a final volume of 25 μL. 8. Mix well and incubate 30 min on ice, in the dark with gentle rocking. 9. Repeat steps 2–4. 10. Resuspend the cells in 300 μL of DPBS + 2%NCS (see Note 5). 11. Analyze results with FloJo software. Report results as number of cells per femur using leukocytes counts recorded for each tissue and each mouse. An example of the gating strategy and result is shown in Fig. 3. 3.7. Preparation of Methylcellulose Semi-Solid Medium for Colony Forming Assays

1. Weigh 8.1 g methylcellulose into Shott bottle. 2. Autoclave bottle and flea on porous load twice. 3. Remove from autoclave and tighten cap. Leave to cool. 4. Place fresh 500 mL bottle on magnetic stirrer in hood. Aseptically add 200 mL of sterile 1× IMDM.

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5. Add a sterile large stirring flea to bottle with IMDM. 6. Start stirring and slowly add methylcellulose powder. Shake vigorously to disperse any large clumps. Once all the powder is added, use another 130 mL of IMDM to wash out the methylcellulose powder bottle and pour into medium bottle. 7. Stir at room temperature overnight with shaking every now and then when possible. Keep covered with aluminum foil at all times. 8. When the solution is homogenous and appears translucent, place back into laminar flow hood and aseptically add 180 mL of sterile FCS, 5.1 mL of Pen/Strep/Glutamine (×100 stock). 9. Aliquot aseptically 40 mL volumes into 50 mL falcon tubes. Label and store at −20°C. Note that Aliquots may be kept at 4°C if using within 2 weeks. 3.8. Colony Forming Assays

Before use, the methylcellulose:IMDM mix first requires addition of cytokines and insulin. 1. Make enough cytokine mix for all the samples in duplicate. The final concentrations are for 1 mL of methylcellulose medium per petri dish. 2. Final concentrations of additives are: Fluconazole 6 μg/mL final, mouse IL3-conditioned medium 1,000 units/mL (or 10 ng/mL recombinant mouse IL-3) final, mouse IL6-CM 1,000 units/mL final (or 10 ng/mL recombinant mouse or human IL-6), recombinant mouse or rat KIT ligand/SCF 100 ng/mL, human insulin at 10 μg (2.9 units)/mL final. 3. Make a master mix containing the required amount of additives per petri dish and then pipette master mix of additives/ cytokines onto a spot (around 50 μL in middle of each labeled 35 mm tissue culture petri dish). 4. On top of this cytokine mix, pipette exactly 10 μL of whole blood per dish or alternatively, 50,000 bone marrow leukocytes, or 50,000 splenocytes or 50,000 lysed blood leukocytes (from mobilized mice) or 200,000 splenocytes (from non mobilized control mice). 5. Using a 5 mL syringe mounted with a cannula, slowly aspirate the thawed methylcellulose and gently squeeze out 1 mL on top of the cells, constantly mixing with a sterile pipette tip to ensure even distribution of cells. 6. Cover with labeled lids and place in flat clean box in humid incubator at 37°C, 5%CO2. 7. Incubate for 7–10 days.

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8. Using dissection microscope, record number of colonies in each petri dish. Each colony is derived from a single deposited myeloid progenitor cell. Colonies should be >50 cells in size. Draw a grid on an overhead transparent sheet and put dishes on the grid on the microscope to help counting. 9. Report the colony forming units (CFU) per mL of blood, per femur, or per spleen using leukocytes counts in blood, bone marrow, and spleen previously recorded. An example of CFU/mL of blood is shown in Fig. 4. 3.9. Competitive Repopulation Assays

Colony forming assays measure the number of myeloid progenitors in blood, bone marrow and spleen. Although it is often a good approximation of HSC, this is however not always the case (17, 18). The long-term competitive repopulation assay is the gold standard to demonstrate the mobilization of genuine HSC able to fully reconstitute all blood lineages in a lethally irradiated host, reviewed by (19). In this assay, donor cells containing an unknown number of HSC are put into competition with a standard dose of HSC from a congenic strain to repopulate the host. This competitive transplant assay has the advantage that (1) all mice survive (ethically more acceptable) and (2) more reliable and consistent results than noncompetitive transplants. In this manner, the level of donor chimerism at the end of the experiment (16 weeks post transplant) is proportional to the number of HSC initially present in the test donor sample and can be calculated as “reconstitution units” (RU) with an equivalent number of HSC as 100,000 healthy competing BM cells using the formulae described by Harrison et al. (19, 20).

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1. Cage 8-week-old female B6.SJL-PtprcaPep3b/BoyJ recipients in order to have ten mice per recipient group. Keep two to three B6.SJL-PtprcaPep3b/BoyJ mice nonirradiated as a source of competing HSC. 2. Lethal-irradiation of recipients: On day before transplant, irradiate recipient mice 5.5 Gy in the morning. Put mice back in cages. Irradiate a second time with 5.5 Gy with a minimum 4 h interval between the 2 irradiation doses. This split dosing decreases acute radio toxicity on other vital organs (kidney, gut, heart). 3. On experimental day, collect heparinized blood by cardiac puncture from mobilized mice as described in Subheading 3.4. 4. Donor blood preparation: Per treated group, pool the same amount of mobilized donor blood from each individual donor mouse in order to have 25–50 μL donor blood per recipient mouse. The exact volume of blood used in a transplant will depend on level of expected donor HSC mobilization. Typically after 4 days of G-CSF injections 25 μL mobilized blood provides equivalent HSC as 200,000 healthy competing BM cells (see Note 6). 5. Competing BM preparation: Sacrifice two to three healthy nonirradiated B6.SJL-PtprcaPep3b/BoyJ mice by cervical dislocation and harvest bone marrow leukocytes by flushing femurs as described in Subheading 3.4. These bone marrow leukocytes will be used as healthy competing CD45.1 HSC. 6. Filter these competitive bone marrow leukocytes from through a 40 μm cell strainer into a 50 mL tube in order to eliminate any bone chips. 7. Count bone marrow cells. 8. Transfer the necessary amount of mobilized CD45.2 blood and competing CD45.1 bone marrow leukocytes into a 14 mL round bottom tube. For each individual recipient within a group, count 25 or 50 μL of mobilized donor blood together with 200,000 competitive bone marrow leukocytes from the B6.SJL-PtprcaPep3b/BoyJ mice in a final volume of 180 μL per recipient. Multiply these numbers by the number of recipients in the recipient group (usually 10). 9. Top up tube to 10 mL with saline + 2%FCS. Spin at 300 × g, for 10 min at 4°C. Resuspend cells in saline + 2%FCS in order to have a volume of 180 μL per recipient. 10. Just before injection add 20 μL (20 units) of heparin per recipient mouse to the mix. Heparin is an antithrombolytic agent. Administration of 20 units heparin to mice at/or just prior to blood transplant will limit thrombosis forming around transplanted “platelet clumps” which can be fatal.

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11. Inject retroorbitally 200 μL of the blood/bone marrow transplant mix per recipient. 12. Identify each mouse by ear punch. 3.9.2. Bone Marrow Transplant

1. Donor: Per treated group, pool the CD45.2 bone marrow from one femur from each mobilized donor. 2. Filter the bone marrow flush using a 40 μm cell strainer into a 50 mL tube to eliminate any bone chip. 3. Count CD45.2 bone marrow cells. 4. Sacrifice two to three nonirradiated B6.SJL-PtprcaPep3b/ BoyJ CD45.1 mice by cervical dislocation and harvest bone marrow leukocytes by flushing femurs as described in Subheading 3.4. These bone marrow leukocytes will be used as a source of competing HSC. 5. Filter competitive bone marrow leukocytes from B6.SJLPtprcaPep3b/BoyJ mice on a 40 μm cell strainer into a 50 mL tube in order to eliminate any bone chips. 6. Count competing CD45.1 bone marrow cells. 7. Per recipient mouse in each group, mix 200,000 donor CD45.2 mobilized bone marrow leukocytes together with 200,000 competitive bone marrow leukocytes from the B6.SJLPtprcaPep3b/BoyJ CD45.1 mice in a final volume of 200 μL per recipient. 8. Transfer the necessary amount of cells into a 14 mL round bottom tube. Top up tube to 10 mL with saline + 2%FCS. Spin at 300 × g for 10 min at 4°C. Resuspend cells into saline + 2%FCS in order to have a volume of 200 μL per recipient. 9. Inject retroorbitally 200 μL of the bone marrow transplant mix per recipient. 10. Identify each mouse by ear punch.

3.9.3. Spleen Transplant

1. Extract splenocytes from each individual spleen as described in Subheading 3.4. 2. For each donor group, mix the donor splenocytes from each individual donor mouse in equal proportion in order to transplant the equivalent of 1% of one spleen into each recipient. 3. Transfer the necessary amount of cells into a 14 mL round bottom tube. Top up tube to 10 mL with saline + 2%FCS. Spin at 300 × g for 10 min at 4°C. Resuspend cells in saline + 2%FCS. 4. Per recipient mouse, mix 1% of CD45.2 spleen with 200,000 competitive CD45.1 bone marrow cells from B6.SJLPtprcaPep3b/BoyJ in 200 μL final per recipient. 5. Retroorbitally inject 200 μL of the spleen : BM cell mix per recipient. 6. Identify each mouse by ear punch.

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1. Add antibiotics and antifungal to the drinking water: Bactrim 1/100 dilution and Diflucan 1/200 dilution and cover bottle with foil to protect from light damage. Assume that mice drink 5 mL per day (~1/4 of their body weight) and weigh approximately 20 g. Change the antibiotics every 3 days. Administer for 14 days only. 2. Record the weight of the mice on the day of the transplant. 3. Score individual mouse health, well-being, and weight on the day following transplant and daily for first 4–10 days at least. After 10 days continue daily monitoring until mice regain their original weight following the guidelines in Table 1 (see Note 7).

3.9.5. Posttransplant Follow-Up and Test Bleed

Test bleed the transplanted mice 6, 12, and 16 weeks post transplant to record CD45.2 donor versus CD45.1 competitor chimerism in myeloid and lymphoid lineages. 1. Warm up the mice under a red lamp. Make sure not to burn the mice. 2. Using a mouse restrainer, make a small incision on one of the side vein of the tail. 3. In a 1.5 mL Eppendorf tube, collect 30 μL (or 3 drops) of blood into 3 units of heparin (can also add EDTA to final 4 mM). Mix well. Keep tube on ice. 4. On top of collected blood, add 500 μL of 1× Red Cell Lysis Buffer and incubate for 6 min at room temperature. Mix gently. 5. Spin tubes at 370 × g for 5 min at 4°C. 6. All the following steps are on ice. 7. Aspirate supernatants and immediately resuspend cell pellets in 1 mL DPBS + 2%NSC. 8. Transfer the samples to bullet stain tubes. 9. Pool some cells from the samples to constitute the single color controls (one unstained and one for each individual fluorophore). 10. Spin tubes at 370 × g for 5 min at 4°C. 11. Aspirate supernatants. Leave 25 μL cell pellet. Resuspend cells well. 12. Add 25 μL of 2× antibodies mix to the cells. The 2× antibody mix is made of CD45R/B220-APCCy7, CD11b-PECy7, CD45.1-PE, CD45.2-APC, CD3ε-FITC (for final of 1/200 dilution of each) in Fc block buffer. 13. For the single color controls, add 0.2 μL of each single fluorescent antibody into separate tubes containing 25 μL cells.

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14. Mix well and incubate for 30 min on ice, in the dark with gentle rocking. 15. Wash stains with 1 mL of DPBS + 2%NSC. Spin tubes at 370 × g for 5 min at 4°C. 16. Resuspend cells in 300 μL of DPBS + 2%NSC. 17. Analyze on a flow cytometer after addition of 2 μg/mL 7-AAD to gate on viable cells expression of CD45.2 (donor HSC) and CD45.1 (competing HSC) among CD11b+ myeloid cells, B220+ B cells, and CD3+ T cells. An example of the gating strategy is given in Fig. 5. Typical results of long-term competitive repopulation with mobilized blood are given in Fig. 6.

4. Notes 1. As cells from mouse tissues will be transplanted, all buffers must be prepared sterile and autoclaved or sterile filtered through 0.22 μm filters. We recommend using clinical grade “water for irrigation” (Baxter #AHF7114) to prepare and dilute buffers, and clinical grade injectable saline solution. Cell harvesting and processing should be performed in a sterile flow

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Fig. 6. Analysis of HSC mobilization in response to clodronate-loaded liposomes using a long-term competitive reconstitution assay. Test donor CD45.2+ mice were injected with clodronate-loaded liposomes (clo-lip) for 2 or 4 days, or control PBS-loaded liposomes (PBS-lip) for 4 days. 50 μL blood was transplanted in competition with 200,000 competing CD45.1+ BM cells into lethally irradiated CD45.1+ recipients. Donor CD45.2+ contribution was measured in CD45+ leukocytes, myeloid, B and T cells 16 weeks after transplantation following gating strategy shown in Fig. 5. Lineage chimerism was estimated to be positive when proportion of CD45.2+ donor cells was above 0.5% in each lineage. ***p £ 0.001, **0.001 < p £ 0.01 and *p £ 0.05 between PSB-lip and clo-lip treated mice. Multi-lineage chimerism is considered positive when the proportion of CD45.2+ donor cells is above 0.5% simultaneously in each individual lineage.

cabinet. Similarly all tubes and syringes involved in collection and transplantation must be sterile. 2. If the necessary equipment or skills are not available to synthesize liposomes, ready-to-use clodronate-loaded liposomes and control PBS-loaded liposomes can be directly obtained from Dr Nico van Rooijen (contact at www.clodronateliposomes. org). Of note, liposomes are sent in nonrefrigerated FedEx letter pack. It is therefore important to organize delivery on days with temperatures below 30°C. Once received, liposomes should be kept in the fridge and used within 3–4 weeks. Beyond 4 weeks, their activity decreases. 3. Cyclophosphamide should be used within a month following resuspension in saline. Inject 10% more per week of storage to compensate for gradual loss of activity. 4. We use 8–12-week-old male mice in mobilization treatment groups to reduce variability due to estrus cycle. We use females as long-term transplantation recipients to avoid fighting which inevitably occurs between mature males caged together over 16 weeks. 5. Note that 7-AAD can be added to the cells 10 min before analyzing samples at a final concentration of 2 μg/mL to gate viable 7-AAD-negative cells. 6. The maximum volume of heparinized blood that can be safely transplanted is 50 μL diluted in 200 μL final volume per recipient mouse. To calculate how much blood to prepare, for instance, if one elects to administer 50 μL mobilized blood per

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recipient mouse for a group of ten recipients, one will need a total volume of 500 μL pooled mobilized blood. Therefore, if the donor group comprises four mice, the 1 mL pool of mobilized blood will be made by pooling 250 μL aliquots of mobilized blood from each individual donor mouse. 7. As all recipients received HSC contained in the 200,000 healthy competing bone marrow cells, they should all survive. It is critical to maintain clean cages and keep mice away from pathogens in first 3 weeks post transplant. Death within 24 h of transplantation is usually due to thrombolysis (add more heparin). Death/poor condition between 5 and 14 days is usually due either to infections or to failure to engraft (check for small white spleen and liver), which may result from incorrect intravenous injection of transplant or insufficient dose of healthy competing BM cells.

Acknowledgments J.P.L. is supported by a Senior Research Fellowship from the Cancer Council of Queensland, I.G.W. by a CDA fellowship form the National Health and Medical Research Council of Australia. This work was supported by project grants 434515, 543706, and 350406 from the National Health and Medical Research Council. References 1. Liu F, Poursine-Laurent J, Link DC (1997) The granulocyte colony-stimulating factor receptor is required for the mobilization of murine hematopoietic progenitors into peripheral blood by cyclophosphamide or interleukin-8 but not flt-3 ligand. Blood 90:2522–2528 2. Liu F, Poursine-Laurent J, Link DC (2000) Expression of the G-CSF receptor on hematopoietic progenitor cells is not required for their mobilization by G-CSF. Blood 95: 3025–3031 3. Pelus LM, Bian H, King AG, Fukuda S (2004) Neutrophil-derived MMP-9 mediates synergistic mobilization of hematopoietic stem and progenitor cells by the combination of G-CSF and the chemokines GROb/CXCL2 and GRObT/CXCL2D4. Blood 103:110–119 4. Lévesque JP, Hendy J, Takamatsu Y, Williams B, Winkler IG, Simmons PJ (2002) Mobilization by either cyclophosphamide or granulocyte colony-stimulating factor transforms the bone marrow into a highly proteolytic environment. Exp Hematol 30:440–449

5. Lévesque JP, Takamatsu Y, Nilsson SK, Haylock DN, Simmons PJ (2001) Vascular cell adhesion molecule-1 (CD106) is cleaved by neutrophil proteases in the bone marrow following hematopoietic progenitor cell mobilization by granulocyte colony-stimulating factor. Blood 98:1289–1297 6. Lévesque JP, Hendy J, Takamatsu Y, Simmons PJ, Bendall LJ (2003) Disruption of the CXCR4/CXCL12 chemotactic interaction during hematopoietic stem cell mobilization induced by GCSF or cyclophosphamide. J Clin Invest 111:187–196 7. Petit I, Szyper-Kravitz M, Nagler A, Lahav M, Peled A, Habler L, Ponomaryov T, Taichman RS, Arenzana-Seisdedos F, Fujii N, Sandbank J, Zipori D, Lapidot T (2002) G-CSF induces stem cell mobilization by decreasing bone marrow SDF-1 and up-regulating CXCR4. Nat Immunol 3:687–694 8. Lévesque JP, Liu F, Simmons PJ, Betsuyaku T, Senior RM, Pham C, Link DC (2004) Characterization of hematopoietic progenitor

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mobilization in protease-deficient mice. Blood 104:65–72 9. Semerad CL, Christopher MJ, Liu F, Short B, Simmons PJ, Winkler I, Lévesque J-P, Chappel J, Ross FP, Link DC (2005) G-CSF potently inhibits osteoblast activity and CXCL12 mRNA expression in the bone marrow. Blood 106: 3020–3027 10. Christopher MJ, Link DC (2008) Granulocyte colony-stimulating factor induces osteoblast apoptosis and inhibits osteoblast differentiation. J Bone Miner Res 23:1765–1774 11. Christopher MJ, Liu F, Hilton MJ, Long F, Link DC (2009) Suppression of CXCL12 production by bone marrow osteoblasts is a common and critical pathway for cytokine-induced mobilization. Blood 114:1331–1339 12. Katayama Y, Battista M, Kao WM, Hidalgo A, Peired AJ, Thomas SA, Frenette PS (2006) Signals from the sympathetic nervous system regulate hematopoietic stem cell egress from bone marrow. Cell 124:407–421 13. Mendez-Ferrer S, Lucas D, Battista M, Frenette PS (2008) Haematopoietic stem cell release is regulated by circadian oscillations. Nature 452:442–447 14. Winkler IG, Sims NA, Pettit AR, Barbier V, Nowlan B, Helwani F, Poulton IJ, van Rooijen N, Alexander KA, Raggatt LJ, Lévesque J-P (2010) Bone marrow macrophages maintain hematopoi-

etic stem cell (HSC) niches and their depletion mobilizes HSC. Blood 116:4815–4828 15. Van Rooijen N, Sanders A (1994) Liposome mediated depletion of macrophages: mechanism of action, preparation of liposomes and applications. J Immunol Methods 174:83–93 16. van Rooijen N, Hendrikx E (2010) Liposomes for specific depletion of macrophages from organs and tissues. Methods Mol Biol 605: 189–203 17. Adams GB, Martin RP, Alley IR, Chabner KT, Cohen KS, Calvi LM, Kronenberg HM, Scadden DT (2007) Therapeutic targeting of a stem cell niche. Nat Biotechnol 25:238–243 18. Herbert KE, Walkley CR, Winkler IG, Hendy J, Olsen GH, Yuan YD, Chandraratna RA, Prince HM, Lévesque JP, Purton LE (2007) Granulocyte colony-stimulating factor and an RARalpha specific agonist, VTP195183, synergize to enhance the mobilization of hematopoietic progenitor cells. Transplantation 83:375–384 19. Purton LE, Scadden DT (2007) Limiting factors in murine hematopoietic stem cell assays. Cell Stem Cell 1:262–270 20. Harrison DE, Jordan CT, Zhong RK, Astle CM (1993) Primitive hemopoietic stem cells: direct assay of most productive populations by competitive repopulation with simple binomial, correlation and covariance calculations. Exp Hematol 21:206–219

Chapter 12 Combinatorial Stem Cell Mobilization in Animal Models Simon C. Pitchford and Sara M. Rankin Abstract It has long been recognized that single therapies, such as G-CSF, have a limited capacity to mobilize hematopoietic progenitor cells from the bone marrow. As a consequence in ~20% of patients insufficient numbers of HPCs are mobilized to perform a bone marrow transplant. Recent studies have shown synergistic mobilization of HPCs when G-CSF pretreatment is combined with acute administration of a CXCR4 antagonist suggesting that combinatorial therapies may have therapeutic potential. In addition to HPCs, endothelial progenitor cells (EPCs) and mesenchymal stem cells (MSCs) reside in the bone marrow. These progenitor cells contribute to tissue regeneration and there is currently much interest in identifying the factors and mechanisms that regulate their mobilization. We describe a methodology for an in situ perfusion system of the mouse hind limb that permits direct quantification of stem and progenitor cell egress from the bone marrow. Progenitor cells are quantified by colony forming assays and immunohistochemistry. A strength of the methodology described is the ability to simultaneously quantify the mobilization of HPCs, EPCs and MSCs. Using this system we have shown that it is possible to achieve differential mobilization of these stem cell subsets using discrete combination therapies. Identification of such novel pharmacological regimens that stimulate the selective mobilization of EPCs and MSCs might be exploited in the future for tissue regeneration. Key words: Mobilization, HPC, EPC, MSC, Mouse, VEGF, G-CSF, VEGF, CXCR4

1. Introduction Pharmacological agents have been used clinically for many years to stimulate the mobilization of hematopoietic stem/progenitor cells (HSC/HPC) from the bone marrow for bone marrow transplants. The limitation of such therapies has been in the absolute numbers of HPCs mobilized by any given treatment. Thus, in approximately 20% of patients treated with G-CSF, insufficient HPCs are harvested to perform a bone marrow transplant (1). Recently it has

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_12, © Springer Science+Business Media, LLC 2012

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been shown in animal models and clinical trials that combination therapies can more effectively mobilize HPCs from the bone marrow. In particular a synergistic mobilization of HPCs is reported when G-CSF treatment is combined with a CXCR4 antagonist (2–4). Notably the relative timings of these treatments is critical with G-CSF administered over 3–5 days and CXCR4 antagonist as a single dose a couple of hours before stem cell collection. In addition to HPCs, the adult bone marrow contains other distinct subsets of progenitor cells including endothelial progenitor cells (EPCs) and mesenchymal stem cells (MSCs). These progenitor cells are thought to play a role in tissue regeneration and thus therapies that mobilize MSCs and EPCs have a potential clinical application in the field of regenerative medicine. EPCs have been reported to facilitate post natal neovascularization in animal models of disease and multiple studies reveal that the contribution of bone marrow progenitor cells in angiogenesis is greater, the more severe the ischemic signal (5–8). The identification of pharmacological agents that stimulate EPC mobilization might be highly desirable as a therapeutic strategy to promote tissue angiogenesis and thereby aid healing. MSCs exhibit a trilineage differentiation potential, with the ability to readily differentiate into adipocytes, osteoblasts, and chondrocytes (9, 10). There are some reports that MSCs may also differentiate into skeletal muscle, cardiac myocytes, smooth muscle, and neurons, and as such, MSC mobilization from the bone marrow could be exploited to restore function of diverse tissue types that have become injured. However, there are very few reports of successful MSC mobilization using pharmacological agents, with G-CSF treatment giving contradictory results (11–13). Given the lack of understanding pertaining to the molecular mechanisms by which these distinct stem and progenitor cell types are mobilized, it is necessary to develop technologies that will allow the accurate quantification of their rates of egress from the bone marrow. Such techniques would allow comparisons to be made of the efficacy of various pharmacological agents to induce mobilization of HPCs, EPCs, and MSCs and to record total numbers of stem and progenitor cells mobilized over a period of time. This data is difficult, or impossible to collate from peripheral blood measurements of circulating stem cells for the following reasons: (1) The source of circulating progenitor cells is unknown, i.e., progenitor cells present in the blood may be mobilized from the bone marrow, spleen or other extravascular sites; (2) stem cells have the propensity to rapidly home back to bone marrow after mobilization; (3) detection of rare stem cell subsets is technically difficult; and (4) stem cell mobilizing agents may have both local and systemic effects that cause direct and indirect mobilization (14). We have therefore developed an in situ perfusion system of the mouse

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hind limb, combined with the detection of mobilized progenitor cell subsets with carefully characterized colony forming assays, to directly assess stem cell mobilization from the bone marrow. Using this system we have shown that known mobilizing factors (G-CSF, VEGF, and a CXCR4 antagonist) used alone or in combination differentially mobilize HPCs, EPCs, and MSCs (13). Specifically, optimal and synergistic HPC mobilization was achieved when the CXCR4 antagonist was infused into mice pretreated with G-CSF. Optimal and additive EPC mobilization was achieved when the CXCR4 antagonist was infused into mice pretreated with VEGF, but not with G-CSF. Interestingly, VEGF pretreatment combined with CXCR4 antagonist infusion was the only treatment out of all possible permutations where MSC mobilization was achieved (13). Furthermore, HPC mobilization was significantly reduced using the treatment combination of VEGF pretreatment combined with acute infusion of CXCR4 antagonist compared to the infusion of the CXCR4 antagonist alone (13). These results demonstrate that it is possible to directly compare the efficacy of combinatorial stem cell mobilization regimens in vivo, using this technique. This system can therefore be used to determine the optimal strategies, drugs, and dosing regimens to mobilize specific stem cell subsets. The methodology used to achieve this is outlined below.

2. Materials 2.1. Buffers and Reagents for In Situ Perfusion of the Murine Hind Limb

1. Krebs Ringer buffer with phenol red (Sigma, Poole UK) is supplemented with 0.37 g CaCl2/l and stored at −20°C. On the day of experiment: 45 ml Krebs Ringer solution is defrosted and supplemented with 5 ml fetal bovine serum (FBS), 5,000 U/ml penicillin, 5,000 μg/ml streptomycin, 250 μg/ ml amphotericin B (1/500, fungizone), 0.12 g of NaHCO3, and 2 g of ficoll. Adjust pH to 7.4. 2. Anticoagulants: 5,000 U/ml heparin (multiparin) diluted to 500 U/ml in PBS. Acid citrate dextrose (ACD). Store at 4°C. 3. VEGF, G-CSF (Peprotech, NJ, USA) dissolve to 10 μg/ml in PBS (0.1%BSA) and store in single use aliquots at −20°C. 4. CXCR4 antagonist (AMD3100, Sigma) dissolve in PBS to 250 nM and store at 4°C. 5. 25% urethane solution. Dissolve 6.25 g of urethane (Sigma) in 25 ml saline. It is important to weigh the urethane accurately as this will need to maintain anesthesia for 90 min. Store at room temperature.

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6. Sundries: tubing (fine bore tubing, 0.28 mm ID, 0.61 mm OD) (Portex, London, UK). Three way stopcocks (Tyco, Gosport, UK), Minipuls 3 peristaltic pump (Anachem, Luton, UK), peristaltic pump tubes (0.76 mm BK-BK, Anachem), infusion syringe pump (Harvard apparatus, Kent, UK). Perfusate collecting tubes: red top (glass) vacutainers containing no additives (BD Biosciences, NJ, USA). 2.2. Perfusate Preparation for Cell Culture

1. Dulbecco’s Modified Eagle’s Medium (DMEM with 4.5 g glucose, L-glutamine, and pyruvate), supplemented with penicillin/streptomycin (1/500 dilution) and fungizone (1/250 dilution). Store at 4°C. 2. DMEM, with 20% FBS, penicillin/streptomycin (1/500 dilution) and fungizone (1/250 dilution). Store at 4°C. 3. Hypotonic saline (0.2%), Dissolve 0.2 g NaCl in 100 ml deionized water. Store at room temperature. 4. Hypertonic saline (1.6%), Dissolve 1.6 g NaCl in 100 ml deionized water. Store at room temperature. 5. 0.2% Methylene blue cell counting solution: Make 1% w/v solution of methylene blue (Sigma) in water and dilute to 0.2% in 1% acetic acid solution. Store at room temperature.

2.3. Cell Culture of CFU-HPC, CFU-EPC, and CFU-MSC

1. Murine MethoCult with growth supplements (StemCell Technologies, Grenoble, France), mix well and aliquote in 5 ml batches and store at −20°C. 2. Non treated cell Technologies).

culture

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dishes

(StemCell

3. Bovine fibronectin (Sigma). Dissolve in PBS (10 μg/ml). First dissolve powder in small volume (1 mg/ml) at 37°C (2 h) before aliquoting to working dilution. Store at −20°C (see Note 1). 4. EPC Assay medium: EBM + supplements (EGM-2 medium Cambrex, IA, USA). Store basal medium at 4°C, but aliquot supplements 1/10 and store separately at −20°C. 5. Other EPC medium supplements: VEGF (Peprotech), 10 μg/ ml in PBS (0.1%BSA) and store at −20°C. FBS, Pen/Strep, Fungizone. 6. Tissue culture treated 35 mm2 dishes (Corning, NY, USA). 7. Murine MesenCult + supplements, (Stemcell Technologies). Combine basal medium with supplements and store at 4°C for 1 month. 8. 8 well culture (chamber) slides, 0.7 cm2 (BD Biosciences).

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1. Primary antibodies: goat anti-mouse VEGFR1 antibody, rabbit anti-mouse VEGFR2 antibody, rat antimouse CD34 antibody, and goat anti-mouse VE-Cadherin antibody (Santa Cruz Biotechnology, CA, USA). Rat anti-mouse CD45 antibody, rat anti-mouse CD29 antibody, and rat anti-mouse CD105 antibody (BD Pharmingen Oxford, UK). Rat anti-mouse CD115 antibody (eBioscience Wembley, UK). Rabbit anti-human vWF antibody (DAKO, Glostrup, Denmark). Store at 4°C. 2. Isotype controls: Rat IgG1 (BD Pharmingen), goat IgG and rabbit IgG (Santa Cruz Biotechnology). Store at 4°C. 3. Fluorescently labeled secondary antibodies: donkey anti-goat Alexafluor 568 antibody, goat anti-rat Alexafluor 594 antibody, goat anti-rabbit Alexafluor 488 antibody (Molecular Probes, Paisley, UK). Store at 4°C. 4. DIL-labeled acetylated low density lipoprotein (DIL-Ac-LDL, molecular probes), make 1 mg/ml (in PBS) stock solution and store at 4°C. 5. Fluoroscein labeled Griffonia Simplicifolia (GS)-Lectin (Vector Labs, Peterborough, UK) 5 mg/ml (in PBS) stock solution and store at 4°C. 6. In vitro EC Matrix angiogenesis assay (Millipore). Store at −20°C.

3. Methods In order to compare the potencies of various drug candidates for their ability to mobilize stem/progenitor cells, or indeed their ability to mobilize distinct stem cell subsets, it is necessary to measure the absolute numbers of stem/progenitor cells released from the bone marrow over a defined time period. The number of collected stem/progenitor cells can therefore be correctly attributed to their rate of egress out of the bone marrow (14). As discussed above, the comparison of mobilization regimens by enumerating stem cell subsets in peripheral blood is associated with a number of shortcomings (14). We have therefore developed a technique for the in situ perfusion of the mouse hind limb to assess progenitor cell mobilization from the bone marrow. This technique was originally developed by us to investigate leukocyte egress in animal models of inflammation (15–17). Thus while the methodology described below is for murine studies we have previously used the same technique in rats and guinea pigs. In this system, the femoral artery and vein are cannulated in situ such that the vasculature of the femur

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and tibia bone marrow is perfused directly, in isolation from the systemic circulation. Mobilizing agents may be infused via the femoral artery, and mobilized progenitor cells collected via the femoral vein. Of note, because the bone marrow vasculature is perfused with buffer and drugs are infused in buffer directly into the vasculature of the bone marrow it is possible to determine the relative potency of different drugs with respect to their ability to stimulate stem cell mobilization without the impact of differences in their pharmacokinetics (3). This technique can also be used to compare the potencies of combinatorial mobilization therapies (3, 13). Specifically we have examined the impact of pretreating mice over 4 days with G-CSF or VEGF on subsequent acute mobilization of progenitor cells with a CXCR4 antagonist infused directly into the hind-limb of mice (13). Using this system, and permutations thereof of these mobilizing agents, has allowed us to define novel treatment strategies leading to the differential mobilization of HPCs, EPCs, and MSCs. It has also enabled us to acquire the necessary assay sensitivity to detect mobilized MSCs (13). Other rare circulating stem or progenitor cells might, therefore, be detected using this assay. Our method for detecting mobilized stem and progenitor cells from the collected perfusate relies on both a functional aspect of stem cells (their ability to form colonies in vitro, determined as colony forming units-CFU, or burst forming units-BFU), and the identification of antigenic markers on colonies that are particular to distinct stem cell subsets (in order to exclude the possibility of contamination with other proliferative cell types). Whilst flow cytometry methods have been used to quantify mobilized stem cells, this procedure does not evaluate the colony forming ability of mobilized cells. In addition, EPCs and MSCs represent a very small population of circulating cells, even when numbers are boosted in the peripheral circulation by mobilizing agents, thus it is difficult to accurately quantify absolute numbers of these cells by flow cytometry. Further, given the low absolute number of progenitor cells mobilized and the requirement for multiple antigens to identify a single population of stem cells by flow cytometry, use of this methodology would restrict the analysis to a single population of progenitor cells per animal. A major advantage of quantifying progenitor cells by colony assays is, therefore, the ability to perform at least three different colony assays from a single perfusate, permitting direct comparison with respect to mobilization of distinct subsets of progenitor cells. 3.1. In Situ Perfusion of the Murine Hind Limb

1. All in vivo studies (locally) are to be carried out under the local ethical approval, such as United Kingdom’s Animals (Scientific Procedures) Act of 1986. 2. Administer VEGF (2.5 μg/mouse i.p.), G-CSF (2.5 μg/mouse i.p.), or vehicle on 4 consecutive days to Balb/c mice (8–10 weeks old). Volume of injection is 0.2 ml.

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3. On day 5, terminally anesthetize mice with 25% urethane, to last for approximately 90 min (see Note 2). 4. The surgical procedure for preparing the mice for perfusion has been recorded and animated elsewhere (14). The reader is strongly advised to use this resource when learning how to perform the procedure. Initially, pump 70% ethanol through the tubing to ensure that the system is as sterile as possible. Expose the femoral artery and vein and the circulatory system of the hind limb. Then isolate the circulatory system by occlusion of the external iliac artery, superficial epigastric and muscular branch. Insert stretched polyethylene cannulae into the femoral artery and vein (heparinized) as described (14) (see Note 3). Infuse perfusion buffer (modified Krebs Ringer solution, 37°C, gassed with 95% O2 5% CO2) via the arterial cannula and remove from the venous cannula using a Minipuls peristaltic pump. Perfuse the hind limb for an initial 2 min to remove remaining blood from the vasculature and then perfuse for a further 60 min. Infuse either vehicle alone or the CXCR4 antagonist (AMD3100, 0.1 mM, 0.4 ml) over the first 10 min using an infusion/ withdrawal pump connected via a three-way stopcock (14). 5. Collect perfusate in vacutainers with ACD (15% final volume) and then use for assays outlined below to enumerate mobilized HPCs (CFU-HPC), EPCs (CFU-EPC), MSCs (CFU-F). 6. A 60 min perfusion will result in 6–8 ml of perfusate collected. This will appear quite bloody as peripheral blood vessels dilate in an effort to restore blood supply to the isolated hind limb. This is impossible to stop, but importantly, infused drugs and mobilized cells are unable to enter the systemic circulation due to the positioning of the venous cannula (see Note 4). 3.2. Cell Culture of Mobilized Cells

1. In a tissue culture hood (to avoid infection), dispense perfusate into 50 ml Falcon tube, make sure that perfusate is gently (but thoroughly) mixed with ACD anticoagulant (15% final vol). Centrifuge at 180 × g for 5 min at room temperature. Aspirate off top liquid to leave cell pellet (see Note 5). Resuspend pellet up to 1 ml with DMEM (0% FBS). 2. Take 10 μl of cell solution and add to 190 μl methylene blue solution for total leukocyte count (pre lysis). Count with an improved Neubauer hemocytometer using a light microscope and ×20 objective lens (see Note 6). 3. Lyse red cells by washing perfusate pellet for 3–5 s in 12 ml sterile 0.2% NaCl before solution is turned to normal tonicity by addition of 12 ml 1.6% NaCl. Add 24 ml DMEM +20%FBS.

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4. Centrifuge at 180 × g (5 min) and resuspend pellet in 1 ml DMEM +20%FBS. The pellet is still not stuck hard to bottom as it still contains some red blood cells due to the mild lysis protocol (see Note 7). 5. Take 10 μl washed perfusate and add to 40 μl methylene blue solution for total leukocyte count (post lysis). Count with an improved Neubauer hemocytometer using a light microscope and ×20 objective lens. This assesses the degree of inadvertent leukocyte lysis during the red cell lysis procedure. This reading is used to determine seeding density of cells onto dishes as described below. 6. Plate as described in Subheading 3.3–3.5. 3.3. MethoCult Assay for Culture of CFU-HPC

1. Defrost MethoCult at 4°C, not at room temperature. 2. In a tissue culture hood: Add 1.1 ml MethoCult medium to 1.5 ml eppendorf. Add 30 μl of Pen/Strep and 15 μl Fungizone. 3. Add 5 × 104 cells to 1.1 ml medium and vortex. 4. Dispense 1.1 ml of medium to each nontreated 35 mm2 dish using a 1 ml syringe and 19 G needle, making sure that the medium is evenly spread. Air bubbles will disperse with incubation. 5. Place dish in larger petri dish and include a 35 mm2 dish with 3 ml sterile water Incubate for 12 days in a tissue culture incubator (37°C, 5% CO2, 95% O2), then analyze colonies with inverted microscope as described in Subheading 3.6.

3.4. EPC Assay for Culture of CFU-EPC

1. In a tissue culture hood: Coat 35 mm2 dish with fibronectin (10 μg/ml, 1.25 ml per well). Incubate at 37°C for 4 h. Rinse dishes with PBS before second step. 2. Make sufficient EPC Assay medium. For convenience, defrost EGM-2 supplements to make a total of 50 ml fluid. Thus, add supplements to 40 ml of EGM-2 basal medium. Add 50 ng/ ml VEGF (25 μl stock), 16% FBS (8 ml), 500 μl of Pen/Strep and 250 μl Fungizone. Vortex hard to ensure mix of ingredients. 3. Plate cells at 5 × 105 cells/dish. Add cells to 3 ml medium per dish (0.5 × 105 cells/cm2) using tissue culture treated plastic dishes. Incubate in a tissue culture incubator (37°C, 5% CO2, 95% O2). Read plate at 5 days for “early outgrowth” cells using inverted microscope as described in Subheading 3.7. 4. Incubate for 7 days before changing medium. Gently mix medium with plastic Pasteur pipette and discard 2 ml of medium before adding 2 ml of fresh EPC assay medium.

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5. Read plate at 21 days for “late outgrowth cells” using inverted microscope as described in Subheading 3.7. 6. It will also be necessary to distinguish “late outgrowth” colonies from “early outgrowth” colonies and CFU-HPC via immunohistochemical analysis of antigen expression. To prepare colonies for staining, it is necessary to culture cells in 8 well chamber-slides (glass). Coat the chambers with fibronectin (as above), and plate cells at a density of 1 × 106 cells/chamber. Incubate chamber slides in the same way as dishes. 3.5. MesenCult Assay for Culture of CFU-MSC

1. In a tissue culture hood: add 5 × 105 cells (Blood or BM) to 3 ml complete medium (MesenCult + supplements) and add to 35 mm2 dish (tissue culture treated plastic) to give a cell density of 0.5 × 105 cells/cm2. 2. Incubate for 7 days before changing medium. Gently mix medium with plastic Pasteur pipette and discard 2 ml of medium before adding 2 ml of fresh MesenCult assay medium. 3. Read plate at 21 days for MSC colonies using microscope as described in Subheading 3.8 below. 4. It will also be necessary to distinguish CFU-MSC via immunohistochemical analysis of antigen expression. To prepare colonies for staining, culture cells in 8 well chamber slides (glass). Plate cells at a density of 1 × 106 cells/chamber. Incubate chamber slides in the same way as dishes.

3.6. Determination and Enumeration of CFU-HPC Colonies

MethoCult allows quantification of granulocyte/macrophage colonies (CFU-GM), granulocyte/macrophage/erythroid/megakaryocyte colonies (CFU-GEMM), and erythroid burst forming units (BFU-E). 1. Enumerate dishes for CFU-GM, CFU-GEMM, and BFU-E on top of a grid using an inverted microscope under ×2 or ×5 magnification. The dish is thus scanned by the investigator using the grid for positioning. Erythroid forming progenitors (BFU-E) appear scattered, with no dense colony core under low magnification (×2, ×5, or ×10). They are considered a colony if formed of more than 30 cells (Fig. 1). Under high magnification (×20), the individual cells appear tiny, fused together with an irregular, “bunch of grapes” shape. Granulocyte/macrophage forming progenitors (CFU-GM) appear as multiple cell clusters, with a dense core under low magnification (Fig. 1). The colonies will consist of 30–1,000 s cells. Under high magnification it is possible to observe that CFU-GM consist of two cell types: M, large cells with an oval shape and gray center; and G, round cells that are bright and small. Granulocyte/macrophage/megakaryocyte/erythroid

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Fig. 1. Bright light microscopy images of progenitor cell colonies. (a) BFU-E colony grown in MethoCult (framed within circle). The colony is typically dispersed with no obvious core (×10 magnification). (b) CFU-GEMM colony grown in MethoCult. The colony has a dense core with no obvious border between the core and the periphery (×2 magnification). (c) CFU-GM colony grown in MethoCult. The colony has a dense core appearing as multiple cell clusters, there is a distinct border between the core and the periphery (×2 magnification). (d) CFU-EPC (late outgrowth) colony grown in specialized EPC media (×10 magnification). (e) High power image of late outgrowth EPCs, revealing their “cobblestone” morphology (×40 magnification). (f) High power image of early outgrowth EPCs, contrasting in morphology to the late outgrowth EPCs represented in (d) (×40 magnification). (g) MSC colony grown in MesenCult (×20 magnification). (h) MSCs with short reticulated phenotype (×40 magnification). (i) MSCs as large flattened phenotype (×40 magnification).

forming progenitors (CFU-GEMM) are less numerous than CFU-GM. They contain a highly dense core, and there is an indistinct border between the core and the periphery (Fig. 1). The colonies consist of over 500 cells. Under high magnification, it should be possible to identify erythroid clusters around the colony periphery, as well as G/M. 2. Calculate the total number of colony forming units mobilized from the hind limb according to the number of colonies per plate x by the total number of cells mobilized (prelysis count)/ number of cells originally seeded (5 × 104 cells).

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3. It might be necessary to further characterize the cells growing in MethoCult, to help compare them to cells grown in the CFU-EPC and CFU-MSC assays. Thus, cytospins of cells from HPC colonies need to be prepared. Colonies are easily removed from the MethoCult using a disposable pipette tip and then disrupt into a single cell suspension by flushing with a pipette into PBS. Adjust the volume to 1 × 106 cells/ml. Add 100 μl to Shandon Cytospin funnels and fix onto poly-L-lysine slides at 100 g (1 min). Air-dry slides and fix in ice cold methanol for 10 min, before washing in PBS + 1% BSA. Block slides for 30 min with PBS + 10%BSA. 4. Stain slides with the following primary antibodies for 90 min (all 1/100 dilution in PBS + 1% BSA): goat anti-mouse VEGFR1, rabbit anti-mouse VEGFR2, rat anti-mouse CD34, goat anti-mouse VE-Cadherin, rat anti-mouse CD45, rat antimouse CD115, and rabbit anti-human vWF. The following isotype controls must also be used: Rat IgG1, goat IgG, and rabbit IgG. Then wash slides with PBS and incubate with the following appropriate fluorescently labeled secondary antibodies for 45 min (all 1/250 dilution in PBS + 1% BSA): donkey anti-goat Alexafluor 568 antibody, goat anti-rat Alexafluor 594 antibody, goat anti-rabbit Alexafluor 488 antibody. After a further PBS wash, mount slides onto coverslips using a glycerol based mounting medium containing a 4¢,6-diamidino-2-phenylindole (DAPI) counterstain (Vector Labs). Analyze slides using a fluorescent microscope (Leica, Germany). 5. Cells taken from HPC colonies are positive for CD115, CD34, CD45, VEGFR1 antigens, but are negative for VEGFR2, VE-Cadherin, and vWF antigens. 3.7. Determination and Enumeration of CFU-EPC Colonies

1. EPC medium allows the growth of two types of endothelial progenitor cell type. “Early outgrowth” EPC colonies are detected after 5 days in this medium, and represent an angiogenic precursor cell of hematopoietic origin (6, 7, 18). “Early outgrowth” EPC colonies have spindle shaped cells around the colony perimeter, and the cells within the colony disperse by day 14 (Fig. 1). “Early outgrowth” EPC colonies are easily distinguishable from a second type of EPC colony that grows at later stages (2–3 weeks) in this EPC medium and which are therefore termed “late outgrowth” EPC colonies (CFU-EPC). These are identified by their “endothelial like” cobblestone morphology (Fig. 1). 2. As with CFU-HPC enumeration, place dishes containing CFU-EPC (late outgrowth) on top of a grid using a inverted microscope under ×2 or ×5 magnification and scan by the investigator. Again, calculate the total number of colony forming units mobilized from the hind limb according to the

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number of colonies per plate x by the total number of cells mobilized (prelysis count)/number of cells originally seeded (5 × 105 cells). 3. Further analysis of late outgrowth EPC colonies can be conducted by staining for GS-Lectin and uptake of DIL-labeled Acetylated-low density lipoprotein (Ac-LDL). Add Ac-LDL to EPC medium (5 μg/ml) and incubate cells for 4 h at 37°C. Then wash colonies in PBS and fix with 2% PFA for 10 min before incubation with fluoroscein conjugated griffonia-simplicifolia (GS)-lectin (10 μg/ml in PBS) for 60 min. Analyze colonies using a fluorescent microscope (Leica, Germany). 4. An important in vitro functional characteristic of endothelial cells is their ability to form tubules (angiogenesis) in a defined matrix (20, 21). The ability of EPCs to form tubules is assessed using an EC Matrix Gel (Chemicon). EC Matrix Gel solution and EC Matrix diluent buffer are mixed in a ratio of 9:1 at 4°C. Then add 50 μl to wells of a precooled 96 well plate and incubate at 37°C for 1 h to allow the matrix solution to solidify. Harvest late outgrowth EPC colonies previously grown in dishes (as above) by trypsinization and resuspend in EPC growth medium at a density of 1 × 105 cells/ml. Seed (150 μl) onto the surface of the polymerized matrix. Incubate cells for 24 h. Cellular network structures will be fully developed by 12–18 h. 5. Immunostaining of EPC colonies is necessary to verify that they express specific antigens and can thus be confirmed as early- and late-outgrowth EPCs. Stain chamber slides with EPC colonies with the following primary antibodies for 90 min (all 1/100 dilution in PBS + 1% BSA): goat anti-mouse VEGFR1, rabbit anti-mouse VEGFR2, rat anti-mouse CD34, goat anti-mouse VE-Cadherin, rat anti-mouse CD45, rat antimouse CD115, rabbit anti-human vWF, or goat anti-mouse CD14. The following isotype controls need to be used: Rat IgG1, goat IgG, and rabbit IgG. After 90 min, then wash slides in PBS and incubate with the following appropriate fluorescently labeled secondary antibodies for 45 min (all 1/250 dilution in PBS + 1% BSA): donkey anti-goat Alexafluor 568 antibody, goat anti-rat Alexafluor 594 antibody, goat anti-rabbit Alexafluor 488 antibody. After a further PBS wash, mount slides onto cover slips using a glycerol based mounting medium containing a DAPI counterstain. Analyze slides using a fluorescent microscope (Leica, Germany). 6. CD115, CD14, and CD45 antigen expression are indicative of the “early outgrowth” EPC colonies (6, 18). “Late outgrowth” EPC express CD34, VEGFR2, VE-Cadherin, and von-Willebrand Factor (vWF) (6, 7, 13, 19). Importantly, “late outgrowth” CFU-EPC do not express CD115, CD14 or CD45 and are therefore not of a monocyte–macrophage lineage (13, 22).

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1. MesenCult supports the growth of MSC colonies. Under a microscope these colonies appear as either short reticulated cells, or large flattened cells. Combinations of these two morphologies can appear in the same colony (Fig. 1). 2. As with CFU-HPC enumeration, place dishes containing CFU-MSC on top of a grid using an inverted microscope and observe under a ×2 or ×5 objective lens. Again, calculate the total number of colony forming units mobilized from the hind limb according to the number of colonies per plate x by the total number of cells mobilized (prelysis count)/number of cells originally seeded (5 × 105 cells). 3. It may be necessary to characterize MSC colonies in terms of their tri-lineage differentiation potential. Such assays define the ability of colonies to differentiate into adipogenic, osteogenic, and chondrogenic lineages. Harvest MSC colonies by trypsinization (light cell scraping also necessary) and reseed in flasks with MesenCult. When confluent, subject cell monolayers to the following differentiation assays: Osteogenic differentiation is induced by adding 50 μg/ml ascorbic acid-2-phosphate, 10 nM dexamethasone and 10 mM β-glycerol to DMEM +10% FBS (including Pen/Strep and fungizone) (23). Adipogenic differentiation is induced by adding 50 μg/ml indomethacine, 100nM dexamethasone and 10 ng/ml insulin to DMEM +10% FBS (including Pen/Strep and fungizone) (24). Replace the differentiation medium three times a week for a period of 20 days. To promote chondrogenic differentiation a micromass culture technique is used (9, 23, 25). Detach MSCs by trypsinization and pellet in a conical polypropylene tube. Chondrogenic differentiation medium consists of DMEM supplemented with 10 ng/ ml TGF-β, 50 nM ascorbic acid-2-phosphate, 6.25 μg/ml insulin, and 10% FBS (25). Replace the chondrogenic differentiation medium three times a week for a period of 2–3 weeks. 4. To confirm differentiation of MSCs to osteoblasts, adipocytes, and chondrocytes, stain colonies with cell type specific dyes as reported (9, 24). Stain SPCs cultured in osteogenic medium with Alizarin red. This dye is used to demonstrate the presence of calcium. Interaction of Alizarin dye with calcium ions results in a bright red stain. Stain MSCs cultured in adipogenic medium with Oil red O which is a fat-soluble dye that stains triglycerides and lipids of fixed cells with a deep red color. Stain MSCs cultured in chondroinductive medium using Toluidine Blue, which stains the background blue (orthochromatic staining) and the areas of cartilage matrix red-purple (metachromatic staining). 5. It is necessary to further characterize the colonies grown in MesenCult medium to verify that that they express antigens consistent with an MSC phenotype. Stain chamber slides with MSC

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colonies with the following antibodies for 90 min (all 1/100 dilution in PBS + 1% BSA): goat anti-mouse VEGFR1, rabbit anti-mouse VEGFR2, rat anti-mouse CD34, goat anti-mouse VE-Cadherin, rat anti-mouse CD45, rat anti-mouse CD29, rat anti-mouse CD105, rabbit anti-human vWF. The following isotype controls need to be used: Rat IgG1, goat IgG, and rabbit IgG. After 90 min, wash slides in PBS and incubate with the following appropriate fluorescently labeled secondary antibodies for 45 min (all 1/250 dilution in PBS + 1% BSA): donkey anti-goat Alexafluor 568 antibody, goat anti-rat Alexafluor 594 antibody, goat anti-rabbit Alexafluor 488 antibody. After a further PBS wash, mount slides onto coverslips using glycerol based mounting medium containing a DAPI counterstain. Analyze slides using a fluorescent microscope (Leica, Germany). 6. Plastic adherent bone marrow derived MSCs express CD29, CD105, VEGFR2 antigens and are negative for CD45, CD34, vWF, and VE-cadherin (13) as demonstrated in previous studies on stromal progenitor cells (9, 10).

4. Notes 1. Fibronectin has a tendency to stick to pipette tips. A proportion of the powder will not dissolve but this can be ignored (manufacturer’s instructions). 2. Weighing the mouse is essential in order not to overdose. Generally, a 20–22 g mouse requires 170 μl, whereas a 23–26 g mouse requires 190 μl of anesthetic (i.p.) to initially lose consciousness. Then administer an extra 50 μl urethane (i.p.) to keep the mouse under anesthesia. However, there will be batch variation. 3. It is necessary to stretch the cannula in order to further reduce the diameter so that they can be inserted more easily into the femoral vein and artery. 4. Critically, when buffer alone is perfused through the bone marrow, very few progenitors and leukocytes are mobilized from the bone marrow. 5. Pellet is not stuck hard to bottom of tube due to red blood cell contamination. 6. This reading is necessary for when the researcher calculates the total number of progenitor cells mobilized, as described in later sections. 7. Remaining red blood cells will not interfere with the assay, however, and can be ignored.

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References 1. Cashen AF, Link D, Devine S, DiPersio J (2004) Cytokines and stem cell mobilization for autologous and allogeneic transplantation. Curr Hematol Rep 3:406–412 2. Broxmeyer HE, Orschell CM, Clapp DW, Hangoc G, Cooper S, Plett PA et al (2005) Rapid mobilization of murine and human hematopoietic stem and progenitor cells with AMD3100, a CXCR4 antagonist. J Exp Med 201:1307–1318 3. Martin C, Bridger GJ, Rankin SM (2006) Structural analogues of AMD3100 mobilise haematopoietic progenitor cells from bone marrow in vivo according to their ability to inhibit CXCL12 binding to CXCR4 in vitro. Br J Haematol 134:326–329 4. Calandra G, McCarty J, McGuirk J, Tricot G, Crocker SA, Badel K, Grove B, Dye A, Bridger G (2008) AMD3100 plus G-CSF can successfully mobilize CD34+ cells from non-Hodgkin’s lymphoma. Hodgkin’s disease and multiple myeloma patients previously failing mobilization with chemotherapy and/or cytokine treatment: compassionate use data. Bone Marrow Transplant 41:331–338 5. Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T, Witzenbichler B, Schatteman G, Isner JM (1997) Isolation of putatative progenitor endothelial cells for angiogenesis. Science 275:964–967 6. Nolan DJ, Ciarrocchi A, Mellick AS, Jaggi JS, Bambino K, Gupta S et al (2007) Bone marrow-derived endothelial progenitor cells are a major determinant of nascent tumour neovascularization. Genes Dev 21:1546–1558 7. Yoder MC, Mead LE, Prater D, Krier TR, Mroueh KN, Li F et al (2007) Redefining endothelial progenitor cells via clonal analysis and hematopoietic stem/progenitor cell principals. Blood 109:1801–1809 8. Madlambayan GJ, Butler JM, Hosaka K, Jorgensen M, Fu D, Guthrie SM et al (2009) Bone marrow stem and progenitor cell contribution to neovasculogenesis is dependent on model system with SDF-1 as a permissive trigger. Blood 114:4310–4319 9. Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R, Mosca JD et al (1999) Multilineage potential of adult human mesenchymal stem cells. Science 284:143–147 10. Phinney DG, Kopen G, Isaacson RL, Prockop DJ (1999) Plastic adherent stromal cells from the bone marrow of commonly used strains of inbred mice: variations in yield, growth, and differentiation. J Cell Biochem 72:570–585

11. Wexler SA, Donaldson C, Denning-Kendall P, Rice C, Bradley B, Hows JM (2003) Adult bone marrow is a rich source of human mesenchymal ‘stem’ cells but umbilical cord and mobilized adult blood are not. Br J Haematol 121:368–374 12. Roufosse CA, Direkze NC, Otto WR, Wright NA (2004) Circulating mesenchymal stem cells. Int J Biochem Cell Biol 36:585–597 13. Pitchford SC, Furze RC, Jones CP, Wengner AM, Rankin SM (2009) Differential mobilization of subsets of progenitor cells from the bone marrow. Cell Stem Cell 4:1–11 14. Pitchford SC, Hahnel MJ, Jones CP, Rankin SM (2010) Troubleshooting: quantification of mobilization of progenitor cell subsets from bone marrow in vivo. J Pharmacol Toxicol Methods 61:113–121 15. Palframan RT, Collins PD, Williams TJ, Rankin SM (1998) Eotaxin induces a rapid release of eosinophils and their progenitors from the bone marrow. Blood 91:2240–2248 16. Martin C, Burdon PC, Bridger G, GutierrezRamos JC, Williams TJ, Rankin SM (2003) Chemokines acting via CXCR2 and CXCR4 control the release of neutrophils from the bone marrow and their return following senescence. Immunity 19:583–593 17. Burdon PC, Martin C, Rankin SM (2008) Migration across the sinusoidal endothelium regulates neutrophil mobilization in response to ELR + CXC chemokines. Br J Haematol 142:100–108 18. Jones CP, Pitchford SC, Lloyd CM, Rankin SM (2009) CXCR2 mediumtes the recruitment of endothelial progenitor cells during allergic airways remodelling. Stem Cells 27:3074–3081 19. Hur J, Yoon C-H, Kim H-S, Choi J-H, Kang H-J, Hwang K-K et al (2003) Characterization of two types of endothelial progenitor cells and their different contributions to neovasculogenesis. Arterioscler Thromb Vasc Biol 24:288–293 20. Madri JA, Pratt BM (1986) Endothelial cellmatrix interactions: in vitro models of angiogenesis. J Histochem Cytochem 34:85–91 21. Salani D, Taraboletti G, Rosano L, Di Castro V, Borsotti P, Giavazzi R, Bagnato A (2000) Endothelin-1 induces an angiogenic phenotype in cultured endothelial cells and stimulates neovascularization in vivo. Am J Pathol 157: 1703–1711 22. Hirschi KK, Ingram DA, Yoder MC (2008) Assessing identity, phenotype, and fate of endothelial progenitor cells. Arterioscler Thromb Vasc Biol 28:1584–1695

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23. Eslaminejad MB, Nikmahzar A, Taghiyar L, Nadri S, Massumi M (2006) Murine mesenchymal stem cells isolated by low density primary culture system. Dev Growth Differ 48:361–370 24. Zuk PA, Zhu M, Ashjian P, De Ugarte DA, Huang JI, Mizuno H, Alfonso ZC, Fraser JK, Benhaim P, Hedrick MH (2002) Human adi-

pose tissue is a source of multipotent stem cells. Mol Biol Cell 13:4279–4295 25. Pevsner-Fischer M, Morad V, Cohen-Sfady M, Rousso-Noori L, Zanin-Zhorov A, Cohen S, Cohen IR, Zipori D (2007) Toll-like receptors and their ligands control mesenchymal stem cell functions. Blood 109:1422–1432

Chapter 13 Vascular Progenitor Cell Mobilization Kirsten A. Kienstra and Karen K. Hirschi Abstract Blood vessel formation plays a key role in both physiologic and pathologic tissue growth and healing. Thus, a thorough understanding of the mechanisms underlying neovascularization will translate into innovative clinical treatment strategies for a wide variety of disease processes. Vascular precursor/progenitor cell populations have been isolated from several different tissue types and have a rich potential for use in vascular regenerative strategies. Furthermore, levels of circulating endothelial progenitor cells (EPC) have been shown to correlate with outcomes in cardiovascular and vascular diseases. Treatment with EPC has been shown to improve functional outcomes following cardiac and peripheral vascular ischemia. Recent studies have also demonstrated a role for EPC in pediatric disease processes such as retinopathy of prematurity and bronchopulmonary dysplasia. In addition, many of the drugs utilized to treat vascular disease impact EPC mobilization and function. Importantly, the type of vascular injury appears to dictate the mechanism of neovascularization, highlighting the importance of carefully selected vascular regenerative strategies. Key words: Endothelial progenitor cell, Neovascularization, Umbilical cord blood, Mobilization, Neonatal

1. Introduction The vasculature is a ubiquitous organ system, and blood vessel formation plays an essential role in many physiological processes, including tissue growth as occurs during menstruation or placental development, and tissue healing such as vessel collateralization following ischemia. Blood vessel formation is also critical in pathological processes, such as tumorigenesis or development of retinopathy. Blood vessels comprise a luminal layer of endothelial cells with an underlying basement membrane, and a surrounding vessel wall

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_13, © Springer Science+Business Media, LLC 2012

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of mural cells (pericytes or vascular smooth muscle cells). Mechanisms of blood vessel formation include vasculogenesis (1, 2), the process by which vessels are formed de novo from undifferentiated cell types (i.e., multi-lineage progenitors and committed precursors), and angiogenesis (3), the expansion and remodeling of existing vascular networks. Traditionally, vasculogenesis was thought to occur during embryonic development while angiogenesis occurred postnatally. However, accumulating evidence suggests a role for adult precursor/progenitor cells in postnatal neovascularization (4), and has shifted the paradigm to recognize that both mechanisms likely occur during embryonic and postnatal blood vessel formation, maintenance, and repair. A thorough understanding of the mechanisms underlying adult neovascularization will ultimately translate to innovative clinical treatment strategies to enhance tissue growth and healing, or target pathological disease processes. We must also understand the identity and source of precursor/progenitor cells that play a key role in neovascularization. During embryonic blood vessel formation, endothelial cell differentiation and tube formation are followed by, and require, the recruitment and differentiation of mural cells (5). Although both endothelial and mural cell precursors/progenitors have been identified in the adult and may play parallel roles in postnatal neovascularization (6), our discussions focus specifically on endothelial precursor/progenitor cells.

2. Endothelial Progenitor Cell Characteristics Since the sentinel publication by Asahara in 1997 describing the role of circulating endothelial progenitor cells (EPC) in postnatal neovascularization (4), the study of EPC has been complicated by complexities in identification, and functional and phenotypic overlap with hematopoietic cells. Unfortunately, to date, no cell surface marker has been found to be specific for EPC. In terms of cell surface phenotype, the markers most commonly used to identify and isolate EPC are combinations of CD34+, VEGFR2+ (Flk-1, KDR), and CD133+ (AC133). Markers used in each study referenced in this review will be denoted. Numerous studies have shown that these populations of cells play a role in postnatal neovascularization and are predictive biomarkers for various disease states (7–10). While other combinations of phenotypic markers have been utilized to isolate cell populations involved in postnatal neovascularization (as reviewed in refs. 11–13), a phenotypic profile specific to EPC has yet to be identified. In addition to cell surface markers, EPC have been identified using a variety of in vitro adhesion and growth characteristics (4, 14–17). As with cell surface markers, vascular and

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hematopoietic lineages have significant phenotypic overlap and may display similar features in culture. While some populations may be true endothelial progenitors, others may comprise macrophage–monocyte populations that promote neoangiogenesis via mechanisms such as paracrine signaling pathways (18). Thus, it is important to demonstrate that purported EPC populations have bona fide endothelial cell potential using specific tests such as in vitro tube formation (important to demonstrate the presence of a lumen), and de novo vessel formation in vivo documented using high resolution confocal imaging strategies (18).

3. Sources of Vascular Progenitor Cells In addition to sharing many overlapping functional and phenotypic characteristics in the adult, the hematopoietic and vascular systems also share developmental features. During embryogenesis, blood and blood vessels develop in parallel, with hematopoietic stem/progenitors arising from hemogenic endothelium in the yolk sac (19), the embryo (20), and the placenta (21). The relationship between vascular and hematopoietic lineages during postnatal life remains to be clearly delineated, and the ability of adult hematopoietic stem/progenitor cells to generate vascular progenitors and endothelial cells remains controversial (22–25). Nonetheless, given the close developmental relationship between the vascular and hematopoietic systems, it is not surprising that numerous studies have localized vascular precursor/progenitor cells to the peripheral circulation and bone marrow. 3.1. Peripheral and Umbilical Cord Blood

Since the discovery that human CD34+ peripheral blood mononuclear cells have the capacity to become endothelial-like cells in vitro and contribute to ischemia-induced neovascularization in vivo (4), countless studies have aimed to further delineate the specific subpopulation(s) of peripheral blood with vascular potential. Using a heterogeneous combination of markers and culture strategies to isolate putative EPC, investigators have linked peripheral bloodderived EPC to many disease processes, ranging from colonization of vascular prostheses (10), to recovery following hindlimb (4, 26) and coronary ischemia (27). EPC have been isolated from umbilical cord blood using similar protocols as developed for peripheral blood, and studies have shown parallel endothelial cell potential (using subsets of CD34+ cells or endothelial colony-forming cells (ECFC)) (10, 15, 28). Interestingly, compared to matched adult peripheral blood EPC, umbilical cord blood contains a greater proportion of EPC with a higher proliferative capacity (15). Furthermore, the numbers of

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EPC (ECFC) in human cord blood increase with advancing gestational age: while constant from 24 to 31 weeks gestation, EPC levels double by 32–36 weeks, and triple by 37–40 weeks (29). The role for such variation and the mechanisms underlying the change remain unclear. Oxidative stress reduced the proliferative potential of EPC (ECFC) from both peripheral blood and umbilical cord blood, with the multi-potent progenitors most affected (30). Surprisingly, adult EPC were more sensitive to the oxidative stress then umbilical cord blood EPC (30). Based on enriched content and proliferative capacity of umbilical cord blood compared to adult peripheral blood, it warrants continued investigation as a source for vascular progenitor cells. 3.2. Bone Marrow

As the origin of most circulating blood cells, the bone marrow compartment is intricately tied to the peripheral circulation. Thus, it is not surprising that the bone marrow has also been described as a source for vascular progenitor cells. The bone marrow contains a heterogeneous population of cells, with a wide variety of stem, progenitor and differentiated cell types. Several different subfractions have been proposed to possess vascular potential. To assess for a bone marrow origin of EPC, many studies have transplanted labeled (Tie-2- or VEGFR2-LacZ) donor bone marrow mononuclear cells into irradiated recipients and demonstrated donorderived LacZ-positive vascular endothelial cells (31–33). In view of the close developmental association between the vascular and hematopoietic systems, hematopoietic stem cells (HSC) have been highly investigated as a potential source for EPC. Several studies utilizing labeled-HSC bone marrow transplantation, including both single HSC and serial transplantation, have shown that HSC can give rise to both blood and vascular endothelial cells (22–24). In contrast, other studies suggest that HSC do not contribute to endothelial cells (25). Thus, the postnatal relationship between HSC and the vasculature remains controversial. Another proposed bone marrow cell source for vascular regeneration is myeloid progenitor cells. Some studies suggest that myeloid progenitors have the capacity to differentiate into vascular endothelial cells (34, 35), while others propose that they are recruited to a perivascular position and act in a paracrine fashion to promote neovascularization (36, 37). Hence, there appears to be a close relationship between myeloid and endothelial cells lineages, but the precise mechanism(s) of action during vascular regeneration remain to be fully characterized.

3.3. Tissue-Resident

In addition to the peripheral blood and bone marrow, other postnatal tissues have been described as a source for vascular progenitor cells. Notably, vascular progenitors may reside within adult blood vessel walls. EPC have been isolated from the umbilical and aortic vessel wall (38), and from a “vasculogenic zone” in

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human adult vessels between the smooth muscle and adventitial layers (39). The precise phenotype and function of vessel-resident vascular progenitors have yet to be fully characterized. In addition, vascular progenitor cells may be part of a cycle of mobilization, circulation, and tissue deposition, versus distinct progenitor populations existing within different tissues (40).

4. Role of Vascular Progenitors in Adult and Pediatric Disease 4.1. EPC as Biomarkers

Although the precise identification and vascular regenerative mechanism of EPC remain to be completely defined, circulating levels of EPC have been shown to correlate with various disease processes. Several large studies have shown that EPC levels (defined as CD34+/VEGFR2+ or CFU-Hill) in the peripheral blood can predict the occurrence of cardiovascular events and may help to identify patients at increased risk for morbidity and mortality (8, 14). Specifically, increased levels of circulating EPC have been associated with a reduced risk of cardiovascular-related death and first major cardiovascular event (8). Similarly, increased levels of circulating EPC (CD34+ subsets) are predictive of improved outcomes following acute ischemic stroke related to cerebrovascular disease (41). Metabolic syndrome has been associated with decreased levels of circulating EPC (CD34+/VEGFR2+) and may be related to a mobilization defect (42, 43). While the establishment of EPC as a biomarker does not delineate cause or effect, it clearly links EPC with many vascular disease processes and justifies the need for ongoing research.

4.2. EPC Levels and Injury/Disease Recovery

Several models of ischemic vascular disease have been used to study EPC mobilization and tissue recovery. Consistent with human studies demonstrating a benefit of EPC therapy for coronary artery disease and functional cardiac recovery after ischemia (44), a mouse model of myocardial infarction showed that animals with increased EPC (defined by culture assay with Sca-1+/VEGR2+ phenotype) had improved clinical outcomes (27). Similarly, human adults with peripheral vascular disease had significant clinical improvement following injection with stem and progenitor cells from bone marrow, peripheral blood, or umbilical cord blood (45). Mouse models of hindlimb ischemia have shown parallel benefits of EPC (defined by culture assay and Flt1+/VEGFR2+/vWF+), with increased EPC levels associated with improved perfusion, capillary density, and overall functional outcome (46–48).

4.3. Pediatric Models of Injury and Disease

Most of the studies using EPC therapy have focused on models of adult disease. However, many pediatric and neonatal diseases would also benefit from such therapeutic strategies, and researchers

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are now beginning to address this understudied population. For neonates, exposure to hyperglycemia and a diabetic intrauterine environment has been shown to reduce the function of EPC (ECFC) isolated from umbilical cord blood, providing possible mechanistic insight into the long-term cardiovascular complications seen in infants of diabetic pregnancies (49). Preterm infants with retinopathy of prematurity (ROP) had significantly increased levels of EPC (CD34+/CD133+/CD144+) compared to preterm infants without ROP, suggesting a role for circulating factors in the development and progression of ROP (50). Another neonatal disease that may benefit from therapy with vascular progenitor cells is bronchopulmonary dysplasia (BPD). With improved survival of extremely preterm infants, an increasing number of children are living with the consequences and complications of BPD. The lungs of infants with BPD have arrested alveolar and vascular development, and abnormalities in the vasculature have been shown to play a critical role in the disease process (51, 52). In a mouse model of BPD, pups exposed to hyperoxia had decreased alveolar capillary density, abnormal lung structure, and decreased numbers of EPC (CD45−/Sca-1+/CD133+/VEGFR2+) in the bone marrow, peripheral blood, and lung tissue (53). In contrast, mice treated with a population of “bone marrow-derived angiogenic cells” (defined by culture-based assay and Tie2+/VEGFR2−/CD45+/ CD34−/Sca-1−) had complete restoration of their lung structure (54), highlighting both the key role played by the vasculature in BPD and the exciting potential for innovative new therapies. 4.4. Drug-Induced Mobilization

As an extension of the established correlation between vascular progenitor cells and various disease states, an expanding list of medications have been shown to impact EPC mobilization and function. Not surprisingly, most of these therapeutic agents are used to treat vascular disease. Angiotensin-II inhibitors were shown to increase EPC number and improve EPC dysfunction in a rat model of hypertension and stroke, thought to be mediated through an anti-oxidative mechanism (55). Statins have been shown to increase EPC mobilization and differentiation in both mouse and rat models via the PI 3-kinase/Akt pathway and upregulation of integrins (56, 57). The phosphodiesterase inhibitor sildenafil increased numbers of EPC in patients with several forms of pulmonary hypertension, in whom reduction in EPC numbers was correlated with multiple clinical parameters of diminished cardiac function (7). Treatment with the hormone estradiol accelerated reendothelialization and increased EPC mobilization and proliferation in a mouse model of carotid artery injury (58). Treatment of EPC with prostaglandin E1 enhanced EPC number and function in vivo, and improved neovascularization capacity and function in a mouse model of hindlimb ischemia, with effects mediated by increased CXCR4 expression

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and dependent upon nitric oxide synthase (59). Interestingly, physical activity has also been shown to increase EPC numbers in a mouse exercise and vascular injury model via a nitric oxide-dependent mechanism, an effect that could at least partially explain the beneficial impact of exercise on cardiovascular health (60).

5. Type of Injury Dictates Mechanism(s) of Neovascularization

When designing research studies and therapeutic modalities, it is critical to understand that different cell populations and mechanisms may be involved in recovery from different types of tissue injury and vessel regeneration. For example, the cells and pathways involved in tumor neovascularization likely differ from those participating in vascular collateralization following tissue ischemia. Using several different liver injury models, our laboratory has shown that the same cell type has variable contribution to endothelial cell regeneration depending on the precise mechanism of injury (61). Such findings highlight the importance of careful selection of appropriate cell populations and delivery mechanism when embarking on vascular regenerative strategies.

Acknowledgments This work was supported by AHA-TX 0865252F (KAK), AAP NRP (KAK), NIH R01 HL77675 (KKH), NIH R01 HL096360 (KKH), NIH R01 EB005173 (KKH), NIH P20 EB007076 (KKH), and USDA ARS—6250-51000 (KKH). References 1. Goldie LC, Nix MK, Hirschi KK (2008) Embryonic vasculogenesis and hematopoietic specification. Organogenesis 4:257–263 2. Risau W, Flamme I (1995) Vasculogenesis. Annu Rev Cell Dev Biol 11:73–91 3. Risau W (1997) Mechanisms of angiogenesis. Nature 386:671–674 4. Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T et al (1997) Isolation of putative progenitor endothelial cells for angiogenesis. Science 275:964–967 5. Lindahl P, Johansson BR, Leveen P, Betsholtz C (1997) Pericyte loss and microaneurysm formation in PDGF-B-deficient mice. Science 277:242–245

6. Corselli M, Chen CW, Crisan M, Lazzari L, Peault B (2010) Perivascular ancestors of adult multipotent stem cells. Arterioscler Thromb Vasc Biol 30:1104–1109 7. Diller GP, van Eijl S, Okonko DO, Howard LS, Ali O, Thum T et al (2008) Circulating endothelial progenitor cells in patients with Eisenmenger syndrome and idiopathic pulmonary arterial hypertension. Circulation 117:3020–3030 8. Werner N, Kosiol S, Schiegl T, Ahlers P, Walenta K, Link A et al (2005) Circulating endothelial progenitor cells and cardiovascular outcomes. N Engl J Med 353:999–1007 9. Rafii S, Lyden D (2003) Therapeutic stem and progenitor cell transplantation for organ

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vascularization and regeneration. Nat Med 9:702–712 10. Shi Q, Rafii S, Wu MH, Wijelath ES, Yu C, Ishida A et al (1998) Evidence for circulating bone marrow-derived endothelial cells. Blood 92:362–367 11. Chao H, Hirschi KK (2010) Hemato-vascular origins of endothelial progenitor cells? Microvasc Res 79:169–173 12. Hirschi KK (2010) Vascular precursors: origin, regulation and function. Arterioscler Thromb Vasc Biol 30:1078–1079 13. Richardson MR, Yoder MC (2011) Endothelial progenitor cells: Quo Vadis? J Mol Cell Cardiol Feb 50(2):266–72 14. Hill JM, Zalos G, Halcox JP, Schenke WH, Waclawiw MA, Quyyumi AA et al (2003) Circulating endothelial progenitor cells, vascular function, and cardiovascular risk. N Engl J Med 348:593–600 15. Ingram DA, Mead LE, Tanaka H, Meade V, Fenoglio A, Mortell K et al (2004) Identification of a novel hierarchy of endothelial progenitor cells using human peripheral and umbilical cord blood. Blood 104:2752–2760 16. Ito H, Rovira II, Bloom ML, Takeda K, Ferrans VJ, Quyyumi AA et al (1999) Endothelial progenitor cells as putative targets for angiostatin. Cancer Res 59:5875–5877 17. Lin Y, Weisdorf DJ, Solovey A, Hebbel RP (2000) Origins of circulating endothelial cells and endothelial outgrowth from blood. J Clin Invest 105:71–77 18. Hirschi KK, Ingram DA, Yoder MC (2008) Assessing identity, phenotype, and fate of endothelial progenitor cells. Arterioscler Thromb Vasc Biol 28:1584–1595 19. Goldie LC, Lucitti JL, Dickinson ME, Hirschi KK (2008) Cell signaling directing the formation and function of hemogenic endothelium during murine embryogenesis. Blood 112:3194–3204 20. Zovein AC, Hofmann JJ, Lynch M, French WJ, Turlo KA, Yang Y et al (2008) Fate tracing reveals the endothelial origin of hematopoietic stem cells. Cell Stem Cell 3:625–636 21. Lee LK, Ueno M, Van Handel B, Mikkola HK (2010) Placenta as a newly identified source of hematopoietic stem cells. Curr Opin Hematol 17:313–318 22. Grant MB, May WS, Caballero S, Brown GA, Guthrie SM, Mames RN et al (2002) Adult hematopoietic stem cells provide functional hemangioblast activity during retinal neovascularization. Nat Med 8:607–612 23. Jackson KA, Majka SM, Wang H, Pocius J, Hartley CJ, Majesky MW et al (2001)

Regeneration of ischemic cardiac muscle and vascular endothelium by adult stem cells. J Clin Invest 107:1395–1402 24. Larrivee B, Niessen K, Pollet I, Corbel SY, Long M, Rossi FM et al (2005) Minimal contribution of marrow-derived endothelial precursors to tumor vasculature. J Immunol 175:2890–2899 25. Purhonen S, Palm J, Rossi D, Kaskenpaa N, Rajantie I, Yla-Herttuala S et al (2008) Bone marrow-derived circulating endothelial precursors do not contribute to vascular endothelium and are not needed for tumor growth. Proc Natl Acad Sci USA 105:6620–6625 26. Harraz M, Jiao C, Hanlon HD, Hartley RS, Schatteman GC (2001) CD34− blood-derived human endothelial cell progenitors. Stem Cells 19:304–312 27. Jujo K, Hamada H, Iwakura A, Thorne T, Sekiguchi H, Clarke T et al (2010) CXCR4 blockade augments bone marrow progenitor cell recruitment to the neovasculature and reduces mortality after myocardial infarction. Proc Natl Acad Sci USA 107:11008–11013 28. Timmermans F, Van Hauwermeiren F, De Smedt M, Raedt R, Plasschaert F, De Buyzere ML et al (2007) Endothelial outgrowth cells are not derived from CD133+ cells or CD45+ hematopoietic precursors. Arterioscler Thromb Vasc Biol 27:1572–1579 29. Javed MJ, Mead LE, Prater D, Bessler WK, Foster D, Case J et al (2008) Endothelial colony forming cells and mesenchymal stem cells are enriched at different gestational ages in human umbilical cord blood. Pediatr Res 64:68–73 30. Case J, Ingram DA, Haneline LS (2008) Oxidative stress impairs endothelial progenitor cell function. Antioxid Redox signal 10: 1895–1907 31. Asahara T, Masuda H, Takahashi T, Kalka C, Pastore C, Silver M et al (1999) Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res 85:221–228 32. Llevadot J, Murasawa S, Kureishi Y, Uchida S, Masuda H, Kawamoto A et al (2001) HMGCoA reductase inhibitor mobilizes bone marrow-derived endothelial progenitor cells. J Clin Invest 108:399–405 33. Murayama T, Tepper OM, Silver M, Ma H, Losordo DW, Isner JM et al (2002) Determination of bone marrow-derived endothelial progenitor cell significance in angiogenic growth factor-induced neovascularization in vivo. Exp Hematol 30:967–972 34. Bailey AS, Willenbring H, Jiang S, Anderson DA, Schroeder DA, Wong MH et al (2006)

13 Myeloid lineage progenitors give rise to vascular endothelium. Proc Natl Acad Sci USA 103:13156–13161 35. Fernandez Pujol B, Lucibello FC, Gehling UM, Lindemann K, Weidner N, Zuzarte ML et al (2000) Endothelial-like cells derived from human CD14 positive monocytes. Differentiation 65: 287–300 36. Grunewald M, Avraham I, Dor Y, BacharLustig E, Itin A, Jung S et al (2006) VEGFinduced adult neovascularization: recruitment, retention, and role of accessory cells. Cell 124:175–189 37. Rehman J, Li J, Orschell CM, March KL (2003) Peripheral blood “endothelial progenitor cells” are derived from monocyte/macrophages and secrete angiogenic growth factors. Circulation 107:1164–1169 38. Ingram DA, Mead LE, Moore DB, Woodard W, Fenoglio A, Yoder MC (2005) Vessel wallderived endothelial cells rapidly proliferate because they contain a complete hierarchy of endothelial progenitor cells. Blood 105: 2783–2786 39. Zengin E, Chalajour F, Gehling UM, Ito WD, Treede H, Lauke H et al (2006) Vascular wall resident progenitor cells: a source for postnatal vasculogenesis. Development 133:1543–1551 40. Majka SM, Jackson KA, Kienstra KA, Majesky MW, Goodell MA, Hirschi KK (2003) Distinct progenitor populations in skeletal muscle are bone marrow derived and exhibit different cell fates during vascular regeneration. J Clin Invest 111:71–79 41. Yip HK, Chang LT, Chang WN, Lu CH, Liou CW, Lan MY et al (2008) Level and value of circulating endothelial progenitor cells in patients after acute ischemic stroke. Stroke 39:69–74 42. Jialal I, Devaraj S, Singh U, Huet BA (2010) Decreased number and impaired functionality of endothelial progenitor cells in subjects with metabolic syndrome: implications for increased cardiovascular risk. Atherosclerosis 211:297–302 43. Jialal I, Fadini GP, Pollock K, Devaraj S (2010) Circulating levels of endothelial progenitor cell mobilizing factors in the metabolic syndrome. Am J Cardiol Dec 1;106(11):1606–8 44. Dimmeler S, Zeiher AM (2009) Cell therapy of acute myocardial infarction: open questions. Cardiology 113:155–160 45. Burt RK, Loh Y, Pearce W, Beohar N, Barr WG, Craig R et al (2008) Clinical applications of blood-derived and marrow-derived stem cells for nonmalignant diseases. JAMA 299:925–936 46. Hu Z, Zhang F, Yang Z, Yang N, Zhang D, Zhang J et al (2008) Combination of simvasta-

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tin administration and EPC transplantation enhances angiogenesis and protects against apoptosis for hindlimb ischemia. J Biomed Sci 15:509–517 47. Oh IY, Yoon CH, Hur J, Kim JH, Kim TY, Lee CS et al (2007) Involvement of E-selectin in recruitment of endothelial progenitor cells and angiogenesis in ischemic muscle. Blood 110:3891–3899 48. Ruifrok WP, de Boer RA, Iwakura A, Silver M, Kusano K, Tio RA et al (2009) Estradiolinduced, endothelial progenitor cell-mediated neovascularization in male mice with hind-limb ischemia. Vasc Med 14:29–36 49. Ingram DA, Lien IZ, Mead LE, Estes M, Prater DN, Derr-Yellin E et al (2008) In vitro hyperglycemia or a diabetic intrauterine environment reduces neonatal endothelial colony-forming cell numbers and function. Diabetes 57:724–731 50. Machalinska A, Modrzejewska M, Kotowski M, Dziedziejko V, Kucia M, Kawa M et al (2010) Circulating stem cell populations in preterm infants: implications for the development of retinopathy of prematurity. Arch Ophthalmol 128:1311–1319 51. Jobe AJ (1999) The new BPD: an arrest of lung development. Pediatr Res 46:641–643 52. Thebaud B, Ladha F, Michelakis ED, Sawicka M, Thurston G, Eaton F et al (2005) Vascular endothelial growth factor gene therapy increases survival, promotes lung angiogenesis, and prevents alveolar damage in hyperoxiainduced lung injury: evidence that angiogenesis participates in alveolarization. Circulation 112:2477–2486 53. Balasubramaniam V, Mervis CF, Maxey AM, Markham NE, Abman SH (2007) Hyperoxia reduces bone marrow, circulating, and lung endothelial progenitor cells in the developing lung: implications for the pathogenesis of bronchopulmonary dysplasia. Am J Physiol Lung Cell Mol Physiol 292:L1073–L1084 54. Balasubramaniam V, Ryan SL, Seedorf GJ, Roth EV, Heumann TR, Yoder MC et al (2010) Bone marrow-derived angiogenic cells restore lung alveolar and vascular structure after neonatal hyperoxia in infant mice. Am J Physiol Lung Cell Mol Physiol 298: L315–L323 55. Yu Y, Fukuda N, Yao EH, Matsumoto T, Kobayashi N, Suzuki R et al (2008) Effects of an ARB on endothelial progenitor cell function and cardiovascular oxidation in hypertension. Am J Hypertens 21:72–77 56. Dimmeler S, Aicher A, Vasa M, Mildner-Rihm C, Adler K, Tiemann M et al (2001) HMGCoA reductase inhibitors (statins) increase

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endothelial progenitor cells via the PI 3-kinase/ Akt pathway. J Clin Invest 108:391–397 57. Walter DH, Rittig K, Bahlmann FH, Kirchmair R, Silver M, Murayama T et al (2002) Statin therapy accelerates reendothelialization: a novel effect involving mobilization and incorporation of bone marrow-derived endothelial progenitor cells. Circulation 105:3017–3024 58. Iwakura A, Luedemann C, Shastry S, Hanley A, Kearney M, Aikawa R et al (2003) Estrogenmediated, endothelial nitric oxide synthasedependent mobilization of bone marrow-derived endothelial progenitor cells contributes to reendothelialization after arterial injury. Circulation 108:3115–3121

59. Herrler T, Leicht SF, Huber S, Hermann PC, Schwarz TM, Kopp R et al (2009) Prostaglandin E positively modulates endothelial progenitor cell homeostasis: an advanced treatment modality for autologous cell therapy. J Vasc Res 46:333–346 60. Laufs U, Werner N, Link A, Endres M, Wassmann S, Jurgens K et al (2004) Physical training increases endothelial progenitor cells, inhibits neointima formation, and enhances angiogenesis. Circulation 109:220–226 61. Kienstra KA, Jackson KA, Hirschi KK (2008) Injury mechanism dictates contribution of bone marrow-derived cells to murine hepatic vascular regeneration. Pediatr Res 63:131–136

Chapter 14 Evaluation of Circulating Endothelial Precursor Cells in Cancer Patients Francesco Bertolini, Patrizia Mancuso, Liat Benayoun, Svetlana Gingis-Velitski, and Yuval Shaked Abstract Results obtained from preclinical studies have shown that endothelial progenitor cells (EPCs) play a crucial role in tumor growth and metastasis. In the clinic, EPCs are present in the peripheral blood of cancer patients in higher numbers than in healthy subjects. These cells are mobilized from the bone marrow compartment to the periphery in response to certain cytokines and growth factors. Growing body of evidence suggests that following acute cytotoxic drug therapy levels of circulating EPCs (CEPs) can change significantly in both mouse and human. These changes may predict the efficacy of some anticancer drug treatments. Therefore, the validation and standardization of a procedure to detect CEPs and monitor their kinetic is an important step towards the use of such cells as a possible biomarker to predict clinical outcome. In this chapter, we describe a flow cytometry technique to detect CEPs obtained from human blood specimens stored in both fresh and frozen conditions. Key words: Bone marrow-derived proangiogenic cells, Cellular biomarker, Chemotherapy, Angiogenesis, Antiangiogenic therapy

1. Introduction Over the last decade, a number of different types of bone marrow-derived cells (BMDCs) have emerged as important regulators of tumor angiogenesis, growth, and metastasis (1). Many of these cell types originate from the hematopoietic lineage (for review, see ref. 2). They found to contribute to tumor angiogenesis by different mechanisms such as paracrine secretion of different growth factors, and by contributing to the stability of the vasculature in different ways. In addition to hematopoietic BMDCs, a

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_14, © Springer Science+Business Media, LLC 2012

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subset of non-hematopoietic bone marrow-derived endothelial progenitor cells (EPCs), have been thought to promote tumor angiogenesis by acting as an alternative source of endothelial cells (3). In contrast to all other proangiogenic BMDC types that reside at the perivascular site, EPCs are thought to merge with the wall of a growing blood vessel, where they differentiate into mature endothelial cells, and thus can contribute to vessel growth. However, the relative contribution of EPCs to tumor vessel generation was found to be extremely variable in different preclinical models of cancer (4, 5). Benezra’s group demonstrated that the recruitment of EPCs into tumor vasculature is dependent on the tumor grade (6). They showed that EPCs are key contributors to the first steps of tumor vascularization in small tumors. However, following the establishment of cancer vessels, their relative contribution to neoplastic angiogenesis is quantitatively less than in small tumors, and thus these cells become progressively diluted with the division of differentiated endothelial cells (7). Preclinical observations in lymphoma-bearing NOD/SCID mice treated with intensive 6 day cycles of maximum tolerable dose (MTD) cyclophosphamide, separated by 2 week drug-free breaks, exhibited an initial decline in circulating EPCs (CEPs) followed by substantial increases in their viability and mobilization (8). Such a mobilization effect in CEP levels may contribute to and facilitate tumor cell repopulation during the subsequent drug-free break period that is necessary to allow recovery from the toxic side effects of the drug (8). In another study, we demonstrated that only certain chemotherapy drugs can contribute to the acute mobilization and subsequent tumor homing of CEPs. For example, we found that MTD paclitaxel therapy induces an acute mobilization in CEP levels whereas MTD gemcitabine treatment has no effect on such cells (9). Similar results were obtained in cancer patients treated with paclitaxel-based and gemcitabine-based therapies (9). Furthermore, in circumstances where an antiangiogenic drug known to inhibit CEP mobilization by neutralizing VEGF pathways (10), was combined with chemotherapy-induced acute CEP mobilization, we observed a significant reduction in CEP levels, and enhanced chemotherapy treatment efficacy (9, 11). These results could explain how antiangiogenic drugs act as chemosensitizing agents, and that the efficacy of this combination therapy may be dependent, at least in part, on the type of chemotherapy drug used. These results also suggest that the evaluation of changes in CEP levels during the treatment of chemotherapy and antiangiogenic drugs may serve as a biomarker to predict treatment outcome (12). The development of monoclonal antibodies (Mo-Abs) to detect endothelial cell-associated antigens has led to the growing use of various cytometry methodologies to evaluate CEPs. For example, the multiparametric flow cytometry device is currently

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used for the analysis of CEPs in both mouse and human. CEPs are commonly defined as negative to the pan-hematopoietic marker, CD45, and positive to progenitor markers such as CD34 and CD133. In addition, VEGFR2 and CD146 are usually used as endothelial cell associated markers (for review, see ref. 2). In this chapter, we present protocols the laboratory of Dr. Bertolini has developed to evaluate CEPs in peripheral blood of patients following different types of therapies (13). We should stress at the outset that the lack of standardized method to detect CEPs and mature endothelial cells circulating in the blood which are termed circulating endothelial cells (CECs) has resulted in different values of the number of cells reported using flow cytometry techniques. This has caused increasing confusion about the nature of these cells, both clinically and preclinically. Currently, growing efforts have been made to try and standardize the methodology for detecting CEPs and CECs in peripheral blood, especially for clinical use.

2. Materials 2.1. Blood Specimens

Collected into designated tubes as discussed below for fresh and frozen sample procedures. Usually, the first tube should be discarded in order to avoid increased number of endothelial cells due to venipuncture procedure.

2.2. Antibodies and Reagents

Mo-Abs directly labeled with fluorochromes are purchased from different companies (Table 1). The Mo-Abs were used according to the manufacturer’s instructions. The combination of Mo-Abs used to evaluate CEPs is summarized in Table 2.

2.3. Solutions

1. Red blood cell lysis solution: Dissolve 1.6520 g NH4Cl, 0.2 g KHCO3, and 0.0074 g EDTA (tetra) in 200 ml of distilled water. Keep at room temperature. This reagent must be prepared daily, because CO2 dissolved in water can react with NH4Cl to form the carbonate (NH4)2CO3 that is ineffective in erythrocytes lysis. 2. Flow cytometry buffer solution: Dissolve 1 tablet PBS, 0.6 g of EDTA, and 0.4 g of BSA in 200 ml of distilled water. Keep at room temperature for up to 1 week. 3. TURK solution for white blood cell (WBC) count: Dissolve 0.01 g of crystal violet and 1 ml of 100% acetic acid in 100 ml of distilled water. Keep at room temperature. 4. 7-Aminoactinomycin D (7AAD) solution: Dissolve 1 mg of 7AAD and 50 μl of 100% acetone in 950 μl of PBS. Keep in the dark frozen in aliquots (−20°C).

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Table 1 List of Mo-Abs: The Mo-Abs can be purchased from the indicated supplier: BD—BD Pharmingen, R&D—R&D systems, MB—Myltenji Biotech, BC—Beckman Coulter, Ext—excitation, Ems—emission List of antibodies MoAb

Fluorochrome

Supplier

Cat. no.

Dilution

Volume in tube

Syto 16

FITC

Invitrogen

57578

1:1,000

100 μl/I (for 106 cells)

IgG1

PE

BD

555749

Neat

10 μl

IgG1a

APC

BD

82108

Neat

5 μl

IgG1a

PeCy7

BD

557872

Neat

5 μl

Vegf R2

PE

R&D

FAB357P

Neat

10 μl

CD34

PeCy7

BC

A2169

Neat

5 μl

7-AAD

Ext: 543-555, Ems: SIGMA 655–665

A9400

Neat

10 μg/ml

CD133

APC

MB

130-090-826

Neat

5 μl

CD45

APC-H7

BD

641399

Neat

7.5 μl

Table 2 Mo-Abs master-mix: Master-mix should be prepared for all samples acquired at the same time Antibody master-mix FITC

PE

PerCP

PeCy7

APC

APC-H7

Tube 1

Syto 16

IgG1

7-AAD

IgG1a

IgG1a

CD45

Tube 2

Syto 16

Vegf R2

7-AAD

CD34

CD133

CD45

Slight differences in antibody concentration may change CEP counts. Master-mix should be kept in the dark, and should not exceed 1 week when stored at 4°C

5. Freezing medium: Dissolve 20% DMSO in medium RPMI 1640 to final DMSO concentration of 10%. 2.4. Equipment, Consumable, and Software

1. 5 ml polystyrene round bottom tubes. 2. FACS-Canto flow cytometer (BD) device (or equivalent) which can gather information of eight different parameters with the relevant lasers and fluorescent filters as designated in Table 1. 3. Diva software (or equivalent) for sample acquisition and analysis.

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3. Methods CEPs are evaluated by six-color flow cytometry using nuclear staining Syto16 and 7AAD and a panel of Mo-Abs (see Note 1). Both fresh and frozen sample conditions are discussed (see Note 2). 3.1. Sample Preparation 3.1.1. Whole Blood Fresh Sample Collection

1. Collect 4 ml blood in EDTA-containing tube to avoid coagulation (see Note 3). 2. Process fresh samples—do not exceed 24 h from collection. 3. Keep samples at room temperature until processed. 4. Perform a WBC count using TURK solution. 5. Incubate samples with Mo-Abs according to Table 2.

3.1.2. Whole Blood Frozen Sample Collection (see Notes 4 and 5)

1. Collect blood in cell processing tubes containing sodium citrate and Ficoll. 2. Process samples within 24 h. 3. Centrifuge tubes at room temperature for 25 min at 1,600 × g. 4. Transfer the mononuclear cells into a cryotube. 5. Add an equal volume of freezing medium to cell suspension. 6. Freeze cryotubes in isopropanol cryobox at −80°C freezer for 24 h. 7. Transfer cryotubes to a cryobox, and store in liquid nitrogen until acquired. 8. To thaw frozen cells, place cryotubes immediately in a 37°C bath. 9. Transfer samples from cryotubes into 10 ml tubes. 10. Add 5 ml of flow cytometry buffer solution. 11. Centrifuge for 5 min at 470 × g and subsequently discard supernatant. 12. Add 500 μl of flow cytometry buffer solution to cell pellet. 13. Perform a WBC count using TURK solution. 14. Incubate samples with Mo-Abs according to Table 2 (see Note 6).

3.2. Immunostaining and Lysis of Blood Samples

1. Perform a WBC using TURK solution and prepare 1.5 × 106 total cells for staining. 2. Add antibody mixture (according to Table 2) to 1.5 × 106 total cells (see Note 7). 3. Mix well (vortex) and incubate at 4°C for 20 min in the dark. 4. Vortex tubes briefly and add 4 ml of 1× RBC lysis buffer.

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5. Incubate at 4°C for 10 min. 6. Centrifuge for 5 min at 300–500 × g at 4°C. 7. Discard the resulting supernatant. 8. Resuspend the pellet with 350 μl of flow cytometry buffer and vortex. 9. Samples are now ready for acquisition by flow cytometer (see Note 8). 3.3. Sample Acquisition and Data Analysis

1. Acquire at least 1 × 106 cells per blood sample (see Note 9). 2. Perform post acquisition compensation (see Note 7). 3. Set a gate on FSC/SSC parameter to exclude cellular fragments (debris) from the analysis. 4. Draw a dot plot of SSC vs. CD45 and set a gate on CD45− population to exclude WBC from the analysis. 5. Draw a dot plot of Syto 16 vs. CD45 and analyze only Syto 16 positive cell (nucleated cells) (13). 6. CEPs are defined as positive for the nuclear staining Syto16, negative for anti-CD45 (used in this panel to exclude hematopoietic cells), positive for CD34 and CD133 (progenitor cell marker) and positive for VEGFR-2 (an endothelial cell marker). Figure 1 provides an example of CEP analysis (see Note 10).

4. Notes 1. A skilled flow cytometry operator should perform the procedure. 2. Staining, acquisition, and analysis procedures are the same for both fresh and frozen/thawed samples. 3. The volume of blood to be used is dependent on the WBC count. CEPs are rare, and in order to perform adequate acquisition, at least 100 events of CEPs should be collected at designated gates. 4. To assess the reproducibility of the procedure and its validation, three separate series of studies on both fresh and frozen/ thawed samples should be conducted. These procedures should be performed to assess the intra-operator and inter-operators variability. 5. To test variability of this procedure over time: Evaluate fresh samples on the same day of collection and after 24, 48, and 72 h; and evaluate frozen samples after 0, 2, 7, and 14 days of storage in liquid nitrogen.

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Fig. 1. Analysis of viable CEPs from a human blood specimen. After red blood cell lysis, and an acquisition of 106 cells, gate on mononuclear cells (a) followed by gating on Cyto16+/CD45− population (b). Cell viability is evaluated by being positive to the 7AAD dye (c). To evaluate CEP population, cells gated in (b) that are also CD133+/CD34+ (f) and VEGFR2+/CD34+ (g) are defined as CEPs. (d and e) Isotype control dot-plots.

6. Confirm that all antibodies have been stored correctly according to the manufacturer’s instructions and have not exceeded their expiration date. 7. Note: Prepare a master mix of antibodies if multiple samples are immunostained at the same time. It is important to use single color controls for compensation settings. 8. If using fluorescence-labeled antibodies from other manufacturers, the concentration should be optimized. 9. The samples should be held at 4°C and protect from light as bleaching of fluorochromes can occur. Samples should be acquired within an hour. 10. The combination of Syto16 and 7AAD is used to gain insight into cell viability according to van der Pol et al. (14). Necrotic cells were identified as Syto16low/7-AAD+; apoptotic cells as Syto16low/7-AAD−; and viable cells as Syto16bright/7-AAD−.

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References 1. Shaked Y, Voest EE (2009) Bone marrow derived cells in tumor angiogenesis and growth: are they the good, the bad or the evil? Biochim Biophys Acta 1796:1–4 2. Bertolini F, Shaked Y, Mancuso P, Kerbel RS (2006) The multifaceted circulating endothelial cell in cancer: towards marker and target identification. Nat Rev Cancer 6:835–845 3. Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T, Witzenbichler B, Schatteman G, Isner JM (1997) Isolation of putative progenitor endothelial cells for angiogenesis. Science 275:964–967 4. Lyden D, Hattori K, Dias S, Costa C, Blaikie P, Butros L, Chadburn A, Heissig B, Marks W, Witte L, Wu Y, Hicklin D, Zhu Z, Hackett NR, Crystal RG, Moore MA, Hajjar KA, Manova K, Benezra R, Rafii S (2001) Impaired recruitment of bone-marrow-derived endothelial and hematopoietic precursor cells blocks tumor angiogenesis and growth. Nat Med 7:1194–1201 5. Ruzinova MB, Schoer RA, Gerald W, Egan JE, Pandolfi PP, Rafii S, Manova K, Mittal V, Benezra R (2003) Effect of angiogenesis inhibition by Id loss and the contribution of bonemarrow-derived endothelial cells in spontaneous murine tumors. Cancer Cell 4:277–289 6. Li H, Gerald WL, Benezra R (2004) Utilization of bone marrow-derived endothelial cell precursors in spontaneous prostate tumors varies with tumor grade. Cancer Res 64:6137–6143 7. Ciarrocchi A, Jankovic V, Shaked Y, Nolan DJ, Mittal V, Kerbel RS, Nimer SD, Benezra R (2007) Id1 restrains p21 expression to control endothelial progenitor cell formation. PLoS One 2:e1338 8. Bertolini F, Paul S, Mancuso P, Monestiroli S, Gobbi A, Shaked Y, Kerbel RS (2003) Maximum tolerable dose and low-dose metronomic chemotherapy have opposite effects on the mobilization and viability of circulating endothelial progenitor cells. Cancer Res 63:4342–4346

9. Shaked Y, Henke E, Roodhart JM, Mancuso P, Langenberg MH, Colleoni M, Daenen LG, Man S, Xu P, Emmenegger U, Tang T, Zhu Z, Witte L, Strieter RM, Bertolini F, Voest EE, Benezra R, Kerbel RS (2008) Rapid chemotherapy-induced acute endothelial progenitor cell mobilization: implications for antiangiogenic drugs as chemosensitizing agents. Cancer Cell 14:263–273 10. Shaked Y, Bertolini F, Man S, Rogers MS, Cervi D, Foutz T, Rawn K, Voskas D, Dumont DJ, Ben-David Y, Lawler J, Henkin J, Huber J, Hicklin DJ, D’Amato RJ, Kerbel RS (2005) Genetic heterogeneity of the vasculogenic phenotype parallels angiogenesis; implications for cellular surrogate marker analysis of antiangiogenesis. Cancer Cell 7:101–111 11. Shaked Y, Ciarrocchi A, Franco M, Lee CR, Man S, Cheung AM, Hicklin DJ, Chaplin D, Foster FS, Benezra R, Kerbel RS (2006) Therapy-induced acute recruitment of circulating endothelial progenitor cells to tumors. Science 313:1785–1787 12. Bertolini F, Marighetti P, Shaked Y (2010) Cellular and soluble markers of tumor angiogenesis: from patient selection to the identification of the most appropriate postresistance therapy. Biochim Biophys Acta 1806(2):131–137 13. Mancuso P, Antoniotti P, Quarna J, Calleri A, Rabascio C, Tacchetti C, Braidotti P, Wu HK, Zurita AJ, Saronni L, Cheng JB, Shalinsky DR, Heymach JV, Bertolini F (2009) Validation of a standardized method for enumerating circulating endothelial cells and progenitors: flow cytometry and molecular and ultrastructural analyses. Clin Cancer Res 15:267–273 14. van der Pol MA, Broxterman HJ, Westra G, Ossenkoppele GJ, Schuurhuis GJ (2003) Novel multiparameter flow cytometry assay using Syto16 for the simultaneous detection of early apoptosis and apoptosis-corrected P-glycoprotein function in clinical samples. Cytometry B Clin Cytom 55:14–21

Chapter 15 Tracking Inflammation-Induced Mobilization of Mesenchymal Stem Cells Erika L. Spaeth, Shannon Kidd, and Frank C. Marini Abstract The act of migration is similar for many cell types. The migratory mechanism of mesenchymal stem cells (MSC) is not completely elucidated, yet many of the initial studies have been based on current understanding of the leukocyte migration. A normal function of MSC is the ability of the cell to migrate to and repair wounded tissue. This wound healing property of MSC originates with migration towards inflammatory signals produced by the wounded environment [1]. A tumor and its microenvironment are capable of eliciting a similar inflammatory response from the MSC, thus resulting in migration of the MSC towards the tumor microenvironment. We have shown MSC migration both in vitro and in vivo. In this chapter, we elucidate several in vivo methods to study MSC migration and mobilization to the tumor microenvironment. The first model is an exogenous model of MSC migration that can be performed in both xenograft and syngenic systems with in vitro expanded MSC. The second model utilizes transgenic-fluorescentcolored mice to follow endogenous bone marrow components including MSC. The third technique enables us to analyze data from the transgenic model through multispectral imaging. Furthermore, the migratory phenotype of MSC can be exploited for use in tumor-targeted gene delivery therapy has been efficacious in animal model studies and is an anticipated therapeutic device in clinical trials. Key words: MSC, Inflammation, Tumor bone marrow transplant, Transgenic mice

1. Introduction For the multipotent population of mesenchymal stem (stromal) cells (MSC), migration is an innate function. These cells naturally migrate to wounded environments whether it be a cutaneous lesion, infarcted tissue, or tumor. The MSC is a structurally supportive participant in wound closure, but is also able to promote an immune suppressive response to minimize the inflammation and thus promote healing (2–4).

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_15, © Springer Science+Business Media, LLC 2012

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MSC have been shown to migrate towards several tumor types including ovarian, breast, colon, prostate, and pancreatic carcinomas as well as lung metastases, sarcomas, and gliomas (5–14). Discordant evidence suggests both pro- and anti-tumorigenic effect of the MSC once in the tumor microenvironment (12, 15–18); however, when used as an anticancer agent delivery vehicle, the capabilities of the cell to migrate is maintained to ensure successful delivery of the MSC to the local tumor environment (9–11, 19, 20). Migration can be monitored in vitro using several common methods including the Transwell (Boyden chamber) Assay, the Scratch Assay, and the modified Ouchterlony Assay. In this chapter, we address in vivo migration models (bioluminescence and fluorescence) that will be evaluated using either syngenic or xenograft murine models that enable the investigator to follow the migration of a labeled cell through noninvasive techniques that allow the potential analysis of long-term physiological relevance of the migration without sacrificing the animal. PET/CT and MRI are not covered herein although the methods have been used to follow MSC in animal models (21–23). In addition, we address the use of fluorescently labeled transgenic murine model bone marrow transplant models that address the question of MSC mobility in an enclosed, endogenous system. These techniques will allow investigators to study the movement and localization of endogenous MSC to wounds and injured tissues including tumors, ischemic tissues, bone fractures, and muscle lacerations. Using multispectral imaging techniques, we are able to follow fluorescently expressing cell position within tissue sections to better understand the incorporation and involvement of MSC within injured tissues on a whole body/whole organ/whole tissue scale. While the following chapter addresses only one aspect of an inflamed environment, the tumor, these protocols are easily adapted to enable investigators to pursue many other forms of inflammatory induced migration of MSC.

2. Materials 2.1. MSC Isolation and Culture

1. Alpha Modified Eagle’s Medium (α-MEM) with supplements (L-glutamine and penicillin–streptomycin mixture and 20% fetal bovine serum (FBS)). 2. 0.25% Trypsin-EDTA. 3. Phosphate-buffered saline (PBS). 4. Ficoll-Hypaque gradient (for human MSC isolation from harvested bone marrow). 5. Sterile mortar and pestle (for murine MSC isolation).

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Table 1 Common MSC surface markers for human and murine MSC MSC surface markers Human

Murine

+

−

+

−

CD90

CD45

Sca-1

CD45

CD105

CD34

CD44

CD34

CD44

CD31

CD106

CD31

CD73

CD11b

CD140b

CD11b

CD166

C-Kit

CD146 CD140b GD2 CD271a a

CD271 is only found on freshly isolated human MSC, not on cultured MSC

6. 3 mg/ml Collagenase Type I (Worthington Biochemical) in α-MEM. 7. 180 cm2 Tissue culture dish. 8. Conjugated CD11b antibody (for murine MSC isolation— during immunodepletion step). 9. See Table 1 for list of additional antibodies used for MSC characterization. 2.2. MSC Differentiation 2.2.1. MSC Adipocyte Differentiation

1. Adipogenic induction medium (Dulbecco’s Modified Eagle’s Medium (DMEM), 10% FBS, penicillin, streptomycin, L-glutamine, 10 μg/ml insulin, 500 μM 3-isobutyl-1-methylxanthine, 1 μM dexamethasone, and 200 μM indomethacin). 2. Adipogenic maintenance medium: DMEM, 10% FBS, penicillin, streptomycin, L-glutamine, and 10 μg/ml insulin. 3. Isolated MSC. 4. 12-Well cell culture plate. 5. 10% Formalin. 6. Isopropanol (100 and 60%). 7. Oil Red O (Sigma) solution: Prepare Oil Red O working solution immediately prior to staining. 4 ml Water + 6 ml of 0.35%

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Oil Red O dissolved in 100% isopropanol. Filter twice with a 0.22 μm syringe filter. 2.2.2. MSC Osteoblast Differentiation

1. Isolated MSC. 2. 12-Well cell culture plate. 3. OsteoDiff Medium for osteoblast differentiation (Miltenyi Biotec, Auburn, CA). 4. Chromogenic alkaline phosphatase substrate BCIP/NBT (SIGMA FAST). 5. Alizarin Red S solution. 6. 100% Methanol. 7. PBS.

2.2.3. MSC Chondrocyte Differentiation

1. Isolated MSC. 2. 15 ml Polypropylene Falcon tube. 3. Chondrocyte differentiation medium: DMEM, penicillin, streptomycin, L-glutamine, 50 μg/ml ascorbic acid, 100 nM dexamethasone, and 10 ng/ml transforming growth factor β3 (TGF-β3). 4. PBS. 5. Formalin. 6. Histology core or processing capabilities. 7. Xylene. 8. 100% Alcohol (and dilutions 95, 90, 80, 70%). 9. 1% Alcian blue in 5% acetic acid. 10. Distilled water. 11. Nuclear fast red (Lab Vision Corporation). 12. Aqueous mounting medium.

2.3. In Vivo Mobilization and Tracking

1. αMEM with supplements (L-glutamine and penicillin– streptomycin mixture and FBS): 0–2% (for starvation) or 20% (for normal MSC). 2. 0.25% Trypsin-EDTA. 3. PBS. 4. Labeled tumor cell line of choice and MSC. 5. LipofectAmine and PLUS Technologies) (optional).

transfection

reagents

(Life

6. Plasmid-encoding GFP or other markers (optional). 7. CO2-independent medium (Optional). 8. Xenogen IVIS bioluminescence/fluorescence optical imaging system (Caliper Life Sciences).

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9. IVIS Living Image software (Caliper Life Sciences). 10. Isofluorane. 11. In vivo source—e.g. mouse. 12. Insulin syringe (1 cm3; 28 G ½). 13. Chemiluminescent substrate reagent depends on the bioluminescent expression system in your cells. Most common luciferase substrates: —

—

2.4. Mouse Bone Marrow Transplantation

Renilla requires colenterazine (40 mg/ml) (Biotium, Inc.) to prepare—resuspend in methanol. Firefly requires D-luciferin firefly-potassium salt (125 mg/ kg) (Biosynth).

1. Ubiquitously expressing GFP and RFP transgenic mice. 2. Insulin syringe. 3. Mouse restraint device for tail vein injection or isoflurane for ocular vein injection. 4. Sterile PBS. 5. Clear, Lucite box with air holes. 6. Approved radiation facility. 7. Antibodies for FACS sorting MSC from bone marrow population (investigator’s discretion).

2.5. Immunohistochemisrty/Immunofluorescence/ Multispectral Imaging

1. Tissues stored in formaldehyde or snap-frozen in liquid nitrogen and stored at −80°C. 2. OTC compound (Miles, Inc.) or formaldehyde. 3. Sodium citrate buffer: 10 mM sodium citrate, 0.05% Tween 20, pH 6.0. 4. Blocking buffer: 3% BSA, 1% FBS, and 1% species of secondary antibody (see Note 18). 5. Wash buffer: T-PBS: PBS with 0.5% Tween-20. 6. Antibodies of choice—extensive multicolor staining resources can be acquired elsewhere. 7. Hematoxylin-eosin and immunohistochemical (IHC) staining kits for IHC and fluorescently labeled antibody for choice for IF. 8. Research optical microscope with reflected and transmitted light sources (CRi Nuance attachment (Caliper Life Sciences) for multispectral imaging can alleviate concerns of background autofluorescence and multiple markers, but is not necessary). 9. A high resolution digital camera. 10. Adobe Photoshop, Slidebook, or InForm Software can be used to analyze image data.

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3. Methods 3.1. MSC Isolation and Characterization

3.1.1. MSC Isolation (Human)

While validated MSC cell lines can be purchased from the ATCC, they are also easily isolated from human or mouse (as explained below). Briefly, harvested bone marrow can be plated on plastic and the population of adherent cells that grows out from is enriched in the cell population we define as MSC. Phenotypic characterization by flow cytometry is one method to validate MSC population although there is no consensus on a single set of markers. Table 1 lists the most commonly used markers; however, those listed are not all inclusive nor should they be used exclusively to define MSC. The other standard to define MSC is to analyze the differentiation potential. Adipocyte, osteoblast, and chondrocyte differentiation assays are briefly discussed in this chapter (Fig. 1). 1. Collect clinical bone marrow sample (according to institutional protocol). 2. Separate mononuclear cells by centrifugation over FicollHypaque gradient (Sigma, St. Louis, MO). 3. Plate at initial seeding density of 1 × 106 cells/cm2. Size of tissue culture plate/flask can vary by experimental conditions. 4. After 3 days, remove the non-adherent cells by washing with PBS. 5. Culture adherent cell monolayer until confluency. 6. Trypsinize (0.25% trypsin with 0.1% EDTA) cells and subculture at densities of 5,000–6,000 cells/cm2. 7. Use cell passages 3–4 for the experiments (see Note 1).

3.1.2. MSC Isolation (Murine)

1. Anesthetize and sacrifice mouse according to institution approved protocol (see Note 2). 2. Remove the two hind limbs (femur and tibia) and the iliac crest (hip) (see Note 3). 3. Place clean bones in warmed αMEM (no serum needed—in a Petri dish or conical tube). 4. When all bones are removed, in sterile environment, fill an 18 G needle with serum media, cut the ends (both proximal and distal) of the bone and insert the needle to flush marrow out into a new, sterile tissue culture dish with warmed, serum (20%) αMEM medium. 5. Flushed bones can be crushed with mortal and pestle in warmed media. 6. Spin-down crushed bone (at 250 g) and resuspend in 3 mg/ ml Collagenase Type I 2 ml/mouse, and placed in a 37°C shaker at 50 × g for 45 min.

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Fig. 1. MSC characterization based on flow cytometry and differentiation assays. (a) Freshly isolated murine bone marrow sample stained with CD45. The subpopulation that contains MSC is found within the CD45 negative gate. Additional markers can be applied to the samples to select a more specific population. (b) First passage human MSC and freshly isolated murine MSC in culture are fibroblastic in appearance. (c) Adipogenic differentiation assay stains lipids with Oil Red O. Osteogenic differentiation assay has two stains, one to depict alkaline phosphatase activity and one to detect calcium deposits. Chondrogenic differentiation assay uses Alcian blue to detect mucopolysaccharides associated with chondroblast differentiation.

7. After incubation, bones were filtered out and cells were washed in PBS and added to the sterile culture dish in 20% αMEM. 8. Incubate at 37°C for about 3–5 days. 9. Discard supernatant and floating cells, culture adherent cell monolayer (MSC) until confluent.

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10. Trypsinize (0.25% trypsin with 0.1% EDTA) cells and subculture at densities of 5,000–6,000 cells/cm2. Size of tissue culture plate/flask can vary by experimental conditions. 11. Use cell passages 3–4 for the experiments (see Note 4). 3.2. Cell Differentiation Assays 3.2.1. Adipocyte Differentiation

1. Plate 2 × 104 MSC in a 12-well plate. 2. After cells reach confluence, change the medium to adipogenic induction medium for 72 h. 3. Change medium to the adipogenic maintenance medium for 24 h. 4. Repeat steps 2 and 3, induction/maintenance three times. 5. At the end of the third round, continue the adipogenic maintenance medium for 10 days changing the medium two times per week. 6. Fix adipocytes in 10% formalin for 1 h. 7. Wash adipocytes with 60% isopropanol and dry. 8. Lipid vacuoles are stained with Oil Red O solution. Place on dry adipocytes for 10 min. 9. Adipocytes and red lipid vacuoles can be imaged on any available microscope.

3.2.2. Osteogenic Differentiation

1. Plate 2.2 × 104 MSC subconfluently in a 12-well plate in OsteoDiff Medium. 2. Culture cells for 3 weeks, change medium 2× per week, with a volume of 1 ml per well. 3. Harvest cell culture between day 21 and day 30. 4. Fix cells with ice cold 100% methanol for 5 min then wash with PBS. 5. To visualize alkaline phosphatase activity, add alkaline phosphatase substrate to the well number 1 and incubate at 37°C for 10 min. Then, wash with water. 6. To visualize calcium deposits, add Alizarin Red S solution to the well number 2 and incubate at room temperature for 10 min. Then, wash with water. 7. Osteoblasts can be visualized on microscope of choice.

3.2.3. Chondrogenic Differentiation

1. Pellet 3.5 × 105 MSC in a 15 ml Falcon tube at 250 × g. 2. Resuspend the cells in chondrocyte differentiation medium. 3. Pellet suspension again and place into culture as a pellet— loosen the cap on the 15 ml Falcon tube for gas exchange. 4. Add 10 ng/ml TGF-β3 daily. Change chondrocyte differentiation medium three times per week—do not disturb the pellet.

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5. After 21 days, rinse pellet in PBS and fix in 10% formalin. 6. Paraffin embed and section for histology. 7. Deparaffinize in xylene and rehydrate in a series of alcohols. 8. To detect mucopolysaccharides associated with chondroblasts, stain with 1% Alcian blue in 5% acetic acid for 30 min. 9. Rinse with distilled water. 10. Counterstain with Nuclear Fast Red. 11. Mount in aqueous mounting medium. 12. Visualize on any microscope available. 3.3. In Vivo Mobilization and Migration of MSC

3.3.1. Tracking Exogenously Injected MSC Migration Towards Tumor

The therapeutic utilization of MSC has been attempted in numerous applications. It is well understood that MSC migrate towards sites of inflammation. Inflamed tissues produce factors that attract MSC to aid in wound repair. In this chapter, the site of inflammation that will be thoroughly addressed is the tumor microenvironment; however, MSC have been shown to migrate to other tissue environments including ischemic heart, ischemic brain, subcutaneous incision, and bone fracture (24–27). Location of tumor engraftment will depend on the tumor subtype as well as on analytical method at hand and can be placed subcutaneously, intravenously, intraperitoneally, or intracranially. Herein, we review three techniques that improve the in vivo analysis of MSC migration. Live cell bioluminescent (or fluorescent) imaging allows investigators to follow MSC migration within a live animal without the need to sacrifice it. Transgenic mice like ubiquitously expressing or promoter-specific expressing fluorescent-colored mice can be used to follow endogenous migration of MSC without exogenous manipulation of the system. This simple technique can be modified to address multiple questions. Finally, multispectral immunohistochemical/immunofluorescence imaging is a method that can be used to analyze complex histological markers in combination with one another. 1. First, the tumor cell line of choice needs to be stably labeled using a lenti or retroviral system. For this example, our system will label the breast cancer cell line, 4T1 with a GFP-tagged renilla-luciferine (GFP+/rLuc+) construct. Fluorescent Acquired Cell Sorting (FACS) sort labeled cells and expand for engraftment (see Note 5). 2. Prepare an adequate number of tumor cells for orthotopic tumor engraftment of 1 × 105 cells per injection in Nod-Scid mice (see Note 6). 3. Grow and label MSC in vitro similarly to tumor cell labeling but use an alternative luciferin construct (e.g., RFP-labeled firefly luciferin) (see Note 7).

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4. Engrafted tumors should be monitored weekly until appropriate size (see Note 8), larger than 5 mm diameter ensures visible and palpable tumor target. 5. To image rLuc + 4T1 in vivo, coelenterazine should be prepared immediately prior to injection. Substrate should be kept on ice and in the dark to maintain efficacy. Injections can be given intravenously or intraperitoneally—i.v. injections are more sensitive but take longer to complete and given the short half-life of the luciferase substrate. 6. Once tumors are adequate in size. Prepare MSC: 1 × 106 cells per injection resuspended in 100 μl of PBS. Intravenously inject MSC into Scid mice (see Note 9). 7. MSC can be imaged 0–3 h following injection to confirm the presence of MSC, which will be primarily in the lungs of the mice. The imaging substrate will be D-luciferin for the ffLuclabeled MSC, and should be prepared on ice and kept in the dark until IP injection similar to the coelenterazine. 8. To catch peak MSC recruitment to the tumor site, mice should be imaged every 24 h post MSC injection. Peak recruitment is between 48 and 72 h (Fig. 2). But imaging can be continued according to investigator’s discretion (see Note 10). 9. This system of exogenous recruitment of circulating MSC can be applied to many other injury/inflammatory models systems including cutaneous lacerations, muscular lacerations, bone fractures, and ischemic heart or brain models. This model can also be carried out using fluorescent cells instead of bioluminescent ones; however, there are a few limitations of the fluorescence model: (1) a nude, or shaved mouse is better; (2) autofluorescence can be a limitation when using fluorescent markers on the lower end of the spectrum; (3) depth of the tumor/signal within the animal can also be a problem that compounds the autofluorescent effect—the fluorescence may never be detected. 3.3.2. Mobilization of MSC Towards the Tumor Microenvironment

In this example, we address this question of endogenous MSC mobilization to the tumor microenvironment by using two fluorescently labeled transgenic C57/B6 mice: GFP (green) and RFP (red) (28). 1. Syngenic bone marrow transplant C57/B6 recipients should be between 6 and 10 weeks of age when lethally irradiated. Briefly, irradiate mice with a single dose of 9.5 Gy 4 h before donor bone marrow reconstitution (see Note 11). 2. Donor mice should be sacrificed and bone marrow collected. RFP-MSC from RFP transgenic bone marrow will be isolated and sorted by FACS. Briefly, bone marrow from the tibia, fibia, and iliac crest of the mice should be flushed, crushed with mortal and pestle, and resuspended in collagenase I for 1 h at

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Fig. 2. Bioluminescent images using luciferin substrates. (a) r-luc expressing 4T1 in a 96-well plate can be imaged. Average bioluminescent radiance observed can be plotted against number of tumor cells per well showing a linear correlation. (b) 2 h Post i.v. injection of mMSC shows bioluminescent activity in the lungs and at the injection sites in the tail. (c) In vivo image of murine breast cancer line, 4T1 labeled with r-luc on the left. Labeled tumor cells luminesce in the limbs of the mouse (in the inguinal adipose tissue where the two intra-fat tumor injections were initially given) and in the lungs (sites of metastases). On the right panel, ff-luc labeled mMSC 3 days after injection. The luminescent areas are dimmer, but overlap with the bilateral tumors as indicated by the arrowheads.

37°C at 50 × g (see MSC isolation in Subheading 3.1). Suspension can be collected and filtered (40 nm cell filter) before being labeled for FACS analysis (see Note 12). 3. Donor GFP transgenic bone marrow will be collected and depleted for MSC in the identical manner as described above and will be called the “non-MSC” bone marrow population. 4. Bone marrow fractions collected and isolated from either the GFP donor or the RFP donor will be admixed in vitro in a tube. Cell mixture ratio for one i.v. injection for one mouse will consist of 1 × 106 (can use between 1 × 106 and 1 × 107 bone

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marrow cells) GFP “non-MSC” bone marrow plus 1 × 106 RFP-MSC. See Fig. 3 for a diagram of the procedure. Conversely, alternative controls can be prepared using 1 × 106 RFP “non-MSC” bone marrow plus 1 × 106 GFP-MSC or the “non-MSC” populations alone (see Note 13).

Fig. 3. Diagram of bone marrow transplant schematic. Recipient mice are non-fluorescent and will be lethally irradiated 3–6 h prior to bone marrow re-derivation. Meanwhile, donor mice will be sacrificed and bone marrow will be collected and sorted by FACS to retrieve an “MSC” population derived from the RFP mouse and a total bone marrow minus MSC (or “nonMSC”) population from the GFP mouse. These populations will then be admixed according to an assay-dependent ratio and intravenously injected into the recipient mouse. The GFP/RFP bone marrow re-derivation can be confirmed after 21 days by flow cytometric analysis of drawn blood sample. Upon confirmation, tumors can be engrafted.

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5. Four hours following lethal irradiation, sorted RFP and GFP bone marrow mixtures will be i.v. injected into the recipient mouse (see Note 14). 6. Three weeks following bone marrow transplantation, the control mouse that received PBS alone will have died. Blood can be drawn from the mouse and analyzed by flow cytometry to confirm the presence of the fluorescent bone marrow. 7. Now that we have a C57/B6 mouse with GFP-labeled “nonMSC” or hematopoietic system and RFP-labeled MSC, we can engraft tumors into the mouse to elucidate the contributions of both the MSC and the “non-MSC” components that mobilize in response to tumor engraftment. The tumor model used in this model is the EO771, a C57/B6 breast tumor. Subcutaneous or orthotopic injections of 5 × 104 EO771 tumor cells will engraft into substantial 10–20 mm diameter tumors within three to five weeks. 8. Mice can be sacrificed at any number of time points leading up to the endpoint of the experiment. At time of sacrifice, tumors, organs, and other tissues can be collected for analysis by flow cytometry to quantify the number of RFP versus GFP cells are found within the tumor (or organ in question). The tissues can also be collected for immunofluorescence and/or immunohistochemistry, for which they can be snap frozen in OTC compound on dry ice and 100% ethanol, or they can be fixed in formalin (see Note 15). 9. This syngenic model with fluorescently labeled endogenous cells can be applied to many different models of inflammation including those previously mentioned like subcutaneous incisions or ischemic tissue models. The potential of syngenic models such as these leaves great potential for the elucidation of MSC as an important stromal cell involved in wound healing (see Note 16). 3.3.3. Imaging of Tumor Sections

1. The slides from the aforementioned bone marrow transplant experiment can be analyzed by immunofluorescent (IF) or IHC depending on the microscopy system available to the investigator. In this section, we focus mainly on IF with a multispectral imaging camera and software (CRi Inc., Woburn, MA) that enables the investigator to identify true fluorescence within high background tissue autofluorescence as well as identify multiple colors based on precise wavelength emission differences that might not be separated by a normal fluorescent microscope camera (see Note 17). 2. Tumor sections from paraffin-embedded blocks can be sliced in 5 nm sections. Briefly, slides can be deparaffinized in a series of xylene and ethanol washes prior to antigen retrieval for 20 min in boiling sodium citrate buffer. Tumor sections can alternatively be snap frozen instead of formalin preserved.

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3. Incubate slides in blocking buffer (see Note 18) for 30 min at room temperature (or overnight at 4°C). 4. If using an antibody, prepare dilution as per manufacturer’s instruction in blocking buffer. Incubate for 2 h at room temperature (or overnight at 4°C). In this example, we will be using a rabbit anti-RFP at a 1:200 dilution (see Note 19). 5. Wash in T-PBS for 5 min (3×). 6. Prepare a fluorescently labeled secondary antibody solution in blocking buffer at a 1:1,000 dilution. In this case, we will use goat-anti-rabbit Alexafluor 594 for 1 h at room temperature. The slides should be kept covered during this step. 7. Wash as done in step 5. 8. DAPI stain nucleus for 1 min with 5 mg/ml DAPI stock solution diluted 1:10,000. 9. Wash as done in step 5. 10. Rinse slides in water before applying fluorescent mounting media and a 1½ cover slip (170 nm thickness for optimal fluorescent image quality). 11. Let dry (protect from light) and apply nail hardener to the edges of the slide. 12. Imaging can be done on a microscope with a proper CRi Nuance camera attachment according to manufacturer’s manual. Figure 4 shows the image of a tumor section using the system described herein. 13. Data analysis software InForm (CRi Inc., Woburn, MA) allows for quantification of fluorescence-labeled cells within the image sections and can quantify based on cellular location or co-localization with additional color (see Note 20).

4. Notes 1. Cells can be sorted by flow cytometry for human MSC markers if a subpopulation is desired (CD44+, CD90+, CD105+, CD73+, CD166+, CD146+, CD140b+, and CD34−, CD45−). 2. Remove bone immediately after sacrificing the mouse. 3. Remove as much muscle tissue, skin, and fur from the bone using a scapula to scrape the remaining tissue away before placing in warmed αMEM. 4. Cells can be sorted by flow cytometry for mouse MSC markers if a subpopulation is desired (CD44+, Sca1+, NG2+ and CD34−, CD45−).

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Fig. 4. Immunofluorescent image of EO771 breast cancer tumor section. The Alexa594 staining for a stromal marker (big arrowheads) is depicted in white in the smaller, unmixed image and black in the large multispectral image. GFP cells (thin arrows) within the tumor are depicted in white in the smaller unmixed image, and in dark gray in the large multispectral image. DAPI nuclear staining is depicted in white in the smaller unmixed image and in light gray in the large multispectral image.

5. The tumor model system can be xenograft (human tumor and human MSC) or syngenic (murine tumor and murine MSC) depending on the system of choice. Because many transgenic mouse models exist, the potential for studying the mobilization of gene-modified (either +/− geneX) MSC towards the tumor microenvironment is extensive. 6. Number of tumor cells injected depends on the system in use. To get a significant (1–10 mm diameter tumor) of 4T1 cells in about 4 weeks, inject 2 × 105 cells per subcutaneous or orthotopic tumor site in 100 μl PBS. About 8 × 106 4T1 per T-175 flask at 85% confluency. 7. Start expanding MSC early enough to achieve 1 × 106 MSC per intravenous injection. MSC are readily labeled with lentiviral vectors according to common protocols. About 3 × 106 MSC per T-175 flask at 80% confluency.

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8. In these methods, the IVIS System from Caliper Life Sciences will be used, alternative bioluminescent and fluorescent live imaging systems can be used including Kodak’s In Vivo Multispectral Imaging System by Carestream or LI-COR’s Pearl Imager. 9. Intravenous injections can be given via lateral tail vein or ophthalmic plexus routes. The latter method requires the mouse to be anesthetized in order to insert the needle on the inner side of the ocular cavity, whereas the former method requires a restraint device, but no anesthetic. 10. If your study permits, mice can be sacrificed at any number of time points following MSC injection to confirm the in vivo bioluminescent MSC detection with more conventional IHC or FACS analysis of tumor sections to confirm the presence of MSC. 11. Syngenic bone marrow transplant requires good planning, and long time commitment to complete and will vary depending on the number of animals in the study. 12. Because the surface markers for MSC are still controversial, a select number of markers can be used based on the investigators discretion. Negative gating on CD45 and CD11b eliminate the hematopoietic and macrophage lineages, gating on the positive markers of known MSC subsets like NG2, PDGFRβ, or Sca-1 can be further used to derive a population of MSC. However, this population is not all inclusive. Collect both gated populations. These populations will be known as the “MSC” and “non-MSC” bone marrow fractions. 13. Bone marrow from the tibia, femur, and iliac crest of one mouse will give 3 × 107 total bone marrow cell, but only 1–3 × 106 MSC, therefore to prospectively isolate 1 × 106 MSC for each recipient mouse, you need at 3–5 donor mice per recipient. Alternatively, number of MSC used per bone marrow transplant are at the investigator’s discretion. The ratio of MSC to total bone marrow can be 1:1 or 1:100 (1:100 is a more biologically realistic ratio). 14. Intravenous injections can be given via lateral tail vein or ophthalmic plexus routes. The bone marrow mixture of RFP-MSC and GFP non-MSC or vice versa will be resuspended in 100 μl of PBS. As a control, one mouse will receive PBS alone. 15. Both frozen and paraffin-embedded tissue sections have been shown to produce good naïve images of fluorescent-labeled cells. 16. The most significant limitation is one of MSC characterizations. The heterogeneous population of MSC and the lack of defined surface markers or protein expression leaves a lot of room for interpretation. Therefore, initially defining your

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population of “MSC” is significant to the outcome of your experiment. As MSC are further studied, the use of MSCspecific promoters to drive reporter expression specific to MSC will be of great use in this model, where we will no longer need to prospectively isolate MSC based on unrefined marker expression. 17. If you naïve fluorescence is not visible under the microscope, respective fluorescent antibodies can be used to enhance the visualization of the fluorescently labeled cell. 18. For blocking buffer, you can use serum from the species that your secondary antibody is made in, e.g., using a goat-antirabbit Alexa Fluor secondary, use normal goat serum in the blocking buffer. 19. Proper controls are important to the experiment. When working with a multicolored tissue, it is good to have single colored controls for each color in use. For example, our final slide will contain four colors: DAPI, Alexa647, GFP, and RFP. Therefore, we will need five control slides. One background slide of tumor only to account for background tissue autofluorescence. One tumor slide with RFP cells present only. One tumor slide with GFP cells present only. One tumor slide with DAPI only and one tumor slide with Alexa647 only. These controls will allow us to set up a proper “library” of positive controls for our final, multicolor slide. 20. These methods are not limited to the CRi system, any functional fluorescent microscope can be used to suite the investigators needs. References 1. Spaeth E, Klopp A, Dembinski J, Andreeff M, Marini F (2008) Inflammation and tumor microenvironments: defining the migratory itinerary of mesenchymal stem cells. Gene Ther 10:730–738 2. Sasaki M, Abe R, Fujita Y, Ando S, Inokuma D, Shimizu H (2008) Mesenchymal stem cells are recruited into wounded skin and contribute to wound repair by transdifferentiation into multiple skin cell type. J Immunol 180: 2581–2587 3. Wu Y, Chen L, Scott PG, Tredget EE (2007) Mesenchymal stem cells enhance wound healing through differentiation and angiogenesis. Stem Cells 25:2648–2659 4. Li H, Fu X, Ouyang Y, Cai C, Wang J, Sun T (2006) Adult bone-marrow-derived mesenchymal stem cells contribute to wound healing of skin appendages. Cell Tissue Res 326: 725–736

5. Studeny M, Marini FC, Champlin RE, Zompetta C, Fidler IJ, Andreeff M (2002) Bone marrowderived mesenchymal stem cells as vehicles for interferon-beta delivery into tumors. Cancer Res 62:3603–3608 6. Dwyer RM, Potter-Beirne SM, Harrington KA, Lowery AJ, Hennessy E, Murphy JM et al (2007) Monocyte chemotactic protein-1 secreted by primary breast tumors stimulates migration of mesenchymal stem cells. Clin Cancer Res 13:5020–5027 7. Coffelt SB, Marini FC, Watson K, Zwezdaryk KJ, Dembinski JL, Lamarca HL et al (2009) The pro-inflammatory peptide LL-37 promotes ovarian tumor progression through recruitment of multipotent mesenchymal stromal cells. Proc Natl Acad Sci USA 106:3806–3811 8. Sordi V, Malosio ML, Marchesi F, Mercalli A, Melzi R, Giordano T et al (2005) Bone marrow mesenchymal stem cells express a restricted set

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of functionally active chemokine receptors capable of promoting migration to pancreatic islets. Blood 106:419–427 9. Kim SM, Lim JY, Park SI, Jeong CH, Oh JH, Jeong M et al (2008) Gene therapy using TRAIL-secreting human umbilical cord bloodderived mesenchymal stem cells against intracranial glioma. Cancer Res 68:9614–9623 10. Sonabend AM, Ulasov IV, Tyler MA, Rivera AA, Mathis JM, Lesniak MS (2008) Mesenchymal stem cells effectively deliver an oncolytic adenovirus to intracranial glioma. Stem Cells 26:831–841 11. Ren C, Kumar S, Chanda D, Kallman L, Chen J, Mountz JD et al (2008) Cancer gene therapy using mesenchymal stem cells expressing interferon-beta in a mouse prostate cancer lung metastasis model. Gene Ther 15:1446–1453 12. Khakoo AY, Pati S, Anderson SA, Reid W, Elshal MF, Rovira II et al (2006) Human mesenchymal stem cells exert potent antitumorigenic effects in a model of Kaposi’s sarcoma. J Exp Med 203:1235–1247 13. Kanehira M, Xin H, Hoshino K, Maemondo M, Mizuguchi H, Hayakawa T et al (2007) Targeted delivery of NK4 to multiple lung tumors by bone marrow-derived mesenchymal stem cells. Cancer Gene Ther 14:894–903 14. Hung S, Deng W, Yang WK, Liu R, Lee C, Su T et al (2005) Mesenchymal stem cell targeting of microscopic tumors and tumor stroma development monitored by noninvasive in vivo positron emission tomography imaging. Clin Cancer Res 11:7749–7756 15. Hata N, Shinojima N, Gumin J, Yong R, Marini F, Andreeff M et al (2010) Platelet-derived growth factor BB mediates the tropism of human mesenchymal stem cells for malignant gliomas. Neurosurgery 66:144–156 16. Spaeth EL, Dembinski JL, Sasser AK, Watson K, Klopp A, Hall B et al (2009) Mesenchymal stem cell transition to tumor-associated fibroblasts contributes to fibrovascular network expansion and tumor progression. PLoS One 4:e4992 17. Kidd S, Spaeth E, Klopp A, Andreeff M, Hall B, Marini FC (2008) The (in) auspicious role of mesenchymal stromal cells in cancer: be it friend or foe. Cytotherapy 10:657–667 18. Nakamura K, Ito Y, Kawano Y, Kurozumi K, Kobune M, Tsuda H et al (2004) Antitumor effect of genetically engineered mesenchymal stem cells in a rat glioma model. Gene Ther 11:1155–1164

19. Studeny M, Marini FC, Dembinski JL, Zompetta C, Cabreira-Hansen M, Bekele BN et al (2004) Mesenchymal stem cells: potential precursors for tumor stroma and targeted-delivery vehicles for anticancer agents. J Natl Cancer Inst 96:1593–1603 20. Kidd S, Caldwell L, Dietrich M, Samudio I, Spaeth EL, Watson K et al (2010) Mesenchymal stromal cells alone or expressing interferon-beta suppress pancreatic tumors in vivo, an effect countered by anti-inflammatory treatment. Cytotherapy 12:615–625 21. Ju S, Teng G, Zhang Y, Ma M, Chen F, Ni Y (2006) In vitro labeling and MRI of mesenchymal stem cells from human umbilical cord blood. Magn Reson Imaging 24:611–617 22. Gonzalez-Lara LE, Xu X, Hofstetrova K, Pniak A, Chen Y, McFadden CD et al (2011) The use of cellular magnetic resonance imaging to track the fate of iron-labeled multipotent stromal cells after direct transplantation in a mouse model of spinal cord injury. Mol Imaging Biol 13:702–711 23. Lo Celso C, Fleming HE, Wu JW, Zhao CX, Miake-Lye S, Fujisaki J et al (2009) Live-animal tracking of individual haematopoietic stem/ progenitor cells in their niche. Nature 457:92–96 24. Chen D, Zhang Z, Wu X, Lin JH (2007) Distribution of intravenously grafted bone marrow mesenchymal stem cells in the viscera tissues of rats before and after cerebral ischemia. J Clin Rehab Tissue Eng Res 11: 10160–10164 25. Barbash IM, Chouraqui P, Baron J, Feinberg MS, Etzion S, Tessone A et al (2003) Systemic delivery of bone marrow-derived mesenchymal stem cells to the infarcted myocardium: feasibility, cell migration, and body distribution. Circulation 108:863–868 26. Wu GD, Nolta JA, Jin YS, Barr ML, Yu H, Starnes VA et al (2003) Migration of mesenchymal stem cells to heart allografts during chronic rejection. Transplantation 75:679–685 27. Kidd S, Spaeth E, Dembinski JL, Dietrich M, Watson K, Klopp A et al (2009) Direct evidence of mesenchymal stem cell tropism for tumor and wounding microenvironments using in vivo bioluminescence imaging. Stem Cells 10:2614–2623 28. Kidd S, Spaeth E, Watson K, Burks J, Lu H, Klopp A et al (2012) Origins of the tumor microenvironment: quantitative assessment of adipose-derived and bone marrow-derived stroma. PLoS One 7:e30563

Chapter 16 Differentiation of Circulating Monocytes into Fibroblast-Like Cells Darrell Pilling and Richard H. Gomer Abstract Monocytes are produced in the bone marrow and enter the blood. They generally leave the blood and enter a tissue, and then become macrophages. In healing wounds, circulating monocytes also enter the tissue and instead of becoming macrophages, can differentiate into fibroblast-like cells called fibrocytes. Fibrocytes are also present in the lesions associated with fibrosing diseases such as congestive heart failure, end stage kidney disease, and pulmonary fibrosis. We have found that culturing blood monocytes, or white blood cell preparations containing monocytes, in serum-free media permits some of the monocytes to differentiate into fibrocytes within 5 days, and that this differentiation is inhibited by the blood plasma protein serum amyloid P. Key words: Fibrocytes, Fibrosis, Inflammation, Monocytes, Pentraxin, Serum-free culture, Serum amyloid P

1. Introduction Fibrocytes are spindle-shaped fibroblast-like cells that are involved in both tissue repair and fibrosis (1–6). Fibrocytes differentiate from peripheral blood monocytes, and express markers of both hematopoietic cells (CD34, CD45, LSP-1, and MHC class II) and stromal cells (collagen and prolyl-4-hydroxylase) (1, 2, 7–10). Fibrocytes have both beneficial and detrimental effects on health. Fibrocytes can regulate innate and adaptive immune responses, by the secretion of a variety of cytokines and growth factors (2, 11– 14). Fibrocytes also promote wound healing both directly and also by secreting factors that activate fibroblasts (6, 15). Fibrocytes are also implicated in chronic inflammation and fibrosis, and have been detected in tumors, hypertrophic scars, bronchial asthma, pulmonary

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_16, © Springer Science+Business Media, LLC 2012

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fibrosis, renal fibrosis, some diseased heart valves, and nephrogenic systemic fibrosis (3, 15–21). The original in vitro culture conditions developed by Bucala and colleagues using DMEM-based culture media supplemented with 10% fetal bovine serum, has been used to generate human, murine, and porcine fibrocytes from peripheral blood (1, 2, 11, 13, 22). This system involves incubating peripheral blood mononuclear cells (PBMC), a mixture of monocytes, B, NK, and T cells, for 2 days before the removal of the non-adherent B, NK, and T cells, and then culturing the remaining adherent cells for 10–14 days (22). After 10–14 days, fibrocytes are then recovered from culture by trypsin-EDTA and used for subsequent experiments. In addition, fibronectin is often used to coat the tissue culture vessels prior to the addition of the PBMC. The fibronectin may promote fibrocyte differentiation and also permits the removal of the fibrocytes from the tissue culture plastic after the 14-day culture period (22). We observed that human, murine, and rat PBMC cultured in serum-free medium gave rise to fibrocytes and some macrophages within 5 days. However, at 5 days in medium containing serum no fibrocytes appear, and more macrophages were observed (8, 9, 23, 24). This time-period appears closer to the 3–7 days that fibrocytes take to appear in wounds, inflammatory sites, and fibrotic lesions (1, 2, 12, 15–17, 25–27). The other major advantage of a serumfree culture system is that all the components are defined and there are no unknown compounds from complex biological fluids such as serum. The fibrocytes from serum-free cultures appear to be comparable to those generated over 2 weeks in the presence of serum as they express the same markers, and secrete and respond to the same range of cytokines and growth factors (10, 23, 24). We have also been able to use the serum-free culture of fibrocytes to identify factors that affect fibrocytes differentiation. We have found that glucose, immunoglobulins, insulin, some cytokines, and some extracellular matrix proteins, can regulate fibrocyte differentiation (9, 23, 28). We have also found that the plasma protein serum amyloid P (SAP) inhibits fibrocyte differentiation in vitro, and that in vivo SAP can reduce experimentally induced fibrosis in both lungs and heart, and also inhibit dermal wound healing (8, 26, 27, 29, 30).

2. Materials All materials and solutions used should be sterilized either by irradiation, 0.2 μm filtering, or exposure to ethylene oxide gas. Also, all materials used should be (if possible) “virgin plastic” disposable cell culture grade; standard laboratory glassware should be avoided to reduce exposure to endotoxins, detergents, or other contaminants. All reagents are stored at room temperature unless indicated.

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1. Heparin vacutainer tubes (#367874, 150 U per tube; BD Bioscience; San Diego, CA). 2. Ficoll-Paque Plus (GE Healthcare Biosciences, Piscataway, NJ). 3. Phosphate buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4 pH 7.4). 4. 15 ml and 50 ml polypropylene tubes. 5. Sterile plastic transfer pipettes.

2.2. Cell Culture Reagents

1. FibroLife basal medium (Lifeline Cell Technology, Walkersville, MD). Store at 4°C. 2. 1 M HEPES (Sigma-Aldrich, St. Louis, MO). Store at 4°C. 3. 100× Nonessential amino acids (NEAA, Sigma-Aldrich). Store at 4°C. 4. 100 mM sodium pyruvate (Sigma-Aldrich). Store at 4°C. 5. 200 mM glutamine (Hyclone, Thermo Scientific, Logan, UT). Store at −20°C. 6. Mixture of penicillin (10,000 U/ml) and streptomycin (10,000 μg/ml) (Hyclone). Store at −20°C. 7. 100× ITS-3 (Sigma-Aldrich). Store at 4°C. 8. Flat-bottomed 24, 48, or 96-well tissue culture plates (BD Biosciences). 9. Eosin and methylene blue solutions diluted 1:5 (v:v) in water (Hema 3 Stain, Fisher Scientific, Middletown, VA).

2.3. Preparation of Serum Amyloid P

1. Human, mouse, rat, or pig serum (Gemini Bio-Products, West Sacramento, CA). Store at −80°C. 2. 50 ml and 1000 ml, 0.2 μm tissue culture filter units (Nalgene, Thermo Scientific). 3. ECH Sepharose 4B beads (GE healthcare Life Sciences, Piscataway, NJ). Store at 4°C. hydro4. N-(3-Dimethylaminopropyl)-N¢-ethylcarbodiimide chloride, also called N-ethyl-N¢-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC or EDAC) (Sigma-Aldrich). Store desiccated at −20°C. 5. Phosphoethanolamine (Sigma-Aldrich). Store at −20°C. 6. Phosphocholine beads (Pierce, Thermo Scientific, Rockford, IL). Store at 4°C. 7. Agarose beads (SP Sepharose FF, GE-Healthcare Biosciences). Store at 4°C. 8. Amicon Ultra-15 ml centrifugal filters with 10 kDa cutoff (Millipore, Billerica, MA). 9. Amicon Ultra-0.5 ml centrifugal filters with 10 kDa cutoff (Millipore).

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10. Solution A. 500 ml H2O, pH 4.5 (~1 μl of 12 N HCl in 500 ml H2O). 11. Solution B. 500 ml of 500 mM NaCl (14.6 g in 500 ml H2O). 12. Acetate buffer. 61.5 ml of 0.2 M acetic acid (11.55 ml of glacial acetic acid per liter), 13.5 ml of 0.2 M Na acetate (27.2 g Na acetate trihydrate per liter), and 4.38 g of NaCl (500 mM). Adjust to 150 ml total with H2O. 13. 100 mM Tris buffer. 15 ml of stock 1 M Tris–HCl pH 8 and 4.38 g NaCl, adjust to 150 ml with H2O. 14. Tris/NaCl/CaCl2 binding buffer. 20 mM Tris pH 8, 140 mM NaCl, 2 mM CaCl2. 15. Tris/NaCl/EDTA elution buffer. 20 mM Tris pH 8, 140 mM NaCl, 10 mM EDTA. 16. 20 mM sodium phosphate buffer pH 7.4. For 200 mM sodium phosphate buffer, mix 19 ml of 200 mM sodium phosphate monobasic (27.6 g NaH2PO4 ·H2O per liter) with 81 ml of 200 mM sodium phosphate dibasic (53.65 g Na2HPO4 · 7H2O per liter). Dilute to 20 mM with water. 17. Filter sterilize all solutions with 0.2 μm Nalgene vacuum filter sterilization units.

3. Methods As Bucala and colleagues have recently published methods on the culture of fibrocytes in serum-containing conditions (22), we have focused this chapter on the culture of fibrocytes in serum-free conditions. This chapter also focuses on the culture of human fibrocytes, but we have also recently published protocols for the serum-free culture of murine fibrocytes from blood (24) and we and others have cultured fibrocytes from murine spleen preparations (20, 50). 3.1. Preparation of Human Peripheral Blood Mononuclear Cells

All procedures must comply with and have the specific approval of your institution’s Institutional Review Board. All procedures must use appropriate BSL-2 laboratory facilities, in accordance with your institution’s Office of Biosafety. 1. 20 ml of human peripheral blood is collected by venipuncture from healthy adult volunteers using heparin as an anticoagulant (see Note 1). For 20 ml of blood, two 10 ml vacutainer tubes will be needed (see Note 2). 2. After the blood is collected, invert tubes 3–4 times to mix the blood and heparin.

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3. If the PBMC are not to be isolated from the blood immediately, the tubes can be stored for 1–3 h in the dark at room temperature (20–23°C). 4. Transfer the blood from the vacutainer tube to a 50 ml polypropylene tube, either using a sterile plastic transfer pipette, or by simply pouring the blood into the 50 ml polypropylene tube. Then add an equal volume of PBS to a final volume of approximately 40 ml (see Note 3). 5. Add 3 ml of Ficoll-Paque to a 15 ml polypropylene tube, repeat for three additional tubes (see Note 4). 6. Gently layer 10 ml of diluted blood over the 3 ml Ficoll-Paque (see Note 5), repeat for the remaining three tubes. These volumes leave 2 ml of extra space which will help in the following steps when a transfer pipette is inserted through the plasma to remove the PBMC at the Ficoll interface, see step 11. 7. Isolate the PBMC by centrifugation at room temperature for 40 min at 400 × g, using swing-out buckets with screw top lids attached. 8. Carefully remove the centrifuge buckets from the centrifuge, so as not to disturb the PBMC at the Ficoll interface. For this step, and all subsequent steps, place the centrifuge buckets with the lids still attached inside a class II cabinet, and only then unscrew the lids and remove the tubes from the centrifuge bucket (see Note 6). 9. Carefully remove the PBMC from the diluted serum-Ficoll interface with a sterile plastic transfer pipette (see Note 7). This procedure should remove approximately 2–3 ml of liquid containing the PBMC. Add the PBMC to a new 15 ml polypropylene tube. Repeat the procedure for the three remaining tubes. We generally combine two tubes of recovered material into one 15 ml tube. Therefore, for four tubes of blood-Ficoll mix, two new 15 ml polypropylene tubes will be required. 10. Add 10 ml of PBS to each tube, mix by inversion, and collect the PBMC by centrifugation for 10 min at 300 × g at room temperature (see Note 8). 11. Pour off the excess PBS and resuspend the cell pellet with 1–2 ml of PBS, and then add an additional 10 ml of PBS to each tube. Recentrifuge cells for 10 min at 300 × g. 12. Again resuspend the cell pellet, and then combine the four cell pellets into one tube, and add PBS to 13 ml total (see Note 9). 13. Repeat the centrifugation step. 14. Repeat the process of resuspending the cells in PBS and collecting the cells by centrifugation for a total of six times (see Note 10).

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15. After the sixth centrifugation step, resuspend the PBMC in 10 ml of PBS and count the cells with a hemocytometer. Collect the cells by centrifugation as above. At this point the cells can be resuspended in 1 ml Fibrolife serum-free medium (see recipe below) and stored on ice. If necessary, the PBMC can be kept in a covered ice bucket in a cold-room for up to 24 h, although we rarely store cells for more than 3 h. We recover approximately 20 × 106 cells from 20 ml of blood. 3.2. Differentiation of Fibrocytes in Serum-Free Culture

1. Prepare FibroLife serum-free medium (FibroLife-SFM) by mixing 47.0 ml FibroLife basal medium in a 50 ml polypropylene tube with 0.5 ml of each of the following supplements; HEPES, NEAA, sodium pyruvate, glutamine, penicillin/streptomycin, and ITS-3 (see Note 11). 2. Resuspend the PBMC to 5 × 105 cells/ml in FibroLife-SFM (see Note 12). We have also shown that density has an effect on fibrocyte differentiation, and that PBMC cultured between 2.5 and 5 × 105 cells/ml is optimal for fibrocyte differentiation (24). 3. Add 0.1 ml cell suspension per well to 96-well plates, along with 0.1 ml of Fibrolife-SFM or Fibrolife-SFM containing a test compound. 4. Incubate cells in a humidified incubator containing 5% CO2 at 37°C. Fibrocytes will begin to be visible by microscopy in approximately 2 days. 5. After 5 days, fibrocytes will be very obvious on a low power inverted microscope, due to their elongate morphology (50– 200 μm long spindle-shaped cells), compared to small round lymphocytes (8–10 μm cells), dendritic cells (10–15 μm cells, with many dendritic-like processes protruding from the cell), and larger irregular-shaped macrophages (15–20 μm cells, with a large nucleus and pronounced cytoplasm) (Fig. 1a).

Fig. 1. Appearance of human fibrocytes in serum-free conditions. (a) Viable human PBMC cultured in SFM for 5 days. (b) PBMC air-dried and stained with eosin and methylene blue. Solid arrow points to a fibrocyte, white arrow points to a dendritic cell, and asterisk is to the left of a macrophages. Bar is 50 μm.

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We observe approximately 1,000–2,000 fibrocytes per 1 × 105 PBMC (see Note 13). 6. At this point the cells can be air-dried, fixed, stained, and then stored at room temperature in the dark. We have plates fixed and stained in 2001 that are still easily viewable in 2010. 7. Remove the liquid either by aspirating off the liquid with an eight (or 12)-well multichannel pipette, or simply by flicking out the liquid into a biosafety bag containing paper tissues to soak up the excess liquid. Then invert the plate over the vents inside a class II safety cabinet and air dry the plates for at least 2 h. Cells can then be fixed by adding 0.1 ml methanol to each well using a multichannel pipette and leaving the plate for 10 min at room temperature. Methanol is then flicked out as above, and the plates air dried again (see Note 14). 8. Stain the cells by adding 0.1 ml of the diluted solution of eosin to each well with a multichannel pipette. Incubate the plate for 1–2 min. Then flick out the eosin and add 0.1 ml of the diluted solution of methylene blue to each well. Again leave for 1–2 min and then flick out the liquid, and gently rinse the whole plate with standard distilled water, and then air dry the plate. Store plates in the dark at room temperature. 9. Fibrocytes can then be counted using a standard inverted microscope (Fig. 1b). We routinely count duplicate wells for fibrocytes from five different 900 μm diameter fields per well. All cultures are counted by at least two independent observers blinded to the experimental design. If necessary, macrophages, lymphocytes, and dendritic cells can also be enumerated in the same wells. Fibrocytes are defined as adherent 50–200 μm long spindle-shaped cells with an oval nucleus, as described previously (8–10, 23). 3.3. Preparation of Serum Amyloid P from Serum or Plasma

We have previously shown that fibrocyte differentiation is inhibited by a variety of molecules but that SAP appears to be the dominant inhibitor of fibrocyte differentiation (8, 23, 26, 29). The original methods used to isolate SAP from plasma, serum, or amyloid deposits used a variety of techniques. These methods included using ultracentrifugation through sucrose gradients (31); chromatography using cross-linked dextran (Sephadex) (32) or polyacrylamide (Bio-Gel P) (33) beads; and calcium-dependent binding to agarose (34, 35) or phosphoethanolamine (36, 37). However, agarose varies in the ability to bind SAP (37, 38), due to the differential amounts of pyruvate present in the agarose preparations (39). Therefore, as the calcium-dependent binding of human and murine SAP to phosphoethanolamine beads is specific, this method has become the standard matrix to purify SAP from human and murine serum (37, 40).

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3.4. Preparing Phosphoethanolamine Beads

Day 1 1. Take 25–30 ml of ECH-Sepharose 4B gel slurry and place it in a 50 ml tube top filter device (Nalgene). Apply vacuum, and rinse first with 40 ml of the pH 4.5 H2O wash (solution A), followed by 40 ml of the 500 mM NaCl wash (solution B). Repeat this alternating wash process four times. 2. Dissolve 100–150 mg of phosphoethanolamine in 25 ml pH 4.5 H2O (solution A) and add to the ECH-Sepharose slurry contained in the top of the filtration device. Transfer slurry to a 50 ml polypropylene tube. 3. Add 400–500 mg EDC (Sigma) powder to the Sepharose/ phosphoethanolamine slurry and mix using a rotary mixer. 4. Leave overnight at 4°C on a rotary mixer. Day 2 5. Recover the beads in the 50 ml polypropylene tube by centrifugation at 300 × g for 60 s. Pipette off the excess liquid, do not pour off liquid as beads will be lost. Then wash the beads with 25 ml acetate buffer. Mix for 2 min, spin 300 × g for 60 s, and pipette off excess liquid. Next, add 25 ml Tris buffer, and repeat mixing, centrifugation, and removal of liquid steps. 6. Repeat washes three additional times (4 × 25 ml washes for each wash buffer), alternating between the acetate and Tris wash buffers. 7. Wash beads as above twice with 25 ml H2O by centrifugation at 300 × g for 60 s, and remove excess liquid. Follow with two washes in 25 ml of Tris/NaCl/CaCl2 binding buffer. 8. Store beads at 4°C in an equal volume Tris/NaCl/CaCl2 binding buffer. All batches of phosphoethanolamine beads should be tested for their ability to bind to commercial SAP preparations. Take 50 μl of the phosphoethanolamine bead slurry and mix with 200 μl of 50 μg/ml SAP. Incubate for 60 min and then separate the beads and non-bound material by centrifugation. Analyze the presence of SAP in the unbound fraction, following the protocols in Subheading 3.5, step 9.

3.5. Purification of SAP from Human, Murine, Rat, or Pig Serum

This procedure describes a simple batch process. The following purification can also be performed using column chromatography following the recommended protocols of the manufacturers of the specific columns. 1. Remove 100 ml of human, mouse, rat, or pig serum from −80°C and defrost overnight at 4°C.

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2. Mix serum by inversion and then pipette 12 ml of serum into eight 15 ml polypropylene tubes, and add 3 ml of phosphoethanolamine bead slurry. 3. Mix at room temperature for at least 1 h, using a rotary mixer. 4. Spin down at 300 × g for 60 s and remove supernatant (see Note 15). 5. Wash beads by adding 10 ml Tris/NaCl/CaCl2 binding buffer. Mix for 10 min at room temperature, using a rotary mixer. Centrifuge at 300 × g for 60 s and remove supernatant. To increase bead recovery, one can perform an extra centrifugation step on the supernatant before discarding the supernatant. 6. Add 2 ml of Tris/NaCl/CaCl2 binding buffer to resuspend the beads in each tube and then combine the beads from two tubes. This reduces the amount of buffer used for subsequent steps. Repeat wash procedure with 10 ml Tris/NaCl/CaCl2 binding buffer at least four times. 7. Add an equal volume of Tris/NaCl/EDTA elution buffer to the beads and combine beads into one tube. Mix overnight at 4°C using a rotary mixer. Next day, centrifuge at 300 × g to pellet the beads and remove elution buffer. The elution buffer now contains the purified SAP. Repeat elution by adding fresh elution buffer to the beads and mix for 30 min at room temperature. Centrifuge beads and recover the elution buffer and combine with the first elution. Then centrifuge the elution buffer at 1,000 × g to remove any remaining beads. 8. Desalt and concentrate the elution buffer containing the SAP using a 15 ml centrifugal filter device with 10 kDa cutoff. Prewash the device with 10 ml H2O by centrifugation for 5 min at 3,000 × g to remove residual glycerol present on the filter. Repeat the wash step twice with 10 ml of 20 mM sodium phosphate buffer pH 7.4. Apply the combined elution buffer containing the SAP to a centrifugal filter and add 20 mM Na phosphate buffer to 15 ml final volume. Centrifuge for 10 min at 3,000 × g, so that only approximately 1 ml of material is in the upper filter chamber. Add 14 ml of 20 mM Na phosphate buffer and repeat the buffer exchange at least three times. 9. At this point the material can be tested for purity and concentration using PAGE and/or western blotting using anti-SAP antibodies, as described previously (8, 27, 29, 38). Additionally, to quickly assess the protein concentration take 4 μl of sample and dilute with 396 μl of H2O into an Eppendorf tube, and take 4 μl of sodium phosphate buffer and dilute with 396 μl of H2O into an Eppendorf tube. Take the two tubes and perform an UV scan at 280 and 340 nm, using the phosphate buffer as the blank control. Additionally, the protein content can be assessed using a standard Bradford protein assay.

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Fig. 2. Example of purified SAP from murine serum. Purified mouse SAP was analyzed by PAGE, on a 4–15% reducing gel, and stained with Coomassie. Lane 1 Protein molecular weight markers (BenchMark protein standards, Invitrogen, Carlsbad, CA). Lanes 2–6 300, 100, 30, 10 and 3 μg/ml human SAP loading controls. Lane 7 Undiluted murine SAP preparation. Lanes 8–13 Murine SAP preparation diluted 1/10, 1/20, 1/40, 1/80, 1/100, and 1/160 with sodium phosphate buffer, respectively.

10. We find that a repeat purification step using phosphoethanolamine beads is usually necessary. Dilute the concentrate from step 8 with 20 mM Na phosphate buffer, pH 7.4, to a final volume of 4 ml. Add 6 ml of PE bead slurry with additional CaCl2 (8 μl of 1 M solution) to obtain a final concentration of 2 mM CaCl2 for 10 ml, and mix for at least 1 h at RT. Repeat steps 4–8. 11. Dilute the final concentrated sample to 500 μg/ml SAP using 20 mM sodium phosphate buffer and store at 4°C. The final SAP preparation should be a single 27 kDa band when analyzed by PAGE on a 4–15% reducing gel and stained with Coomassie or silver stain (Fig. 2). 12. As an additional purification step use phosphocholine beads from Pierce/Thermo Scientific (Cat #20307; immobilized p-aminophenyl phosphoryl choline) to remove any residual levels of CRP from the purification. In humans and mice, the concentration of CRP in normal serum is only 1–2 μg/ml, therefore only small amounts of resin are needed. For rat serum, where the CRP levels can be as high as 300–500 μg/ml (41) large volumes of p-aminophenyl phosphoryl choline agarose beads may be required (see Note 16). Alternatively, pneumococcal C-polysaccharide linked beads can be used to remove any contaminating CRP from preparations of SAP (37, 42, 43).

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As discussed above, SAP binds to agarose in a calcium-dependent manner, but different preparations, brands, and batches of agarose have differing capacities to bind SAP. However, we have found that SP Sepharose FF appears to bind SAP efficiently irrespective of the batches tested between 2003 and 2010 (38). 1. Wash 30 ml pre-hydrated agarose beads (SP Sepharose FF) four times in 10 volumes of Tris/NaCl/CaCl2 binding buffer, by collecting the beads by centrifugation at 300 × g for 60 s. Store beads in Tris/NaCl/CaCl2 buffer at 4°C. 2. Resuspend the bead slurry in 10 ml Tris/NaCl/CaCl2 buffer and add 3 ml to a 15 ml conical polypropylene tube containing 12 ml serum. 3. Mix at room temperature for at least 1 h, using a rotary mixer. 4. Collect the beads by centrifugation and remove supernatant as in step 4 of Subheading 3.5. Add 10 ml Tris/NaCl/CaCl2 buffer and mix for 10 min at RT. Spin at 300 × g for 60 s and discard supernatant, as in step 5 of Subheading 3.5. To increase bead recovery, one can perform an extra centrifugation step on the supernatant before discarding the supernatant. 5. Repeat wash procedure four times. 6. Add an equal volume of Tris/NaCl/EDTA elution buffer to beads and combine into 1–2 tubes. Mix overnight at 4°C. Next day, collect the beads by centrifugation and remove supernatant. Repeat elution by adding fresh elution buffer to the beads and mix for 15 min at room temperature. Centrifuge the beads, collect the supernatant, and combine with the first elution. 7. Desalt and concentrate the eluted material as in step 8 of Subheading 3.5. Apply sample to centrifugal filter and fill to 15 ml using 20 mM Na phosphate buffer, pH 7.4. Concentrate by centrifugation to 1 ml and repeat buffer exchange at least three times. 8. Dilute the final concentrated samples to 500 μg/ml SAP using 20 mM sodium phosphate buffer and store at 4°C.

3.7. Desalting Commercial Human SAP

Many commercial preparations of human SAP (#565190, EMDCalbiochem) or rat SAP (#1895-SA-050, R&D Systems) contain azide, detergent, or EDTA, which may interfere with many biological processes. Either a buffer control has to be used in all experiments, or a better solution is to desalt the SAP preparation by buffer exchange. 1. Wash two Amicon Ultra-0.5 ml centrifugal filters (this is a 10 kDa cut-off, so for a 130 kDa protein the loss is negligible) with 0.5 ml H2O for 5 min at 10,000 × g. Then wash twice with

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20 mM sodium phosphate pH 7.4. The liquid should pass through the filter into the bottom Eppendorf tube. Running at slower speeds and shorter time periods will prevent the filter from drying out, which is to be avoided (see Note 17). 2. Add 0.4 ml of the commercial SAP solution to the centrifugal filter. Centrifuge for 5–10 min, or until approx. 0.3 ml of the buffer has passed through the filter into the bottom Eppendorf. 3. Replace the lost liquid with 20 mM sodium phosphate pH 7.4 in the top filter and repeat four times. 4. When approximately 0.3 ml has passed through a fifth time, carefully remove the SAP solution from the top unit and replace the lost volume with 20 mM phosphate buffer. 5. Check the concentration of the solution by protein gel/ Coomassie, using albumin and the original commercial SAP preparations as protein standards. This leaves the SAP in 20 mM sodium phosphate and in this buffer SAP is stable for many weeks/months at 4°C (see Note 18). 6. SAP is very stable in its 130 kDa pentameric form and only dissociates into its individual 27 kDa protomers when exposed to temperatures above 100°C in the presence of SDS (44, 45). In the presence of 10 mM calcium, SAP is also resistant to a variety of proteases (44). However, SAP will form decamers or aggregates above 1 mM calcium, or in the presence of 140 mM NaCl (46–48). Nevertheless, in plasma and in solutions containing at least 10 mg/ml albumin SAP remains a pentamer, even in the presence of 2 mM calcium (48, 49). Therefore, SAP should be stored either in the presence of albumin or EDTA (to chelate calcium), or in 20 mM sodium phosphate (not PBS) to reduce the formation of decamers or aggregates.

4. Notes 1. This procedure is usually performed by a licensed and/or trained phlebotomist. 2. We have also used EDTA tubes to collect blood for fibrocyte studies. 3. All procedures involving blood are performed inside class II biological safety cabinets, using sterile technique and materials. Diluting the blood reduces the viscosity of the blood which helps in the following separation steps. 4. Store Ficoll-Paque at room temperature, as Ficoll-Paque stored at 4°C has a density greater than the defined density of 1.077.

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5. Care is to be taken to prevent mixing of the two liquids, as the PBMC are collected at the Ficoll-blood interface and a disturbed interface will be less efficient—read manufacturer’s leaflets for additional directions on layering blood on density gradient liquids. These volumes can be scaled up for larger blood volumes using 10 ml Ficoll in 50 ml polypropylene tubes, and adding 40 ml of blood diluted with PBS. 6. This reduces the risk of exposure to aerosols if the tubes break during centrifugation. 7. Read manufacturer’s leaflets for additional directions on removing the cell interface from density gradient liquids. 8. This process removes excess Ficoll which is toxic to cells after long exposure, along with any dead cells and platelets. These molecules and cells may also interfere in biological assays. 9. This reduces the amount of PBS need for the subsequent washes. 10. We have found that this number of steps is necessary to remove platelets. 11. These supplements are designed to promote cell survival and differentiation in the absence of serum. We have previously shown that glucose, sodium pyruvate, and insulin all affect fibrocyte differentiation (28). HEPES is an additional buffering agent, as serum has the capacity to buffer biological fluids. NEAA prolongs the viability of cells in culture and may reduce the biosynthetic burden on cells. Sodium pyruvate is an additional energy source, which is necessary for fibrocyte differentiation. ITS-3 contains insulin to promote the uptake of glucose; transferrin, an iron-transport protein necessary for cell differentiation and proliferation; selenium, an essential trace element normally found in serum; and albumin with oleic and linoleic acids, which promote cell differentiation. 12. As fibrocytes differentiate from highly adherent peripheral blood monocytes, it is an advantage to use medium that is cold to reduce the possibility of the monocytes adhering to the polypropylene tubes before being added to the cell culture plates. 13. These protocols can also be used to culture fibrocytes on glass slides for staining with antibodies (8, 10, 24). 14. Due to the inhalation hazard of methanol, ethanol can be used instead of methanol to fix the cells. Methanol and ethanol are not only effective fixatives but potent anti-biological agents. 15. Retain supernatant to check that the majority of the SAP has been removed. 16. Read manufacturer’s data sheets for the amount of CRP bound per mg of p-aminophenyl phosphoryl choline beads.

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17. Read manufacturer’s data sheets for additional information on centrifugation speeds and times. 18. This procedure should only take a few 5 min centrifuge steps at room temperature, for a buffer exchange.

Acknowledgments We would like to thank Nehemiah Cox and Jeff Crawford for critical reading of the manuscript. This work was supported by NIH grant HL083029. References 1. Bucala R, Spiegel LA, Chesney J, Hogan M, Cerami A (1994) Circulating fibrocytes define a new leukocyte subpopulation that mediates tissue repair. Mol Med 1:71–81 2. Abe R, Donnelly SC, Peng T, Bucala R, Metz CN (2001) Peripheral blood fibrocytes: differentiation pathway and migration to wound sites. J Immunol 166:7556–7562 3. Quan TE, Cowper S, Wu SP, Bockenstedt LK, Bucala R (2004) Circulating fibrocytes: collagen-secreting cells of the peripheral blood. Int J Biochem Cell Biol 36:598–606 4. Gomperts BN, Strieter RM (2007) Fibrocytes in lung disease. J Leukoc Biol 82:449–456 5. Bellini A, Mattoli S (2007) The role of the fibrocyte, a bone marrow-derived mesenchymal progenitor, in reactive and reparative fibroses. Lab Invest 87:858–870 6. Wang JF, Jiao H, Stewart TL, Shankowsky HA, Scott PG, Tredget EE (2007) Fibrocytes from burn patients regulate the activities of fibroblasts. Wound Repair Regen 15:113–121 7. Yang L, Scott PG, Giuffre J, Shankowsky HA, Ghahary A, Tredget EE (2002) Peripheral blood fibrocytes from burn patients: identification and quantification of fibrocytes in adherent cells cultured from peripheral blood mononuclear cells. Lab Invest 82:1183–1192 8. Pilling D, Buckley CD, Salmon M, Gomer RH (2003) Inhibition of fibrocyte differentiation by serum amyloid P. J Immunol 17:5537–5546 9. Pilling D, Tucker NM, Gomer RH (2006) Aggregated IgG inhibits the differentiation of human fibrocytes. J Leukoc Biol 79:1242–1251 10. Pilling D, Fan T, Huang D, Kaul B, Gomer RH (2009) Identification of markers that distinguish monocyte-derived fibrocytes from monocytes, macrophages, and fibroblasts. PLoS One 4:e7475

11. Chesney J, Bacher M, Bender A, Bucala R (1997) The peripheral blood fibrocyte is a potent antigen-presenting cell capable of priming naive T cells in situ. Proc Natl Acad Sci USA 94:6307–6312 12. Chesney J, Metz C, Stavitsky AB, Bacher M, Bucala R (1998) Regulated production of type I collagen and inflammatory cytokines by peripheral blood fibrocytes. J Immunol 160:419–425 13. Balmelli C, Ruggli N, McCullough K, Summerfield A (2005) Fibrocytes are potent stimulators of anti-virus cytotoxic T cells. J Leukoc Biol 77:923–933 14. Balmelli C, Alves MP, Steiner E, Zingg D, Peduto N, Ruggli N, Gerber H, McCullough K, Summerfield A (2007) Responsiveness of fibrocytes to toll-like receptor danger signals. Immunobiology 212:693–699 15. Yang L, Scott PG, Dodd C, Medina A, Jiao H, Shankowsky HA, Ghahary A, Tredget EE (2005) Identification of fibrocytes in postburn hypertrophic scar. Wound Repair Regen 13: 398–404 16. Schmidt M, Sun G, Stacey MA, Mori L, Mattoli S (2003) Identification of circulating fibrocytes as precursors of bronchial myofibroblasts in asthma. J Immunol 171:380–389 17. Mori L, Bellini A, Stacey MA, Schmidt M, Mattoli S (2005) Fibrocytes contribute to the myofibroblast population in wounded skin and originate from the bone marrow. Exp Cell Res 304:81–90 18. Mehrad B, Burdick MD, Zisman DA, Keane MP, Belperio JA, Strieter RM (2007) Circulating peripheral blood fibrocytes in human fibrotic interstitial lung disease. Biochem Biophys Res Commun 353:104–108 19. Sakai N, Wada T, Yokoyama H, Lipp M, Ueha S, Matsushima K, Kaneko S (2006) Secondary lym-

16 Differentiation of Circulating Monocytes into Fibroblast-Like Cells phoid tissue chemokine (SLC/CCL21)/CCR7 signaling regulates fibrocytes in renal fibrosis. Proc Natl Acad Sci USA 103:14098–14103 20. Niedermeier M, Reich B, Rodriguez Gomez M, Denzel A, Schmidbauer K, Gobel N, Talke Y, Schweda F, Mack M (2009) CD4+ T cells control the differentiation of Gr1+ monocytes into fibrocytes. Proc Natl Acad Sci USA 106:17892–17897 21. Barth PJ, Koster H, Moosdorf R (2005) CD34+ fibrocytes in normal mitral valves and myxomatous mitral valve degeneration. Pathol Res Pract 201:301–304 22. Quan TE, Bucala R (2007) Culture and analysis of circulating fibrocytes. Methods Mol Med 135:423–434 23. Shao DD, Suresh R, Vakil V, Gomer RH, Pilling D (2008) Pivotal advance: Th-1 cytokines inhibit, and Th-2 cytokines promote fibrocyte differentiation. J Leukoc Biol 83:1323–1333 24. Pilling D, Vakil V, Gomer RH (2009) Improved serum-free culture conditions for the differentiation of human and murine fibrocytes. J Immunol Methods 351:62–70 25. Phillips RJ, Burdick MD, Hong K, Lutz MA, Murray LA, Xue YY, Belperio JA, Keane MP, Strieter RM (2004) Circulating fibrocytes traffic to the lungs in response to CXCL12 and mediate fibrosis. J Clin Invest 114:438–446 26. Naik-Mathuria B, Pilling D, Crawford JR, Gay AN, Smith CW, Gomer RH, Olutoye OO (2008) Serum amyloid P inhibits dermal wound healing. Wound Repair Regen 16:266–273 27. Haudek SB, Xia Y, Huebener P, Lee JM, Carlson S, Crawford JR, Pilling D, Gomer RH, Trial J, Frangogiannis NG, Entman ML (2006) Bone marrow-derived fibroblast precursors mediate ischemic cardiomyopathy in mice. Proc Natl Acad Sci USA 103:18284–18289 28. Pilling D, Gomer RH (2007) Regulatory pathways for fibrocyte differentiation. In: Bucala R (ed) Fibrocytes-new insights into tissue repair and systemic fibroses. World Scientific, Singapore, pp 37–60 29. Pilling D, Roife D, Wang M, Ronkainen SD, Crawford JR, Travis EL, Gomer RH (2007) Reduction of bleomycin-induced pulmonary fibrosis by serum amyloid P. J Immunol 179: 4035–4044 30. Haudek SB, Trial J, Xia Y, Gupta D, Pilling D, Entman ML (2008) Fc receptor engagement mediates differentiation of cardiac fibroblast precursor cells. Proc Natl Acad Sci USA 105: 10179–10184 31. Cathcart ES, Wollheim FA, Cohen AS (1967) Plasma protein constituents of amyloid fibrils. J Immunol 99:376–385

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32. Thompson AR, Enfield DL (1978) Human plasma P component: isolation and characterization. Biochemistry 17:4304–4311 33. Binette P, Binette M, Calkins E (1974) The isolation and identification of the P-component of normal human plasma proteins. Biochem J 143:253–254 34. Pepys MB, Dash AC (1977) Isolation of amyloid P component (protein AP) from normal serum as a calcium-dependent binding protein. Lancet 1:1029–1031 35. Painter RH (1977) Evidence that C1t (amyloid P-component) is not a subcomponent of the first component of complement (C1). J Immunol 119:2203–2205 36. Pontet M, Engler R, Jayle MF (1978) One step preparation of both human C-reactive protein and CIt. FEBS Lett 88:172–175 37. de Beer FC, Pepys MB (1982) Isolation of human C-reactive protein and serum amyloid P component. J Immunol Methods 50: 17–31 38. Gomer RH, Pilling D, Kauvar L, Ellsworth S, Pissani S, Real L, Ronkainen SD, Roife D, Ma F, Davis SC (2009) A serum amyloid P-binding hydrogel speeds healing of partial thickness wounds in pigs. Wound Repair Regen 17: 397–404 39. Hind CR, Collins PM, Renn D, Cook RB, Caspi D, Baltz ML, Pepys MB (1984) Binding specificity of serum amyloid P component for the pyruvate acetal of galactose. J Exp Med 159:1058–1069 40. Schwalbe RA, Dahlback B, Coe JE, Nelsestuen GL (1992) Pentraxin family of proteins interact specifically with phosphorylcholine and/or phosphorylethanolamine. Biochemistry 31:4907–4915 41. de Beer FC, Baltz ML, Munn EA, Feinstein A, Taylor J, Bruton C, Clamp JR, Pepys MB (1982) Isolation and characterization of C-reactive protein and serum amyloid P component in the rat. Immunology 45:55–70 42. Tillett WS, Francis T (1930) Serological reactions in pneumonia with a nonprotein somatic fraction of pneumococcus. J Exp Med 52: 561–571 43. Bach BA, Gewurz H, Osmand AP (1977) C-reative protein in the rabbit: isolation, characterization and binding affinity to phosphocholine. Immunochemistry 14:215–219 44. Kinoshita CM, Gewurz AT, Siegel JN, Ying SC, Hugli TE, Coe JE, Gupta RK, Huckman R, Gewurz H (1992) A protease-sensitive site in the proposed Ca(2+)-binding region of human serum amyloid P component and other pentraxins. Protein Sci 1:700–709

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45. Pepys MB, Booth DR, Hutchinson WL, Gallimore JR, Collins PM, Hohenester E (1997) Amyloid P component. A critical review. Amyloid 4:274–295 46. Baltz ML, de Beer FC, Feinstein A, Pepys MB (1982) Calcium-dependent aggregation of human serum amyloid P component. Biochim Biophys Acta 701:229–236 47. Coker AR, Purvis A, Baker D, Pepys MB, Wood SP (2000) Molecular chaperone properties of serum amyloid P component. FEBS Lett 473:199–202

48. Hutchinson WL, Hohenester E, Pepys MB (2000) Human serum amyloid P component is a single uncomplexed pentamer in whole serum. Mol Med 6:482–493 49. Sorensen IJ, Andersen O, Nielsen EH, Svehag SE (1995) Native human serum amyloid P component is a single pentamer. Scand J Immunol 41:263–267 50. Crawford JR, Pilling D, Gomer RH (2010) Improved serum-free culture conditions for spleen-derived murine fibrocytes. J Immunol Methods 363(1):9–20

Chapter 17 Enumeration of Very Small Embryonic-Like Stem Cells in Peripheral Blood Rui Liu and Mariusz Z. Ratajczak Abstract In addition to hematopoietic stem and progenitor cells (HSPCs) also other types of stem cells (e.g., mesenchymal stem cells [MSCs], endothelial progenitor cells [EPCs], and very small embryonic-like stem cells [VSELs]) circulate under steady-state conditions at detectable levels in peripheral blood (PB), with their numbers increasing in response to stress, inflammation, tissue organ injury (e.g., heart infarct, stroke, or colitis), or mobilizing agents (e.g., granulocyte colony-stimulating factor; G-CSF). This mobilization process may be envisioned as a danger-sensing response mechanism triggered by hypoxia or mechanical- or infection-induced tissue damage that recruits into peripheral blood various types of stem cells that play a role not only in immune surveillance but also in organ/tissue regeneration. Thus, stem cells circulating in PB could be envisioned as “cellular paramedics” that are involved in immune surveillance (HSPCs) or tissue/organ rejuvenation (MSCs, EPCs, VSELs). In this chapter we focus on detection and enumeration of VSELs circulating in human PB, which are circulating in steady-state conditions or after administration of G-CSF or as a consequence of various pathological events. Key words: Hematopoietic stem cells, Very small embryonic-like stem cells, Mesenchymal stem cells, Endothelial progenitors, Mobilization

1. Introduction A small percentage of hematopoietic stem and progenitor cells (HSPCs) are continuously released from bone marrow (BM) niches into peripheral blood (PB) (1, 2). Thus, PB may be envisioned as a highway by which HSPCs can relocate between distant stem cell niches to keep the pool of BM stem cells in balance. In addition to HSPCs, some other rare stem cells (e.g., mesenchymal stem cells [MSCs], endothelial progenitor cells [EPCs], and very small

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_17, © Springer Science+Business Media, LLC 2012

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embryonic-like stem cells [VSELs]) may also appear in the PB during various stress situations (1). Both HSPCs and non-hematopoietic stem cells are mobilized into peripheral blood also during different pathological situations including heart infarct, stroke, liver injury, kidney injury, limb ischemia, colitis, and organ transplants (1–6). Enumeration of these cells may be of clinical/prognostic value (1, 5, 6). In this chapter we focus on enumeration of VSELs, MSCs, and EPCs. VSELs—These small cells identified by our team express several morphological (e.g., relatively large nuclei containing euchromatin) and molecular (e.g., expression of SSEA-1, Oct4, Nanog, Rex1) markers characteristic for embryonic stem cells (ESCs) (7, 8). The true expression of Oct4 and Nanog in BM-derived VSELs (BM-VSELs) was recently confirmed by demonstrating transcriptionally active chromatin structures of Oct4 and Nanog promoters (9). We also described a mechanism based on parent-of-origin-specific reprogramming of genomic imprinting that keeps VSELs quiescent in a dormant state in tissues. The expression of germ line markers (Oct4 and SSEA-1) and modulation of somatic imprints suggest a potential developmental similarity between VSELs and germ line-derived primordial germ cells (PGCs) (9). Mobilization of VSELs into PB was reported so far both in murine models and in human patients after heart infarct, stroke, retina damage, and acute colitis (1, 5, 6). MSCs—MSCs are a population of BM-derived adherent bone/ cartilage-forming progenitor cells. It is known that BM-adherent cells grow colonies of fibroblastic-like cells which have a high replating potential (colony-forming units of fibroblasts; CFU-F) (10). It is now widely accepted that MSCs contribute to the regeneration of mesenchymal tissues (e.g., bone, cartilage, muscle, ligament, tendon, adipose, and stroma). Because various inconsistencies have come to light in the field of MSC research, in particular if they truly represent a population of stem cells, the International Society for Cellular Therapy recently recommended avoiding the name of MSCs and changing it to multipotent mesenchymal stromal cells instead (10). Of note recently it has been demonstrated that VSELs may give rise to population of MSCs (11). EPCs—It is postulated that the BM is endowed with neoangiogenetic activity and EPCs, which is a rare and very primitive founder population of endothelial cells, may be released during stressed situations and circulate in PB (12). Furthermore, BM was also identified as a source of more differentiated circulating endothelial cells (CECs). BM-derived EPC and CEC that subsequently circulate in PB at very low levels (0.0001 and 0.01%, respectively) may play a role in the repair of damaged endothelium and contribute to postnatal neo-angiogenesis (12). While EPCs are probably progeny of PSC or perhaps direct descendants of hemangioblasts, the more differentiated CECs originate in the myeloid

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compartment from a common myeloid progenitor (CMP). The level of contribution of BM-derived cells to organ/tissue vascularization, however, still requires further study. In adult organisms, circulating stem cells (1) show a circadian rhythm in the circulation with the peak occurring early in the morning and the nadir at night; (2) are mobilized during strenuous exercise, inflammation, and tissue organ injury (e.g., heart infarct or stroke); and (3) may increase in number up to 100-fold after administration of certain drugs (3–6). This enforced translocation of HSPCs from BM into PB induced by pharmacological agents (e.g., G-CSF or AMD3100) is called “mobilization” and mobilized PB (mPB) as discussed in other chapters in this book is an easily accessible source of HSPCs for hematopoietic transplantation. In this chapter we discuss cytometry-based methods to detect and enumerate circulating in PB other non-hematopoietic cells with a special emphasis on VSELs, MSCs, and EPCs. These cells as reported are detectable in murine models of pharmacological mobilization and tissue injury (13), as well as in analogical situations in humans (1–6). Identification of these cells, in particular enumeration of circulating VSELs, requires unique gating strategies to focus on small events that are slightly smaller than red blood cells (7, 14). Furthermore, since VSELs due to the both small size and density are lost (up to 50%) during Ficoll-Paque centrifugation (15), the recommended way to preserve these cells is lysis of peripheral blood samples to preserve VSELs and all other nucleated cells (HSPCs, MSCs, EPCs) for staining and subsequent analysis.

2. Materials 2.1. Preparation of Peripheral Blood for Analysis

1. Laboratory tubes containing anticoagulant or medium supplemented with anticoagulant (see Note 1). 2. Lysing buffer (PharmaLyse, BD Biosciences). 3. RPMI 1640 medium with 2% fetal bovine serum (FBS). 4. 50 ml plastic tissue culture-grade tubes (BD Biosciences). 5. Centrifuge with 50 ml tube holders. 6. Hemocytometer to enumerate nucleated blood cells.

2.2. Staining of Total PB-Derived Nucleated Cells for Analysis

1. Medium used for staining RPMI 1640 (no serum) and RPMI 1640 with 2% FBS. 2. Flow Cytometry (Invitrogen).

Size

Calibration

Kit

microspheres

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3. 50 ml plastic tissue-grade culture tubes and 5 ml round-bottom tubes. 4. 70 and 40 μm strainer/mesh filters. 5. Centrifuge with 50 ml and 5 ml tube holders. 6. Monoclonal antibodies used for staining human HSCs, VSELs, MSCs, and EPCs include mouse monoclonal antibodies against human epitopes, predominantly directly conjugated with fluorochromes—are listed in Tables 1, 2, and 3. 7. Flow cytometer.

Table 1 Antibodies employed in staining for identification and sorting of human PB-derived HSCs and VSELs by flow cytometry Antibody

Clone

Fluorochrome

Vendor

Anti-CD2

RPA-2.10

FITC

BD Biosciences

Anti-CD3

UCHT1

FITC

BD Biosciences

Anti-CD14

M5E2

FITC

BD Biosciences

Anti-CD16

3G8

FITC

BD Biosciences

Anti-CD19

HIB19

FITC

BD Biosciences

Anti-CD24

ML5

FITC

BD Biosciences

Anti-CD56

NCAM16.2

FITC

BD Biosciences

Anti-CD66b

G10F5

FITC

BD Biosciences

Anti-CD235a

GA-R2

FITC

BD Biosciences

Anti-CD45

HI30

PE-Cy7

BD Biosciences

Anti-AC133

AC133

APC

Miltenyi Biotec

Anti-CD34

581/CD34

APC

BD Biosciences

Anti-CD184

12G5

APC

BD Biosciences

Isotype controls Mouse IgG1, κ Mouse IgG2a, κ Mouse IgG2b, κ Mouse IgG1, κ Mouse IgG1 Mouse IgG1, κ Mouse IgG2a, κ

MOPC-21 G155-178 27-35 MOPC-21 IS5-21F5 MOPC-21 G155-178

FITC FITC FITC PE-Cy7 APC APC APC

BD Biosciences BD Biosciences BD Biosciences BD Biosciences Miltenyi Biotec BD Biosciences BD Biosciences

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Table 2 Antibodies employed in staining for identification and sorting of human PB-derived MSCs by flow cytometry Antibody

Clone

Fluorochrome

Vendor

Anti-CD45

HI30

FITC

BD Biosciences

Anti-CD90

5E10

PE

BioLegend

Anti-CD34

4H11

PE-Cy7

eBioscience

Anti-CD105

43A3

APC

BioLegend

Anti-CD29

TS2/16

APC-Cy7

BioLegend

Anti-CD31

WM59

FITC

BioLegend

Anti-CD105

43A3

PE

BioLegend

Anti-CD45

HI30

PE-Cy7

BD Biosciences

Anti-CD73

AD2

APC

eBioscience

MOPC-21 MOPC-21 MOPC-21 MOPC-21 MOPC-21 MOPC-21 MOPC-21

FITC PE PE-Cy7 APC APC-Cy7 FITC PE PE-Cy7 APC

BD Biosciences BioLegend eBioscience BioLegend BioLegend BioLegend BioLegend BD Biosciences eBioscience

Isotype controls Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ Mouse IgG1, κ

3. Methods Steady-state conditions circulating PB in addition to HSPCs contain also some other types of stem cells that circulate in peripheral blood. Overall a number of all these cells increase during stress situations, infections, strenuous exercise, tissue organ damage, and pharmacological mobilization (1–6). Interestingly, in particular in cytokine-/growth factor-induced mobilization some selectivity of mobilized and then circulating in PB cells was observed—depending on combination of mobilizing agents (16). Accordingly, vascular endothelial growth factor (VEGF) pretreatment enhanced EPC mobilization and mobilization of MSCs was preferentially induced in mice when the CXCR4 antagonist (AMD3100) was administered

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Table 3 Antibodies employed in staining for identification and sorting of human PB-derived EPCs by flow cytometry Antibody

Clone

Fluorochrome

Vendor

Anti-CD14

M5E2

FITC

BD Biosciences

Anti-VEGF R2(KDR)

89106

PE

R&D Systems

Anti-CD34

4H11

PE-Cy7

eBioscience

Anti-AC133

AC133

APC

Miltenyi Biotec

Anti-CD31

WM59

Alex 647

BioLegend

Anti-CD105

43A3

APC

BioLegend

Isotype controls Mouse IgG2a, κ Mouse IgG1 Mouse IgG1, κ Mouse IgG1 Mouse IgG1, κ Mouse IgG1, κ

G155-178 11711 IS5-21F5 MOPC-21 MOPC-21

FITC PE PE-Cy7 APC Alex Flour 647 APC

BD Biosciences R&D Systems eBioscience Miltenyi Biotec BioLegend BioLegend

to mice pretreated with VEGF, but not G-CSF (16). We have observed that the number of VSELs increases in PB after we employ combination of G-CSF + CXCR4 antagonist (AMD3100 or T140) (17). Furthermore, mounting evidence accumulates that more or less primitive subsets of HSPCs are mobilized in response to G-CSF, AMD3100 Gro-β chemokines alone, or in combination (4). In protocols described below human PB-derived samples are collected from the patients into tubes with anticoagulant. To avoid the loss of small cells (e.g., VSELs) during separation of cells on Ficoll-Paque gradient we remove red blood cells by employing lysing buffer. Total nucleated cells are subsequently stained by using antibodies listed in Tables 1, 2, and 3—based on population of cells we would like to identify (HSPCs, VSELs, MSCs, or EPC). For isolation of VSELs we may use size-predefined beads, to define a sorting region containing small objects (2–10 μm), as indicated on the dot plot presenting FSC and SSC parameters of analyzed objects (Fig. 1—region R1). This region contains mostly cellular debris but also includes rare nuclear cellular objects (8, 15, 18). 3.1. Isolation of Total PB-Derived Nucleated Cells

1. Collect patient PB into tubes containing anticoagulant (see Notes 1 and 2). 2. Distribute PB sample into 50 ml plastic tubes in amount of 10 ml of PB per tube, fill the tubes by adding 40 ml 1× PBS,

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Fig. 1. Gating strategy for sorting of human PB-VSELs by FACS. PB-VSELs were isolated from fraction of human PB TNCs by FACs by employing the following gating criteria. (a) All events ranging from 2 μm are included in gate R1 after comparison with six differently sized bead particles with standard diameters of 1, 2, 4, 6, 10, and 15 μm. (b) PB-derived TNCs are visualized on a dot plot based on FSC vs. SSC signals. (d) Cells from region R1 are further analyzed for CD133 and Lin expression: Lin−/CD133+ events are included in region R2. (c) The Lin−/CD133+ population from region R2 is subsequently analyzed based on CD45 antigen expression and CD45− and CD45+ subpopulations visualized on dot plot, i.e., CD133+/ Lin−/CD45− (VSELs: region R3) and CD133+/Lin−/CD45+ (HSCs: region R4).

and centrifuge samples for 10 min at 500 × g at room temperature (RT). 3. Discard supernatant and remove the red blood cells (RBCs) by employing BD Pharm Lyse—lysing buffer (see Note 3). Add 40 ml of 1× lysing solution to each tube. Immediately after adding the lysing solution begin gently vortexing each tube. Incubate at room temperature for 10 min and protect from light. 4. Centrifuge cells for 10 min at 500 × g at RT and remove supernatant carefully. 5. Repeat lysis of RBCs by resuspending the pellet of cells with 20 ml BD Pharm Lyse buffer. Incubate cells for 10 min in RT and then spin the samples for 10 min at 500 × g at 4°C.

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6. Discard supernatant, resuspend the remaining pellet in 50 ml of RPMI 1640 media with 2% FBS, transfer cell suspension to a new 50 ml tube passing through a 70-μm strainer/mesh filter to remove cellular clumps, and then spin down for 10 min at 500 × g at 4°C. 7. Resuspend cells in 1 ml of RPMI 1640 media with 2% FBS, and count total nucleated cells (TNCs) with hemocytometer. 8. Separate the cell suspension into tubes (5 ml round-bottom tubes). The number of cells in each tube should be kept around 3–5 million/tube. Stain cells as described below. 3.2. Staining of PB-Derived TNCs for Flow Cytometric Analysis

1. Stain PB-derived TNCs in RPMI 1640 media supplemented with 2% FBS with antibodies (for list of required antibodies see Tables 1, 2, and 3). Staining should be performed according to recommendations provided by vendor in 5 ml round-bottom tubes kept for 30 min on ice (see Note 4). 2. Wash all samples by adding 3 ml of RPMI 1640 medium with 2% FBS and centrifuge tubes for 10 min at 500 × g at 4°C. 3. Resuspend cells for analysis in 0.5 ml RPMI 1640 medium with 2% FBS. Keep samples on ice until analysis by FACS. 4. Cells to be sorted have to be resuspended in RPMI 1640 medium supplemented with 2% FBS and transferred to new tubes (5 ml round-bottom tubes) after filtration through a 40 μm strainer/mesh filter to remove cell clumps. Adjust volume of cell suspension to 4/tube and keep cells on ice until sorting.

3.3. Setting Up Instrument for FACS Analysis

1. Set up the forward- and side-scatter parameters (FSC and SSC, respectively) in logarithmic or linear scale and adjust the threshold on FSC parameter. 2. Run the mixture of the size-predefined beads (size calibration beads with standard diameters of 1, 2, 4, 6, 10, and 15 μm) and adjust the threshold of cytometer to be able to include for sorting all objects that are bigger/equal 2 μm. 3. Set up minimal threshold to be able to see 2 μm beads on FSC vs. SSC dot plot. 4. Set up the gate which will include all objects larger than 2 μm on dot plot showing objects according to their FSC and SSC parameters. 5. Run the stained samples and adjust the gate to include agranular objects larger in size than 2 μm. 6. Perform compensation calculations and prepare the logical gating strategy resulting in identification. Analyze human VSELs, MSCs, and EPCs by FACS (as shown in Figs. 2, 3, and 4) or isolate CD133+/Lin−/CD45− PB-VSELs by MoFlo (as shown in Fig. 1).

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Fig. 2. Gating strategy for sorting of human PB-VSELs derived by FACS. PB-VSELs were isolated from fraction of human PB TNCs by FACs. (a) PB-derived TNCs are visualized by dot plot based on FSC vs. SSC signals. The TNC events are shown in region P3. (b–f) Cells from region P1 are further analyzed for hematopoietic lineages maker expression and all the Lin− events are included in region P2. (c) The Lin− population from region P2 is subsequently analyzed based on CD133 and CD45 antigen expression and two populations of CD133+ cells are distinguished based on CD45 expression, i.e., Lin−/ CD45−/CD133+ (VSELs: region Q2) and Lin−/CD45+/CD133+ (HSCs: region Q4). (e) The Lin− population from region P2 is subsequently analyzed based on CD34 and CD45 antigen expression and two populations of CD34+ cells are distinguished based on CD45 expression, i.e., Lin−/CD45−/CD34+ (VSELs: region P5) and Lin−/CD45+/CD34+ (HSCs: region P4). (g) The Lin− population from region P2 is subsequently analyzed based on CXCR4 and CD45 antigen expression and two populations of CXCR4+ cells are distinguished based on CD45 expression, i.e., Lin−/CD45−/CXCR4+ (VSELs: region P5) and Lin−/ CD45+/CXCR4+ (HSCs: region P4).

7. The phenotypes of VSELs, MSCs, and EPCs are described as below. 3.4. Identification of Circulating Stem Cell Populations

1. HSPCs (Fig. 2) are identified according to cell surface markers as (1) Lin/CD45+/CD133+, (2) Lin−/CD45+/CD34+, and (3) Lin−/CD45+/CXCR4+ cells. Lineage markers include CD2, CD3, CD14, CD16, CD19, CD24, CD56, CD66b, and CD235 (see Notes 5 and 6). 2. VSELs (Fig. 2) are identified and enumerated as (1) Lin−/CD45−/ CD133+, (2) Lin−/CD45−/CD34+, and (3) Lin−/CD45−/ CXCR4+. Lineage markers include CD2, CD3, CD14, CD16, CD19, CD24, CD56, CD66b, and CD235 (see Note 7).

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Fig. 3. Gating strategy for sorting human PB-derived MSCs by FACS. PB-MSCs were isolated from fraction of human PB TNCs by FACs. (a) PB-derived TNCs are visualized by dot plot based on FSC vs. SSC signals. All the TNC events are included in region P3. (b) Cells from region P1 are further analyzed for CD45− events and are included in region P2. (c) The CD45− population from region P2 is subsequently analyzed based on CD34 and CD105 antigen expression and CD105+/CD34− population is shown in region P4. (d) The CD105+/CD34− population from P4 is subsequently analyzed based on CD90 and CD29 antigen expression and CD90+/CD29+ cells are distinguished by dot plot, i.e., CD45−/CD90+/CD34−/CD105+/CD29+ (MSCs: region P5). (e) Cells from region P1 are further analyzed for MSCs and CD31− events are included in region P2. (f) The CD31− population from region P2 is subsequently analyzed based on CD73 and CD45 antigen expression and CD45−/CD73+ population is included in region P4. (g) The CD45−/CD73+ population from P4 is subsequently analyzed based on CD105 antigen expression and CD105+ events are visualized by dot plot, i.e., CD31−/CD105+/CD45−/CD73+ (MSCs: region P5).

3. MSCs (Fig. 3) are identified in PB samples as population of (1) or CD31−/ CD45−/CD90+/CD34−/CD105+/CD29+ CD105+/CD45−/CD73+ cells. 4. EPCs (Fig. 4) are identified as (1) KDR+/CD34+/AC133+, (2) CD14+/KDR+/CD34+/CD31+, or CD14+/KDR+/CD34+/ CD105+ cells. 3.5. Quantification of Target Cells in PB

1. Calculate TNCs’ number in 1 μl of peripheral blood according to formula—TNCs’ number in 1 μl PB = amount of TNCs (by counting)/volume of PB (μl). 2. Calculate absolute numbers of target cells in 1 μl of PB by employing formula—absolute numbers of target cells in 1 μl of PB = (TNCs’ number/μl PB × identified target cells’ number)/ amount of TNCs recorded in each sample by FACS.

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Fig. 4. Gating strategy for sorting human PB-derived EPCs by FACS. PB-EPCs were isolated from fraction of human PB TNCs by FACs. (a) PB-derived TNCs are visualized by dot plot based on FSC vs. SSC signals. The TNC events are included in region P3. (b) Cells from region P1 are further analyzed for CD34+/CD133+ events and are included in region P2. (c) The CD34+/ CD133+ population from region P2 is subsequently analyzed based on KDR antigen expression and KDR+ population is distinguished based on CD34+/CD133+ expression, i.e., KDR+/CD34+/CD133+ (EPCs: region P4). (d) Cells from region P1 are further analyzed for EPCs and CD34+/CD31+ events are included in region P2. (e) The CD34+/CD31+ population from region P2 is subsequently analyzed based on KDR and CD14 antigen expression, and KDR+/CD14+ population is identified by dot plot, i.e., CD14+/KDR+/CD34+/CD31+ (EPCs: region P4). (f) Cells from region P1 are further analyzed for EPCs and CD34+/ CD105+ events and are shown in region P2. (g) The CD34+/CD105+ population from region P2 is subsequently analyzed based on KDR and CD14 antigen expression and KDR+/CD14+ population is enumerated by dot plot as CD14+/KDR+/CD34+/ CD105+ cells (EPCs: region P4).

3.6. Sorting of Cells

By employing the above-described strategies HSCs, VSELs, MSCs, and EPCs can be sorted from BM, UCB, or PB (see Note 8).

4. Notes 1. Since VSEL, MSCs, and EPCs are very rare cells in PB, we require minimum 10 ml of PB to enumerate these cells—in particular in steady-state conditions. 2. Similar protocol may be employed for enumeration of umbilical cord blood (UCB)-derived or human bone marrow

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(BM)-derived cells. Since both UCB and BM contain more stem cells, the volume of the harvested samples may be decreased to ~5 ml. 3. To remove efficient RBC, the 1× lysing solution should be warmed to room temperature. Lysing buffer prewarmed to room temperature works much better than the cold one. After adding the lysing solution, the cells should be very well resuspended. 4. Remember to prepare single-color-stained samples to prepare proper compensation profile for flow cytometric analysis/sorting as well as samples stained with isotype controls only (for isotype control antibodies—see Tables 1, 2, and 3). 5. In order to obtain valuable data record at least one million TNC events from each sample. 6. The preferable concentration of cells resuspended for analysis/ sort should be between 10 and 15 × 106/ml. 7. While analyzing VSELs and HSCs, the cells are gated in extended to the left lymphocytic cells’ area. For analysis of MSCs and EPCs, add also cells in the monocytic gate as shown in Note 4. 8. During sorting, do not exceed the speed of 20,000 events/s to keep the recovery and purity of sorted cells high. Use typical high-purity sorting mode (e.g., purify 1 drop for MoFlo cell sorter).

Acknowledgments This work was supported by NIH R01 CA106281-01, NIH R01 DK074720, EU structural funds, Innovative Economy Operational Program POIG.01.01.01-00-109/09-01, and the Henry M. and Stella M. Hoenig Endowment to MZR. References 1. Ratajczak MZ, Kim CW, Wojakowski W, Janowska-Wieczorek A, Kucia M, Ratajczak J (2010) Innate immunity as orchestrator of stem cell mobilization. Leukemia 24:1667–1675 2. Lévesque J-P, Helwani FM, Winkler IG (2010) The endosteal ‘osteoblastic’ niche and its role in hematopoietic stem cell homing and mobilization. Leukemia 24:1979–1992 3. Cottler-Fox MH, Lapidot T, Petit I, Kollet O, DiPersio JF, Link D, Devine S (2003) Stem cell mobilization. Hematol Am Soc Hematol Educ Program 419–437

4. Pelus LM, Bian H, King AG, Fukuda S (2004) Neutrophil-derived MMP-9 mediates synergistic mobilization of hematopoietic stem and progenitor cells by the combination of G-CSF and the chemokines GRObeta/ CXCL2 and GRObetaT/CXCL2delta4. Blood 103:110–119 5. Wojakowski W, Tendera M, Kucia M, ZubaSurma E, Paczkowska E, Ciosek J, Hałasa M, Król M, Kazmierski M, Buszman P, Ochała A, Ratajczak J, Machaliński B, Ratajczak MZ (2009) Mobilization of bone marrow-derived

17 Enumeration of Very Small Embryonic-Like Stem Cells in Peripheral Blood Oct-4+ SSEA-4+ very small embryonic-like stem cells in patients with acute myocardial infarction. J Am Coll Cardiol 53:1–9 6. Paczkowska E, Kucia M, Koziarska D, Halasa M, Safranow K, Masiuk M, Arbicka A, Nowik M, Nowacki P, Ratajczak MZ, Machalinski B (2009) Clinical evidence that very small embryonic-like stem cells are mobilized into peripheral blood in patients after stroke. Stroke 40:1237–1244 7. Kucia M, Reca R, Campbell FR, Zuba-Surma E, Majka M, Ratajczak J, Ratajczak MZ (2006) A population of very small embryonic-like (VSEL) CXCR4+SSEA-1+Oct-4+ stem cells identified in adult bone marrow. Leukemia 20:857–869 8. Zuba-Surma EK, Ratajczak MZ (2010) Overview of very small embryonic-like stem cells (VSELs) and methodology of their identification and isolation by flow cytometric methods. Curr Protoc Cytom. Chapter 9, Unit 9.29 9. Shin DM, Liu R, Klich I, Wu W, Ratajczak J, Kucia M, Ratajczak MZ (2010) Molecular signature of adult bone marrow-purified very small embryonic-like stem cells supports their developmental epiblast/germ line origin. Leukemia 24:1450–1461 10. Horwitz EM, Le Blanc K, Dominici M, Mueller I, Slaper-Cortenbach I, Marini FC et al (2005) Clarification of the nomenclature for MSC: The International Society for Cellular Therapy position statement. Cytotherapy 7:393–395 11. Taichman RS, Wang Z, Shiozawa Y, Jung J, Song J, Balduino A, Wang J, Patel LR, Havens AM, Kucia M, Ratajczak MZ, Krebsbach PH (2010) Prospective identification and skeletal localization of cells capable of multilineage differentiation in vivo. Stem Cells Dev 19(10):1557–1570

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12. Wu H, Chen H, Hu PC (2007) Circulating endothelial cells and endothelial progenitors as surrogate biomarkers in vascular dysfunction. Clin Lab 53:285–295 13. Kucia M, Zhang YP, Reca R, Wysoczynski M, Machalinski B, Majka M, Ildstad ST, Ratajczak J, Shields CB, Ratajczak MZ (2006) Cells enriched in markers of neural tissue-committed stem cells reside in the bone marrow and are mobilized into the peripheral blood following stroke. Leukemia 20:18–28 14. Zuba-Surma EK, Kucia M, Wu W, Klich I Jr, Lillard JW, Ratajczak J, Ratajczak MZ (2008) Very small embryonic-like stem cells are present in adult murine organs: ImageStream-based morphological analysis and distribution studies. Cytometry Part A 73A:1116–1127 15. Zuba-Surma EK, Klich I, Greco N, Laughlin MJ, Ratajczak J, Ratajczak MZ (2010) Optimization of isolation and further characterization of umbilical-cord-blood-derived very small embryonic/epiblast-like stem cells (VSELs). Eur J Haematol 84:34–46 16. Pitchford SC, Furze RC, Jones CP, Wengner AM, Rankin SM (2009) Differential mobilization of subsets of progenitor cells from the bone marrow. Cell Stem Cell 4(1):62–72 17. Kucia MJ, Wysoczynski M, Wu W, Zuba-Surma EK, Ratajczak J, Ratajczak MZ (2008) Evidence that very small embryonic-like stem cells are mobilized into peripheral blood. Stem Cells 26:2083–2092 18. Zuba-Surma EK, Abdel-Latif A, Case J, Tiwari S, Hunt G, Kucia M, Vincent RJ, Ranjan S, Ratajczak MZ, Srour EF, Bolli R, Dawn B (2006) Sca-1 expression is associated with decreased cardiomyogenic differentiation potential of skeletal muscle-derived adult primitive cells. J Mol Cell Cardiol 41:650–660

Chapter 18 Generation of a Vascular Niche for Studying Stem Cell Homeostasis Jason M. Butler and Shahin Rafii Abstract Emerging evidence indicates that endothelial cells (ECs) not only form the passive building blocks of blood vessels that deliver oxygen and nutrients, but also instructively participate in organ regeneration and tumorigenesis by producing tissue-specific angiocrine factors. Due to a lack of unbiased, functional angiogenic models, the role of ECs in the homeostasis of tissue-specific stem cells and propagation of malignant cells is unknown. We established a means to maintain primary EC cultures by introducing phospho-ser473 Akt, enabling their survival for weeks under serum-/cytokine-free conditions. This lentiviral-based system maintains the angiogenic repertoire without immortalization and increased tumorigenic potential. Using our novel cytokine-/serum-free in vitro EC-based culture system, we have shown that ECs are endowed with the capacity to expand and maintain bona fide hematopoietic stem cells (HSCs) and survival of leukemic cells. This unbiased system described here can serve as a platform to identify EC-derived growth and to model treatment of a wide variety of hematological and malignant conditions. Key words: Endothelial cells (ECs), Angiocrine factors, Hematopoietic stem cells (HSCs), Human umbilical vein endothelial cells (HUVECs), Adenoviral E4 open reading frame 1 (E4ORF1)

1. Introduction The bone marrow (BM) contains specialized microdomains that constitute the hematopoietic stem cell (HSC) niches. In 1978, Schofield (1) was the first to suggest that HSCs may be supported by their microenvironment by niche cells providing regulatory signals. These microdomains comprise a wide variety of cells that support hematopoiesis through the paracrine expression of released and membrane-bound factors that help regulate the balance of quiescence, self-renewal, survival, and differentiation of HSC.

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3_18, © Springer Science+Business Media, LLC 2012

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The BM microenvironment is made up of multiple hematopoietic niches that include the endosteal, vascular, and perivascular niches. It has been suggested that the endosteal cells might maintain quiescence of the HSCs (2–5). However, since sinusoidal endothelial cells (SECs) comprise the majority of the surface area within the BM microenvironment, and can produce paracrine factors supporting HSC homeostasis, it is likely that SECs play a seminal role in the creation of the hematopoietic niches. Indeed, it has been demonstrated that the phenotypically marked long-term HSCs (LT-HSCs) reside adjacent to SECs in the BM (6, 7). It has also been suggested that perivascular reticular cells also secrete pro-hematopoietic growth factors (8, 9). The complexity of the BM microenvironment raises the important question whether all the various cell types that are within the BM make up the hematopoietic niche or that each of these cell types serves a different function, suggesting that the BM microenvironment comprises multiple hematopoietic niches. Hematopoietic regeneration following myeloablative injury is a highly complex process that requires the self-renewal and differentiation of the HSCs. The precise mechanisms that regulate the balance between self-renewal and lineage-specific differentiation are not fully understood. Studies to determine the role of the endothelial cells (ECs) in homeostasis and expansion of LT-HSCs have been hampered by a lack of functional angiogenic models. Emerging evidence suggests that ECs are not simply passive conduits for delivering oxygen, nutrients, or waste disposal. By contrast, ECs could be conceived as specialized organ-specific vascular niches that upon proper activation could instructively support maintenance and reconstitution of normal and malignant stem/ progenitor cells by production of stem cell active paracrine factors, which we define as angiocrine factors. To circumvent the lack of angiogenic models, we have developed a novel in vitro system demonstrating that ECs are essential for the expansion and maintenance of both mouse HSCs capable of engrafting lethally irradiated recipients. We established EC cultures by introducing the adenoviral E4ORF1 gene into primary human endothelial cells (E4ORF1+ ECs) enabling their survival for weeks under serum-/cytokine-free conditions, while maintaining their angiogenic properties. Using our novel EC-based coculture system we have demonstrated that coculture of mouse cKit+Lineage−Sca1+ with E4ORF1+ ECs in serum-/cytokine-free cultures resulted in the expansion of LT-HSCs beyond 14 days in a contact-dependent manner. The expanded HSCs were able to engraft lethally irradiated primary and secondary recipients (10). These data suggest that E4ORF1+ ECs provide an effective model to identify the permissive and instructive factors that modulate self-renewal and lineage-specific differentiation of LT-HSC and also elucidate into the mechanisms in which hematopoietic cells interact with endothelium.

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2. Materials

2.1. Cell Culture

1. Gelatin powder (J.T. Baker, Phillipsburg, NJ), 0.2% in sterile water and stored at 4°C. 2. Endothelial growth medium (EGM): Medium 199/DBSS (Thermo Scientific, Logan, UT) supplemented with 20% fetal bovine serum, 50 mg/l of endothelial mitogen (Biomedical Technologies Inc., Stoughton, MA) reconstituted in Medium 199/DBSS, 50 mg/500 ml of heparin, 1% antibiotic–antimycotic solution, and 1% L-glutamine. 3. Hyclone Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 20% fetal bovine serum. 4. X-Vivo 20 (Lonza, Walkersville, MO) supplemented with 50 ng/ml of mouse stem cell factor (Peprotech, Rockyhill, NJ), 1% antibiotic–antimycotic solution, and 1% L-glutamine. 5. Trypsin/EDTA 1×. 6. HEPES: 1 M HEPES made in sterile water, pH adjusted to 7.55 at room temperature. 7. HEPES-buffered saline stock (HBSS) for endothelial cell isolation. Dissolve in 1 l of sterile water 88.0 g NaCl, 3.30 g KCl, 26.18 g HEPES, 22.0 g glucose. 8. Collagenase Type 1 reconstituted in HBSS. 9. EDTA reconstituted in HBSS. 10. Bovine serum albumin. 11. 500 ml filter system.

2.2. Isolation of Human Umbilical Cord Vein

1. Gauze pad. 2. 150 ml beaker. 3. Nylon ties. 4. Cannulas. 5. Hemostat. 6. Syringes.

2.3. Virus Production and Infection

1. DMEM supplemented with 20% fetal bovine serum. 2. 293 T cells (American Type Culture Collection (ATCC), Manassas, VA). 3. Solution A (Lentivirus Mix added to deionized water containing 2 M CaCl2): pCCL.PGK gene of interest plasmid: 10–15 mg. VSV-G envelope plasmid: 3 mg.

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pMDL/pRRE packaging plasmid: 5 mg. pRSV-REV packaging plasmid: 2.5 mg. 4. Solution B (2× HEPES balanced salt solution): 281 mM NaCl, 100 mM HEPES, 1.5 mM Na2HPO4. Adjust pH to 7.08–7.12. Store at −20°C or at 4°C for up to ten days. 2.4. Antibodies for Flow Cytometry and Magnetic Separation

1. Hematopoietic Lineage Positive Cocktail v450-conjugated: CD41a (clone MWReg30) (BD Biosciences, San Jose, CA). CD11b (clone M1/70) (BD Biosciences, San Jose, CA). CD3 (clone 17A2) (BD Biosciences, San Jose, CA). Gr-1 (clone RB6-8 C5) (BD Biosciences, San Jose, CA). B220 (clone RA3-6B2) (BD Biosciences, San Jose, CA). Ter119 (clone TER-119) (BD Biosciences, San Jose, CA). 2. Sca-1 PECy7-conjugated (BD Biosciences, San Jose, CA). 3. cKit APC-conjugated (BD Biosciences, San Jose, CA). 4. CD34 FITC-conjugated (BD Biosciences, San Jose, CA). 5. Mouse Hematopoietic Lineage Depletion Kit (Miltenyi Biotech, Auburn, CA). 6. Human Endothelial Cell CD31 Depletion Kit (Miltenyi Biotech, Auburn, CA).

3. Methods Despite major advances in cultivating various cellular components of the BM niches, very few unbiased serum-free coculture systems have been developed to study the mechanisms by which various BM niche cells support the balanced proliferation and differentiation of the authentic repopulating HSCs. Cultivation studies attempted to identify soluble or membrane-bound paracrine factors expressed by BM stromal cell that promoted the expansion of bona fide repopulating HSC. Initial studies, using serum-free cultures, demonstrated that a combination of soluble growth factors, introduction of transcription factors, and/or up-regulation of intracellular signaling pathways could augment expansion of repopulating LT-HSCs (11–16). However, the repopulating ability of the cultured HSCs is gradually decreased, suggesting that the interaction between the LT-HSCs and their microenvironment may be essential in maintaining the balance between self-renewal and differentiation. Many attempts have been made to use immortalized BM stromal cell lines, but with limited success. Supplementation with serum and multiple cytokines needed to expand hematopoietic cells could only support the short-term expansion of HSCs (17). BM stromal

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cells used in these studies comprise heterogeneous populations of cells, including fibroblasts, mesenchymal cells, and a small population of ECs. As culturing of ECs has been cumbersome, the contribution of the BM vascular cells to HSC self-renewal has not been fully elucidated. Deciphering the capacity of ECs to support the expansion of LT-HSCs has been hampered by a lack of serum- and cytokine-free coculture models in which ECs could be cultivated in vitro or selectively targeted in vivo. Withdrawal of cultured ECs from angiogenic factors, such as VEGF-A, FGF-2, IGF, and EGF or serum, for a few hours results in apoptosis, thereby making it impossible to study the role of ECs in sustaining self-renewal of LT-HSC. Furthermore, previous studies in which ECs were cocultured with HSCs in the presence of serum and angiogenic factors have shown only marginal enhancement of hematopoietic progenitor cell expansion (18). Failure of these studies to show a definite role for ECs in supporting HSC could be due to suboptimal serumand growth factor-dependent culture conditions, resulting in depletion of ECs and loss of their angiogenic activity. Moreover, the presence of exogenously added EC growth factors could have also promoted expansion of stem and progenitor cells independent of a true contribution from the ECs. Ultimately, there is a need to generate a durable and reproducible EC culture system that does not require the addition of exogenous growth factors and/or serum. This unbiased culture system would be an ideal platform to study the role of ECs in the regulation of stem cell biology and also to decipher the mechanisms by which organ-specific ECs modulate organ regeneration. 3.1. Gelatin Coating on Tissue Culture Plates

1. Warm up 0.2% gelatin. 2. Add 3 ml per T25. 3. Let stand overnight at 4°C. 4. Remove gelatin right before using (see Note 1).

3.2. Isolation of Human Umbilical Vein Endothelial Cells from Umbilical Cords

1. Thaw 10 ml aliquot of 0.1% collagenase. 2. Place cord in sterile container, containing sterile 1× HEPES buffer, in 37°C water bath. 3. Prepare laminar flow tissue culture hood for cord: (a) Turn off UV light and turn on fluorescent light. (b) Turn on blower. (c) Wipe hood down with 70% ethanol. 4. Remove sterile tray from sterile bag under a laminar flow tissue culture hood. The contents of the sterile bag are as follows: Three sterile gauze pads. One 150 ml beaker.

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Three nylon ties. Two cannulas (one with rubber tubing attached). One hemostat. One aluminum triangle. Two 50 ml syringes. One 30 ml syringe. One 10 ml syringe. 5. Place sterile 10 gauge needle and sterile #10 surgical blade on sterile tray in the hood. 6. Place sterile mat and sterile gloves in the hood. 7. Place thawed collagenase in rack in the hood. 8. Don sterile gloves and remove 10 ml syringe tip and attach 18 gauge needle. 9. Place syringe back into tray and discard tube. Pour 150 ml sterile buffer into beaker. 10. Place all material on sterile mat. Fill the two 50 ml and one 30 ml syringe with buffer (step 9). 11. Remove cord from cup with hemostat. Wipe clean with sterile gauzes. 12. Inspect the cord for clamp marks. If present, cut just below and above the marks. 13. Cannulate one end of the cord with steel cannulas (use the cannulas without the rubber tubing). Hold in place with nylon ties. 14. Rinse cord with 1× HEPES buffer (one 50 ml syringe). 15. Repeat steps 13 and 14 for other end of the cord. 16. Fill cord with collagenase and as collagenase starts to come out of rubber tubing, clamp rubber tubing with hemostat. Continue filing cord with collagenase until it becomes slightly distended. 17. Cover rubber tubing with aluminum foil. 18. Incubate for 10 min in 37°C buffer bath (sterile beaker with PBS). 19. During incubation, add 10 ml of EGM to a 50 ml conical tube. 20. With 30 ml 1× HEPES buffer, flush collagenase–cell mixture from the cord into the tube containing the 10 ml of media from step 19. 21. Spin for 10 min at 1,100 × g. 22. Decant supernatant and add EGM for plating (4 ml per one gelatin-coated T25) (see Note 2).

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1. Remove old EGM, add HEPES buffer, wash over cells gently, and remove buffer. 2. Add 1:1 mixture of 0.05% collagenase and 0.02% EDTA–0.5% BSA. 3. Incubate at 37°C for 5 min. 4. Add equal parts EGM, rinse off cell-side of flask several times, remove cell suspension, and put into appropriate tube (size dependent on volume). 5. Spin at 100 × g for 5 min and remove supernatant. 6. Resuspend cells in 12 ml of EGM and put 2 ml of cell:EGM mixture into each well of a gelatin-coated 6-well tissue culture plate (see Note 3).

3.4. Generation of E4ORF1 Lentivirus

1. Plate 293T cells on gelatin-coated 10 cm tissue culture dishes with DMEM 10% FCS media. 2. Once 50–80% confluent, replace culture media with 10 ml of fresh culture media. 3. Prepare transfection solutions A and B. 4. Carefully and slowly vortex Solution B while adding Solution A. 5. Gently vortex transfection solution (A + B) and add the solution dropwise to the 10 cm dish of 293T cells (see Note 4). 6. Gently move 10 cm dish back and forth to distribute the transfection solution evenly. Allow 8–16 h for incubation of transfection solution and 293T cells. All the following steps should be performed in a laminar flow tissue culture hood dedicated to virus laboratory work. 7. Remove transfection solution and add 10 ml of fresh culture media per 10 cm dish. 8. 24 h later, collect virus media in a 15- or 50 ml tube and store at 4°C. 9. Add 10 ml of fresh culture media per 10 cm dish. 10. 24 h later, collect virus media and combine with the virus media collected in step 8. 11. Centrifuge virus media at 100 × g for 3 min. 12. Pass virus media through a 0.45 mm filter, aliquot in 1 ml tubes, and store in −80°C until use (see Note 5).

3.5. Transduction of HUVECs with E4ORF1 Lentivirus

1. Plate human umbilical vein endothelial cells (HUVECs) on gelatin-coated 6-well tissue culture plates in 2 ml of EGM. 2. Once 90% confluent, replace EGM with 1 ml of fresh EGM. 3. Add 1.5 ml of 4 mg/ml polybrene.

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4. Add 1 ml of E4ORF1 virus stock 3 wells of the 6-well plate and 1 ml of GFP virus stock (virus made the same time as E4ORF1) to the remaining 3 wells. 5. 18 h later, remove virus medium and add 1 ml of fresh EGM to all wells. 6. 48 h later, remove virus medium and add 1 ml of fresh EGM to all wells. 7. Check for GFP expression. Once confirmed, remove EGM and replace with serum-free culture media (X-Vivo 20). HUVECs not transduced with E4ORF1 lentivirus will not survive in serum- and endothelial growth factor-free culture media. 8. Once all HUVECs transduced with GFP lentivirus die, replace X-Vivo 20 culture media with 2 ml of fresh EGM to the wells containing the HUVECs transduced with the E4ORF-1 lentivirus (Fig. 1). 3.6. Passage of E4ORF1 HUVECs (E4ORF1+ ECs)

1. Remove EGM, add 1× PBS without calcium and magnesium, and gently wash cells. 2. Add 0.25% Trypsin–EDTA. 3. Incubate at 37°C for 5 min. 4. Add equal volume of fresh EGM. 5. Spin at 100 × g for 5 min. 6. E4 ECs are split in a 1:3 ratio (see Note 6).

Fig. 1. E4ORF1+ ECs and other primary EC lines, including freshly isolated HUVECs, bone marrow endothelial cells (BMECs), immortalized SV40 Large T, polyoma middle T Antigen, and hTERT BMECs. We found that by day 14 of cultivation in serum-/ cytokine-free only E4+ ECs were able to maintain structural integrity and viability.

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1. Remove femurs and tibias from CD45.2 C57Bl/6 male mice (see Note 7). 2. Denude the bones of all muscle and soft tissues. 3. Carefully remove the ends of the bones. 4. Flush bones with 3 ml of 1× PBS per mouse using a 26 gauge needle. 5. Make bone marrow cells into a single cell suspension by aspirating back up the cells using the same 26 gauge needle and syringe in step 4. 6. Plunge single cell suspension into a 5 ml tube. 7. Filter bone marrow using a 45 mm blue filter cap 5 ml tube. 8. Spin bone marrow at 100 × g for 5 min. 9. Resuspend cells in 1 ml and place on ice. 10. Count cells using hemocytometer and trypan blue exclusion.

3.8. Staining Whole Bone Marrow for Cell Sorting of Hematopoietic Stem and Progenitor Cells

1. Set up two 1.5 ml Eppendorf tubes. Place 1 × 106 into the first tube (no stain control) in 100 ml of 1× PBS and the remaining cells in the other tube (sample) in 300 ml per mouse sacrificed (see Note 8). 2. In the sample tube add the following antibodies (10 ml of each antibody per mouse sacrificed): v450-conjugated Lineage Antibody Cocktail (CD11b; CD41a; CD3; Gr-1; B220; Ter119); APC-conjugated anti-cKit; PECy7-conjugated antiSca-1; and FITC-conjugated anti-CD34. 3. Allow antibodies to stain cells for 30 min at room temperature in the dark. Vortex cell every 5 min to ensure proper staining of all cells. 4. Add 1 ml of 1× PBS and centrifuge cells at 100 × g for 5 min. 5. Aspirate supernatant and resuspend no stain control in 500 ml of 1× PBS and resuspend the sample in 1.5 ml of 1× PBS per mouse sacrificed. 6. Place on ice and immediately begin sorting cells (Fig. 2). 7. Sort hematopoietic stem and progenitor cells (HSPCs) into serum-free X-Vivo media supplemented with 50 ng/ml of mouse stem cell factor (SCF). 8. Spin HSPCs at 100 × g for 5 min. 9. Aspirate supernatant and resuspend in 1 ml of X-Vivo media supplemented with 50 ng/ml of mouse SCF per 10,000 HSPCs (see Note 9).

3.9. Coculture of HSPCs with E4 ECs

1. Passage E4ORF1+ ECs in a 12-well plate. 2. Once confluent, replace EGM with 1 ml of X-Vivo media supplemented with 50 ng/ml of mouse SCF and 10,000 HSPCs (Subheading 3.8).

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Fig. 2. Population gating schema for isolation of phenotypic mouse hematopoietic stem and progenitor cells using fluorescence-activated cell sorting.

3. Every other day supplement X-Vivo media with 50 ng/ml of mouse SCF. 3.10. Passage of HSPCs

1. Once HSPCs reach a cell density of approximately 2 × 106 per well, passage the HSPCs to a fresh well of E4ORF1+ ECs. 2. Remove the EGM from the adjacent well of E4ORF1+ ECs. 3. Replace the EGM with 500 ml of the non-adherent HSPCs from the well that needs to be passaged. 4. Add 500 ml of fresh X-Vivo to each of the two wells. 5. Supplement each well with 50 ng/ml of mouse SCF.

3.11. Enrichment of Cocultured HSPCs

Due to the initial rapid expansion of lineage-committed hematopoietic cells, one must enrich the hematopoietic cells on a weekly basis. By 3 weeks of coculturing with E4ORF1+ ECs, the balance tips towards expanding mostly HSPCs and less mature hematopoietic cells are propagated. 1. Remove all non-adherent hematopoietic cells and place on ice. 2. Add 0.5 ml of 0.25% Trypsin–EDTA to each well to remove all E4ORF1+ ECs and adhered hematopoietic cells. Incubate for 5 min at 37°C.

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3. Deactivate Trypsin–EDTA by adding 0.5 ml of EGM. 4. Spin cells at 100 × g for 5 min. 5. Resuspend cells in 1× PBS supplemented with 2 mM EDTA and 1% BSA. 6. Count cells using a hemocytometer and trypan blue exclusion. 7. According to cell number, follow manufacturer’s instructions for anti-human CD31 microbeads from Miltenyi Biotech. 8. Once hematopoietic cells are separated from the E4ORF1+ ECs, combine them with the non-adherent hematopoietic cells from step 1. 9. Repeat steps 4–6. 10. According to cell number, follow manufacturer’s instruction for anti-mouse lineage depletion microbead from Miltenyi Biotech. 11. Count the lineage-depleted cells (enriched for HSPCs) using a hemocytometer and trypan blue exclusion. 12. Replate 5 × 104 to 1 × 105 lineage-depleted cells on fresh E4ORF1+ ECs per well of a 12-well plate in X-Vivo 20 supplemented with 50 ng/ml of mouse SCF. 13. Stain 2 × 105 the remaining cells with 1 ml of the following antibodies for flow cytometric analysis: APC-conjugated anti-cKit. PECy7-conjugated anti-Sca-1. FITC-conjugated anti-CD34.

4. Notes 1. Avoid letting the gelatin solution evaporate. This will make the surface too thickly coated with gelatin and cause the coating to lift off under aspiration. 2. Plate 1 T25 for every 10 ml of collagenase used. 3. Once HUVECs are attached, feed twice a week with fresh EGM. Once wells are confluent, HUVECs must be passaged or frozen. 4. The dropwise addition of Solution A to Solution B is extremely critical. Addition of Solution A too fast will result in extremely low viral production. 5. Avoid freeze/thaw cycles of the viral stocks, as this will decrease viral titer. 6. The use of gelatin-coated plates is no longer needed. E4ORF1+ ECs have a high affinity to adhere to tissue culture plates.

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7. CD45.2 and male mice are used because hematopoietic cells that are isolated may need to be used in functional transplantation assays. CD45.2 and CD45.1 are congenic C57Bl/6 mice strains. Using the different CD45 cell surface markers allows researchers to distinguish between donor and host cells. Using male mice as a donor gives researchers another tool to distinguish between donor and host by transplanting male donor cells in female host recipients. 8. Compensation beads conjugated to the appropriate fluorophore are used for single color controls for all flow cytometric and FACS protocols. 9. One of the most important cytokines in maintaining HSPCs is SCF. Although E4ORF1+ ECs produce many pro-hematopoietic cytokines, including SCF, human SCF has very little activity on mouse hematopoietic cells. Therefore, it is essential that exogenous mouse SCF is added to ensure the survival of the cocultured mouse HSPCs. References 1. Schofield R (1978) The relationship between the spleen colony-forming cell and the haemopoietic stem cell. Blood Cells 4:7–25 2. El-Badri NS, Wang BY, Cherry, Good RA (1998) Osteoblasts promote engraftment of allogeneic hematopoietic stem cells. Exp Hematol 26:110–116 3. Marusic A, Kalinowski JF, Jastrzebski S, Lorenzo JA (1993) Production of leukemia inhibitory factor mRNA and protein by malignant and immortalized bone cells. J Bone Miner Res 8:617–624 4. Taichman RS, Emerson SG (1994) Human osteoblasts support hematopoiesis through the production of granulocyte colony-stimulating factor. J Exp Med 179:1677–1682 5. Taichman RS, Reilly MJ, Emerson SG (1996) Human osteoblasts support human hematopoietic progenitor cells in vitro bone marrow cultures. Blood 87:518–524 6. Kiel MJ, Yilmaz OH, Iwashita T, Terhorst C, Morrison SJ (2005) SLAM family receptors distinguish hematopoietic stem and progenitor cells and reveal endothelial niches for stem cells. Cell 121:1109–1121 7. Zhang J, Niu C, Ye L, Huang H, He X, Tong WG, Ross J, Haug J, Johnson T, Feng JQ, Harris S, Wiedemann LM, Mishina L, Li L (2003) Identification of the haematopoietic stem cell niche and control of the niche size. Nature 425:836–841

8. Sacchetti B, Funari A, Michienzi S, Di Cesare S, Piersanti S, Saggio I, Tagliafico E, Ferrari S, Robey PG, Riminucci M, Bianco P (2007) Selfrenewing osteoprogenitors in bone marrow sinusoids can organize a hematopoietic microenvironment. Cell 131:324–336 9. Sugiyama T, Kohara H, Noda M, Nagasawa T (2006) Maintenance of the hematopoietic stem cell pool by CXCL12-CXCR4 chemokine signaling in bone marrow stromal cell niches. Immunity 25:977–988 10. Butler JM, Nolan DJ, Vertes EL, VarnumFinney B, Kobayashi H, Hooper AT, Seandel M, Shido K, White IA, Kobayashi M, Witte L, May C, Shawber C, Kimura Y, Kitajewski J, Rosenwaks Z, Bernstein ID, Rafii S (2010) Endothelial cells are essential for the selfrenewal and repopulation of Notch-dependent hematopoietic stem cells. Cell Stem Cell 6:251–264 11. Krosl J, Austin P, Beslu N, Kroon E, Humphries RK, Sauvageau G (2003) In vitro expansion of hematopoietic stem cells by recombinant TATHOXB4 protein. Nat Med 9:1428–1432 12. Miller CL, Eaves CJ (1997) Expansion in vitro of adult murine hematopoietic stem cells with transplantable lympho-myeloid reconstituting ability. Proc Natl Acad Sci USA 94: 13648–13653 13. Willert K, Brown JD, Banenberg E, Duncan AW, Weissman IL, Reya T, Yates JR, Nusse R (2003)

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Wnt proteins are lipid-modified and can act as stem cell growth factors. Nature 423:448–452 14. Kopp HG, Hooper AT, Broekman MJ, Avecilla ST, Petit I, Luo M, Milde T, Ramos CA, Zhang F, Kopp T, Bornstein P, Jin DK, Marcus AJ, Rafii S (2006) Thrombospondins deployed by thrombopoietic cells determine angiogenic switch and extent of revascularization. J Clin Invest 116:3277–3291 15. Antonchuk J, Sauvageau G, Humphries RK (2002) HOXB4-induced expansion of adult hematopoietic stem cells ex vivo. Cell 109:39–45 16. Varnum-Finney B, Xu L, Brashem-Stein C, Nourigat C, Flowers D, Bakkour S, Pear WS,

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Bernstein ID (2000) Pluripotent, cytokinedependent, hematopoietic stem cells are immortalized by constitutive Notch1 signaling. Nat Med 6:1278–1281 17. Moore KA, Ema H, Lemischka IR (1997) In vitro maintenance of highly purified, transplantable hematopoietic stem cells. Blood 89: 4337–4347 18. Rafii S, Shapiro F, Pettengell R, Ferris B, Nachman RL, Moore MA, Asch AS (1995) Human bone marrow microvascular endothelial cells support long-term proliferation and differentiation of myeloid and megakaryocytic progenitors. Blood 86:3353–3363

Chapter 19 Studying Vascular Progenitor Cells in a Neonatal Mouse Model Kirsten A. Kienstra and Karen K. Hirschi Abstract While many murine models have been developed to study adult disease, animal research focused on neonatal and pediatric medicine has been limited by the small size of the mouse pups. Several transplantation, injection, and implantation systems have been used to study the function and role of vascular progenitor populations in adult mice; however, such techniques have been difficult to translate into newborn animals. Herein, we describe a model of neonatal murine intravascular injections that opens opportunity to study many diseases unique to the newborn that might benefit from vascular repair strategies and regenerative medicine. Key words: Endothelial progenitor cell, Neovascularization, Neonatal

1. Introduction Blood vessel formation plays a key role in physiologic and pathologic tissue growth and healing. A clear understanding of the mechanisms regulating postnatal neovascularization, and the precise role(s) played by vascular precursor/progenitor cell populations, will translate into innovative treatment modalities for a wide spectrum of diseases. Various populations of vascular progenitor cells have been isolated from several different tissue types (reviewed in refs. 1–3), and levels have been shown to correlate with outcomes in many vascular disease processes (4–7). Animal models using treatment with endothelial progenitor cells (EPC) have shown improved functional outcomes following cardiac and peripheral vascular ischemia (8–11). Recent neonatal human and animal studies have highlighted the potential role for EPC in

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outcomes of diabetic pregnancies (12), retinopathy of prematurity (7), and bronchopulmonary dysplasia (13, 14). It is important to recognize that due to developmental differences neonates and children are not just small versions of adults. They can have different responses to the same type of injury; for example, following hyperoxia newborns have decreased levels of EPC, while adult levels increase (13). They also are vulnerable to disease processes and injuries unique to their age and developmental stage. As the rate of preterm birth continues to increase and advances in neonatal care improve survival at the extremes of viability, more children survive with often lifelong complications of prematurity, including retinopathy of prematurity, bronchopulmonary dysplasia, and necrotizing enterocolitis. Many of these children might benefit from vascular regenerative medicine strategies to enhance tissue growth and healing. Cord blood, available as an autologous supply at birth, offers rich potential as a source of vascular precursor/progenitor cells for critically ill neonates. Ongoing research is needed to further identify the presence and role of vascular precursor/progenitor cells within newborn and pediatric populations, and to define their function during pediatric-specific disease processes. The increased availability and use of neonatal animal research methods will help develop treatment modalities for a wide variety of newborn diseases, ranging from genetic disorders to complications of prematurity. Transplantation strategies have been used to study the origin and lineage hierarchy of endothelial precursor/progenitor cells, and to further investigate signaling pathways and regulatory mechanisms that play a governing role. Hematopoietic stem cell transplantation techniques have been used to study the relationship between vascular precursor/progenitor cells and the bone marrow compartment (15, 16). Direct injection and/or implantation of various EPC populations have also been widely utilized to investigate the role of progenitor cells in a variety of vascular disease states. To accurately verify incorporation of EPC into the vascular endothelium, the “transplanted” population of cells must be labeled or otherwise distinguishable from endogenous endothelial cells (using eGFP, LacZ, etc.), and the transplanted marker should be colocalized with the expression of mature endothelial cell-specific genes or proteins using high-resolution confocal microscopy (17). While numerous murine models have been developed to study adult vascular disease and regenerative medicine, animal research focused on neonatal and pediatric medicine has been limited by the small size of the mouse pups. Genetic malleability and relatively short reproductive cycles make the mouse an extremely useful model compared with other larger animal models. However, the ability to effectively deliver therapeutic agents intravascularly, whether comprising cell populations or medications, has limited the opportunities for neonatal research using mouse models.

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Adult mice are usually injected via the tail vein or retroorbital venous plexus. In mouse pups, the tail vein is prohibitively small and the eyelids remain fused after birth. Alternative methodology for murine neonatal intravascular injections opens opportunities to study many diseases unique to the newborn that might benefit from vascular repair strategies and regenerative medicine, such as bronchopulmonary dysplasia, retinopathy of prematurity, and necrotizing enterocolitis (18). Thus, we have developed techniques for neonatal injection of various cell populations including HSC and EPC.

2. Materials 2.1. Animals

1. Newborn mouse pups at 0–6 days of life (see Note 1). 2. Lactating dams to provide routine maternal care and continued suckling for experimental pups (see Note 2).

2.2. Neonatal Intravascular Injection

1. 33-gauge, ¼ in. ultra fine gauge needle (Cadence Science, Lake Success, NY). 2. T-connector extension tubing set (4 ¾ in., 0.25 ml; Braun, Bethlehem, PA) (see Note 3). 3. 1 ml tuberculin syringe. 4. Handheld transilluminator (WeeSight, Respironics, Murrysville, PA). 5. Hands-free 2.75× magnification lens (MagEyes, Kerrville, TX). 6. Cotton-tipped swabs.

2.3. Substrates for Injection

1. GIBCO Hanks’ Balanced Salt Solution or other vehicle solution. 2. Cell suspension for injection: £1 × 107 cells suspended in a maximum volume of 100 μl per pup (1 × 108 cells/ml maximum concentration) (see Note 4). 3. Texas Red Dextran, molecular weight 70,000 Da, 50 mg/ml (Invitrogen-Molecular Probes, Eugene, OR) or other fluorochrome.

3. Methods 1. The injectate (specific cellular suspension, medication, fluorescent dye, vehicle control, etc.) is drawn into a 1-ml tuberculin syringe and used to prime the T-connector.

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Fig. 1. Positioning of mouse pup for intravascular injection. Investigator 1 positions the pup obliquely over the transilluminator as shown, using one hand to hold the chest and upper limb, and the other to stabilize the head. The neck is slightly extended and the head rotated to expose the superficial temporal vein.

Bubbles are eliminated to minimize the risk of air emboli. Cell suspensions are not drawn up through the ultra fine gauge needle to avoid cell lysis due to the small diameter of the needle lumen. 2. Consider pup sedation using mild hypothermia. 3. Neonatal injections are best accomplished using a 2-person team, one to hold and position the pup (Investigator 1), and the other to perform the injection (Investigator 2). Injections should be carried out in a darkened room to enhance transillumination and visualization of the vascular tree (see Note 5). For optimal positioning, Investigator 1 places the pup over the transilluminator, using one hand to stabilize the trunk and exposed upper extremity, and the other to position the head and neck. Position the pup obliquely (between supine and lateral) with the neck extended and slightly rotated to the side to best visualize the superficial temporal vein (Fig. 1). During the injection process, pups are closely monitored for signs of airway obstruction or distress (see Note 6). 4. Investigator 2 adjusts the hands-free magnification lenses to optimize visualization of the transilluminated vessels and needle. 5. Investigator 2 positions the ultra fine gauge needle tip with bevel down at a 10–20 ° angle just distal to the bifurcation of

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Fig. 2. Superficial temporal vein injection. Investigator 2 uses the ultra fine gauge needle to access the transilluminated superficial temporal vein. With the needle at a shallow angle and the bevel down, enter the skin at a site just distal to the bifurcation of the vein, and then slowly advance the tip into the vessel lumen.

the superficial temporal vessel, then inserts the needle tip through the skin and slowly advances it into the vessel lumen (Fig. 2). 6. Carefully holding the needle in place, Investigator 2 gently pushes the syringe plunger to begin the infusion. In the case of malposition, injectate is immediately visible in the extravascular space and the injection should be halted and the needle repositioned. After successful injection, we observe blanching of the vessel, followed by visualized flow of injectate throughout the local vascular network (see Note 7). 7. If the injection results in a large degree of extravasation into the local tissues or hematoma formation, we repeat the injection in a more proximal position in the vessel (if still possible to visualize), or access the contralateral side. 8. Following injection, apply pressure to the site with a cottontipped swab for approximately one minute to achieve hemostasis. Avoid excessive pressure to minimize airway compromise. 9. Roll pups in cage bedding to reestablish normal scent and replace in cage with nursing mother (see Note 8). 10. This technique can be used for serial injections. 11. After injection, pups have excellent survival rates with no evidence of morbidity, and studies in our laboratory have demonstrated effective intravascular delivery of cells and other agents (18) (see Note 9).

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4. Notes 1. Injections become difficult after 6 days of life, when pigmentation obscures visualization of the superficial temporal vessel. For the earlier end of the spectrum, we have extended this neonatal intravascular injection methodology to pups delivered prematurely, as early as embryonic day 16. 2. During experiments requiring a high degree of pup manipulation or alterations in routine care, we have occasionally had difficulties with poor maternal care or rejection of pups, especially with C57BL/6 animals. On rare occasions, we have used CD1 surrogate mothers with good success. 3. The T-connector tubing facilitates more stable positioning of the fine gauge needle intravascularly, and helps prevent dislodgement of the needle during syringe manipulation. We prefer small, short tubing to minimize dead space and the need for extra volume injectate. 4. Numbers for total volume and cellular concentration for injection were calculated based on estimates of total blood circulatory volume and published data (19–21). Newborn mice generally weigh 1–3 g in the first week of life (22), with an estimated blood volume of 90–270 μl (approximately 90 ml/kg). 5. A small flash light or other light source can be used to visualize syringe markings to measure the volume of injectate. In addition, we found that a small piece of aluminum foil with a circular hole cut in the center placed over the transilluminator helped to focus the red light source. The transilluminator was enclosed in a clear plastic bag to facilitate clean up. 6. Investigator 2 may choose to exert gentle pressure over the chest, leading to transient venous congestion and improved visualization of the superficial temporal vein. However, this technique increases the risk of airway compression and should be used with caution. 7. As with any injection, the needle must be advanced to a position where the needle lumen is within the vessel lumen. If only the needle tip enters the vessel wall, the injectate will extravasate into the surrounding tissue. While the learning curve for neonatal injections is rapid, it generally takes several practice injections to develop the “feel” for accurate needle insertion and injection. 8. Occasionally, in experiments in which litters are mixed or large amounts of pup manipulation are required, we have also placed a small amount of Vick’s VapoRub topical ointment (Procter and Gamble, Cincinnati, OH) on the nursing mother’s nose to prevent pup rejection.

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9. For neonatal bone marrow transplantation studies, we have treated the pups with sublethal doses of irradiation (4 Gray in two divided doses) to enhance hematopoietic engraftment levels (Hirschi laboratory, unpublished data).

Acknowledgments The authors would like to thank James Adams, M.D. (Baylor College of Medicine, Houston, TX) for sharing his photographic expertise. This work was supported by AHA-TX 0865252F (KAK), AAP NRP (KAK), NIH R01 HL77675 (KKH), NIH R01 HL096360 (KKH), NIH R01 EB005173 (KKH), NIH P20 EB007076 (KKH), and USDA ARS―6250-51000 (KKH). References 1. Chao H, Hirschi KK (2010) Hemato-vascular origins of endothelial progenitor cells? Microvasc Res 79:169–173 2. Hirschi KK (2010) Vascular precursors: origin, regulation and function. Arterioscler Thromb Vasc Biol 30:1078–1079 3. Richardson MR, Yoder MC (2010) Endothelial progenitor cells: quo vadis? J Mol Cell Cardiol 50(2):266–272 4. Jialal I, Devaraj S, Singh U, Huet BA (2010) Decreased number and impaired functionality of endothelial progenitor cells in subjects with metabolic syndrome: implications for increased cardiovascular risk. Atherosclerosis 211:297–302 5. Werner N, Kosiol S, Schiegl T, Ahlers P, Walenta K, Link A et al (2005) Circulating endothelial progenitor cells and cardiovascular outcomes. N Engl J Med 353:999–1007 6. Yip HK, Chang LT, Chang WN, Lu CH, Liou CW, Lan MY et al (2008) Level and value of circulating endothelial progenitor cells in patients after acute ischemic stroke. Stroke 39:69–74 7. Machalinska A, Modrzejewska M, Kotowski M, Dziedziejko V, Kucia M, Kawa M et al (2010) Circulating stem cell populations in preterm infants: implications for the development of retinopathy of prematurity. Arch Ophthalmol 128:1311–1319 8. Hu Z, Zhang F, Yang Z, Yang N, Zhang D, Zhang J et al (2008) Combination of simvastatin administration and EPC transplantation enhances angiogenesis and protects against apoptosis for hindlimb ischemia. J Biomed Sci 15:509–517

9. Jujo K, Hamada H, Iwakura A, Thorne T, Sekiguchi H, Clarke T et al (2010) CXCR4 blockade augments bone marrow progenitor cell recruitment to the neovasculature and reduces mortality after myocardial infarction. Proc Natl Acad Sci USA 107: 11008–11013 10. Oh IY, Yoon CH, Hur J, Kim JH, Kim TY, Lee CS et al (2007) Involvement of E-selectin in recruitment of endothelial progenitor cells and angiogenesis in ischemic muscle. Blood 110:3891–3899 11. Ruifrok WP, de Boer RA, Iwakura A, Silver M, Kusano K, Tio RA et al (2009) Estradiolinduced, endothelial progenitor cell-mediated neovascularization in male mice with hind-limb ischemia. Vasc Med 14:29–36 12. Ingram DA, Lien IZ, Mead LE, Estes M, Prater DN, Derr-Yellin E et al (2008) In vitro hyperglycemia or a diabetic intrauterine environment reduces neonatal endothelial colony-forming cell numbers and function. Diabetes 57: 724–731 13. Balasubramaniam V, Mervis CF, Maxey AM, Markham NE, Abman SH (2007) Hyperoxia reduces bone marrow, circulating, and lung endothelial progenitor cells in the developing lung: implications for the pathogenesis of bronchopulmonary dysplasia. Am J Physiol Lung Cell Mol Physiol 292:L1073–L1084 14. Balasubramaniam V, Ryan SL, Seedorf GJ, Roth EV, Heumann TR, Yoder MC et al (2010) Bone marrow-derived angiogenic cells restore lung alveolar and vascular structure after neonatal hyperoxia in infant mice. Am J Physiol Lung Cell Mol Physiol 298:L315–L323

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15. Jackson KA, Majka SM, Wang H, Pocius J, Hartley CJ, Majesky MW et al (2001) Regeneration of ischemic cardiac muscle and vascular endothelium by adult stem cells. J Clin Invest 107:1395–1402 16. Kienstra KA, Jackson KA, Hirschi KK (2008) Injury mechanism dictates contribution of bone marrow-derived cells to murine hepatic vascular regeneration. Pediatr Res 63: 131–136 17. Purhonen S, Palm J, Rossi D, Kaskenpaa N, Rajantie I, Yla-Herttuala S et al (2008) Bone marrow-derived circulating endothelial precursors do not contribute to vascular endothelium and are not needed for tumor growth. Proc Natl Acad Sci USA 105:6620–6625 18. Kienstra KA, Freysdottir D, Gonzales NM, Hirschi KK (2007) Murine neonatal intravascu-

lar injections: modeling newborn disease. J Am Assoc Lab Anim Sci 46:50–54 19. Luchtman-Jones L, Schwartz AL, Wilson DB (2002) The blood and hematopoietic system. In: Fanaroff AA, Martin RJ (eds) Neonatal-perinatal medicine: diseases of the fetus and infant, 7th edn. Mosby, St. Louis, pp 1183–1254 20. Sands MS, Barker JE (1999) Percutaneous intravenous injection in neonatal mice. Lab Anim Sci 49:328–330 21. Young PP, Hofling AA, Sands MS (2002) VEGF increases engraftment of bone marrowderived endothelial progenitor cells (EPCs) into vasculature of newborn murine recipients. Proc Natl Acad Sci USA 99:11951–11956 22. Theiler K (1972) The mouse house: development and normal stages from fertilization to 4 weeks of age. Pringer-Verlag, New York

Chapter 20 Progenitor Cell Mobilization from Extramedullary Organs Mikhail G. Kolonin Abstract The course of various pathological conditions relies on the mobilization of stem cells and partially differentiated progenitor cells. Bone marrow transplantation studies have demonstrated that medullary hematopoietic and endothelial progenitors can undergo mobilization and trafficking. While the ability of the bone marrow to boost its resources in fighting disease or repairing injury declines with age, other organs have surfaced as reservoirs of various progenitor cell populations. This chapter discusses our current understanding of nonbone marrow-derived progenitor pools, focusing on mesenchymal stem cells. The evidence for the extramedullary progenitor mobilization, with a specific emphasis on white adipose tissue, is presented. Key words: Progenitor cell, Stem cell, Mobilization, Circulation, Bone marrow, Extramedullary

1. Populations of Cells Undergoing Mobilization In the past few decades, it has become increasingly appreciated that mobilization of stem/progenitor cells from adult organs may have high clinical importance (1). Elevated systemic circulation of a number of undifferentiated cell populations has been reported and these precursors are implicated in progression of various pathologies, as well as in the recovery from disease (2). The notion that endogenous stem cells may be activated to mend damaged tissues during injury repair and organ regeneration has prompted the efforts aimed to identify factors mobilizing stem cells for the purposes of regenerative medicine (3). As thoroughly covered in other chapters, mobilization phenomenology and mechanisms have been predominantly established for hematopoietic stem cells (HSC) and their descendent hematopoietic progenitor cells (HPC) (4–6). In the past few years, mobilization of various progenitor cell populations in addition to HSC has been uncovered. Endothelial

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progenitor cells (EPC), giving rise to mature endothelial cells (EC), represent a mobile clinically important cell population (7–9). Recently, stromal mesenchymal progenitors, commonly referred to as mesenchymal stromal cells (MSC), have emerged as a potent cell population capable of migrating throughout the body (10–12). Circulating monocyte-derived fibroblast progenitors (fibrocytes) have been characterized as alternative to the MSC, which become activated in pathological conditions (13). Finally, as recently discovered, very small embryonic-like (VSEL) cells is a progenitor population capable of egressing into circulation, implications of which is yet to be determined (14). All these progenitor types are believed to functionally contribute to vascular, stromal, and parenchymal compartments upon recruitment (2, 15). At present, it is unclear to which extent bone marrow, versus other organs, contributes to the trafficking of each of these populations. Therefore, development of reliable methods to quantify their trafficking and read out their activity is highly important.

2. Non-bone Marrow-Derived Mobilized Cells: Published Evidence

Bone marrow is a bona fide source of progenitor cells mobilized into the bloodstream in response to complex signaling cascades involving chemotactic gradients (16, 17). However, because the quantity and ability of the bone marrow progenitors to respond to mobilization stimuli appears to decline with age, the contribution of cells egressing from this organ to injury repair is likely to progressively decrease in-parallel. On the other hand, hemangioblasts, the precursors of hematopoietic and endothelial cells, as well as HSC have been reported to ectopically accumulate in adult organs other than bone marrow, such as white adipose tissue (WAT) or spleen in certain conditions (18, 19). Tumors present an example of tissue normally located outside of the bone marrow that can disseminate cells into the bloodstream (20). Such mobilization of cancer cells accounts for the distant metastasis of primary tumors, which is largely responsible for cancer lethality. Recent approaches to the detection of circulating tumor cells provide a tool for cancer diagnosis and prognosis (21). The extent at which individual nonmalignant organs other than the bone marrow contribute to the pool of circulating cells is unclear. The current evidence for progenitor cell egress from benign extramedullary organs is predominantly based on the data generated for vascular precursors. In addition to angiogenesis (vascular sprouting through local endothelial cell division), recruitment of circulating progenitors into blood vessels (vasculogenesis) plays an important role in neovascularization (16, 22, 23). For example, bone marrow-derived

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circulating EPC can engraft into tumor vasculature, and promote cancer progression (24, 25), suggesting a possible role in disease progression. Transplantation studies in mice have demonstrated that tumors recruit various populations of progenitor cells from the bone marrow (17, 24, 26); however, several studies have challenged the role of this organ in tumor vascularization (27, 28). The role of other organs in supplying vascular progenitor cells for neovessels is beginning to surface. The concept of non-bone marrow-derived EPC goes back to 2001 when Alessandri and colleagues demonstrated that the human embryonal aorta contains precursors capable of differentiating into mature EC in cell culture (29). In 2005, Ingram et al. presented original evidence that endothelial progenitors reside in the wall of adult blood vessels (30). Later studies identified the localization of endothelial progenitors in a distinct “vasculogenic” zone of the vascular wall between the smooth muscle (tunica media) and connective tissue (tunica adventitia) (31). Such vascular wall resident endothelial progenitor cells (VW-EPC) have been since then discovered in the vasculogenic zone of adult human liver vasculature, as well as in several other organs (15). Subsequent experiments in rodents have unequivocally confirmed that extramedullary precursors contribute to vascularization during wound repair (32). In this study, parabiosis with established cross-circulation between two mice excluded bone marrow as a possible source of progenitor cells for ischemic tissue. Intestine and liver were suggested as possible sources of these progenitors. However, the exact location of precursor cells in these organs has not been defined, and comprehensive analysis of other organs is yet to be done. A population of cells that has begun to surface as a cell type that could be mobilized from organs other than the bone marrow appears to correspond to MSC. These stromal progenitors are found in the majority of adult organs (33). MSC are virtually absent in the peripheral circulation of healthy individuals; however, hypoxia and inflammation signals have been reported to result in MSC mobilization and migration from their niches (34). Desmoplasia is an integral component of cancer progression (35). Therefore, it can be expected that MSC, apparently differentiating into cancer-associated fibroblasts represent a key component of the tumor microenvironment (36, 37). In accordance with this, cancer models are well suitable for investigating MSC mobilization, and the capacity of MSC to “sense” cancer and to home to tumors has been demonstrated (10). Our studies revealing the migration of MSC cells from WAT to experimental tumors in mice (38) challenge the bone marrow as the exclusive source organ. Recently, perivascular cells with clonogenic and proangiogenic potential, which are likely to be an MSC subpopulation, have been isolated from saphenous vein of coronary artery bypass surgery patients (39). Combined, recent observations suggest that not

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only cancer, but also other conditions associated with inflammation can induce the egress of MSC from various resident tissues.

3. Alternative Tissue Sources of Mobilized Cells: Focus on WAT

While the contribution of the majority of extramedullary organs to the pool of circulating progenitor cells mobilized in pathology may be modest, WAT is an exception worth separate discussion. WAT is highly vascularized (40, 41) and, much like tumors, relies on angiogenesis and vasculogenesis for tissue expansion (42, 43). The vasculature of WAT is in the state of constant remodeling and it is possible that adipose EC and EPC have the capacity to exit WAT in certain conditions, such as upon weight loss. WAT expansion, responsible for obesity, relies on proliferation of WAT-specific MSC, which are termed adipose stromal cells (ASC). These cells, abundant in WAT, can be readily harvested for use in regenerative therapies and hundreds of clinical trials are now underway. We and others have shown that ASC comprise the majority of cells in the SVF and display multipotency and proliferation capacity comparable to those of bone marrow MSC, while also having clear unique features (44–46). Apart for serving as progenitor cells, ASC display characteristics of pericytes and cooperate with the endothelium during vascularization by promoting EC proliferation and blood vessel formation at least in part via trophic effects of secreted vascular endothelial growth factor (VEGF), hepatocyte growth factor (HGF), and other angiogenic molecules (46, 47). It has been shown that systemically administered ASC physically migrate to sites of injury and contribute to revascularization of injured mouse organs, including the heart (48). In addition to ASC, various hematopoietic cells can accumulate in WAT quite significantly. Indeed, monocytes/macrophages constitute up to 30% of SVF cells in obese mice. The inherent capacity of this cell population to migrate raises the possibility that they may have overlooked pathological roles outside WAT. Because monocytes are highly plastic and can efficiently acquire fibroblast functions in culture and in vivo (49, 50), both mesenchymal and hematopoietic cells from WAT are potential reservoirs of progenitor cells engaging in disease. A possibility that cells from WAT may undergo spontaneous mobilization and affect the progression of human disease has been recently acknowledged based on our studies demonstrating that obesity is associated with a dramatic increase in systemic MSC circulation (51). Excess WAT-derived progenitors released into the bloodstream in obesity could positively influence postoperative healing and organ regeneration. Mobilization of stromal cells from WAT during operation may explain the “obesity paradox” of better postsurgical recovery observed for obese patients (52). The flip side

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Fig. 1. Design of experiments testing mobilization of WAT cells. WAT-derived cell migration to tumors and their effect on tumor growth is assessed in a transplant model. GFP + WAT is implanted into lean mice prior to tumor grafting, which allows to track mobilization of WAT cells into circulation and their recruitment by the tumor, as well as to measure the resulting effect on cancer progression. Tumor growth is compared to that in control mice.

of excessive stromal cell mobilization has not been realized until recently. According to our hypothesis, WAT-derived cells overabundant in obesity are recruited by tumors at an increased frequency in obese individuals. This phenomenon, confirmed by our data from colorectal cancer patients (12) may account for the epidemiological association between obesity and cancer progression. Consistent with this possibility, obesity is linked with progression of not only cancer, but also of a number of nonmalignant fibrotic conditions. The uncovered function of MSC mobilized from WAT, associated with acceleration of tumor growth, merits investigation of MSC safety in clinical settings involving certain weight loss strategies or adipose stem cell therapies in cancer patients. To determine whether extramedullary cells can be recruited by tumor microenvironment, our laboratory has used WAT as a test source organ in mouse cancer models (38). We have developed an experimental model (Fig. 1) based on transplantation of WAT, all cells of which express green fluorescent protein (GFP) and are thereby traceable. To test whether WAT cells can home to tumors, once GFP-labeled WAT implants assimilated in the host 10 days postimplantation onto lower back, we subcutaneously grafted tumor cells onto upper back. Consistent with adipose cell mobilization into circulation, GFP+ cells were detected in blood by flow cytometry. Immunofluorescence analysis of tissues from WATgrafted animals with anti-GFP and anti-CD31 antibodies demonstrated the presence of vascular and perivascular GFP+ cells within the tumors. Our experiments showed that WAT-derived cells migrate through the systemic circulation and engraft their respective vascular niches in the context of cancer. Importantly, tumors grew significantly faster in mice carrying WAT implants, consistent with the concept of obesity promoting cancer through WAT cells. Inspired by our recent data providing the original evidence that organs other than the bone marrow can serve as a source of cells that can be mobilized and recruited by other organs (38), we have set out to build a model testing whether endogenous (rather than transplanted) organs similarly disseminate vascular/stromal progenitors. In this in vivo competitive repopulation assay, the recruitment of progenitor cells from bone marrow, as opposed to other organs, to tumors is quantified based on transplantation of the bone marrow from mice ubiquitously expressing red fluorescent protein (RFP) into mice ubiquitously expressing GFP. The chimeric

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GFP/RFP mice generated through the bone marrow transplantation allow us to distinguish bone marrow-derived cells (RFP+) from cells derived from other organs (GFP+) in systemic circulation and in growing tumors. Our unpublished results indicate that distinct populations of GFP+ cells undergo mobilization from extramedullary tissues in cancer-bearing mice and are recruited by tumors. Combining this model with diet-induced obesity is currently being used to establish the extent at which WAT contributes to the pool of mobilized cells. We recently identified a molecule expressed uniquely on ASC surface (53). This report, establishing a novel isoform of decorin as a cell surface marker of ASC, gives promise to future prospects on our capacity to track adipose progenitors. The new findings show that it is possible to direct probes to stem cells in vivo in an organ-specific manner. In the future, the identified cell surface biomarker can be exploited for imaging or therapeutic ASC targeting. The ASC-specific peptide probes can be developed into directed cytotoxic compounds to aid in diagnosis, prevention, and/or targeted treatment of adipose-associated disorders such as, but not limited to, body composition disorders, metabolic syndrome, and cancer.

4. Tissue-Resident Stem Cell Pools and Their Migration Capacity: Future Prospects

Recent studies demonstrate that non-bone marrow-derived cells are the predominant source of neointimal smooth muscle cells in experimental in-stent restenosis and transplant arteriosclerosis (54). In this case and in many other studies, the identity of migrating cells has not been assessed. It remains to be determined whether progenitor cell populations other than HPC, HSC, EPC, EC, and MSC discussed above also undergo mobilization. The full spectrum of circulating vascular progenitors, their individual roles, and the identity of the underlying stem cells is yet to be fully understood (55). The example of tumor vascularization illustrates that endothelial mimicry may play an important role obscuring the true progenitor populations typically assumed to be EPC (56). Several non-endothelial populations of circulating cells have been recently implicated in supporting tumor vasculogenesis (57). Tumor-associated dendritic cells (TADCs), a leukocyte population expressing both DC and endothelial markers were shown to promote tumor vascularization (58). TIE2-expressing monocytes (TEMs) (59) as well as other types of recruited blood circulating cells expressing VEGFR1 but not VEGFR2 (60) also participate in cancer vasculogenesis. These CD45+ populations may be similar to monocytoid populations, such as fibrocytes (50) and, therefore, bone-marrow derived. Nevertheless, their transient or prolonged residence in other organs should not be excluded.

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Indeed, monocyte/macrophage accumulation in WAT during obesity progression is well established (61). The discovery of ectopic hematopoiesis taking place in WAT (19) suggests that adipose tissue and possibly other organs (such as spleen and the lymph nodes) can harbor various types of progenitors and may be able to release them into the circulation under certain conditions. Other types of adult precursors not yet studied in the context of mobilization include numerous epithelial stem cell types found in the lung and intestines, as well as mammary stem cells giving rise to both the luminal and myoepithelial cell types of the gland. Wellcharacterized is the migration capacity of neural stem cell found in the adult brain, which are capable of differentiating into neurons and glial cells. Neural crest stem cells, such as those found in hair follicles, gastrointestinal tract, sciatic nerve, cardiac outflow tract, and spinal and sympathetic ganglia, can also generate neurons, Schwann cells, myofibroblasts, chondrocytes, and melanocytes. Olfactory stem cells found in the lining of the nose is a distinct cell population with therapeutic potential. Stem cells of the reproductive organs, such as testicular cells, should not be taken out of the consideration. Finally, dental pulp stem cells is an intriguing population that appears to have multipotency superior to most adult progenitor cell types (62). The capacity to track mobilization of distinct stem cell pools from different tissues will rely on the success in the isolation of markers of the individual cell populations. Our recent success in identifying adipose MSC markers (53) indicates that molecules selectively expressed on the cell surface exist and can be isolated through high-throughput screens. Probes recovered based on their homing to lung, muscle, and bone marrow stromal cells set the foundation for subsequent identification of probes specific for MSC of these organs. The combination of in vivo phage-displayed combinatorial peptide library selection approach with flow cytometrybased isolation of stem cells of interest that we have established could in theory be applied to any organ and stem cell population of interest. Systematic application of such technologies will eventually uncover the tissue-specific cell surface proteomes and the reliable stem cell markers will enable tracking mobilization and migration of individual stem cell populations in development and disease.

Acknowledgments Supported by Komen for the Cure (KG080782), American Heart Association (0835434N), Cancer Prevention and Research Institute of Texas (RP100400), NIH Prostate SPORE (CA140388) and American Cancer Society (CNE-119003) awards to MGK.

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INDEX

A Adipocytes ........................................140, 151, 175, 178, 180 Adipose stromal cells (ASC) ................................... 246, 248 Adverse events ..................................................... 42, 43, 112 Angiocrine factors ...........................................................222 Angiogenesis ...........................................140, 143, 150, 156, 157, 165, 166, 244, 246 Apheresis ......................................... 2, 37–39, 43, 71, 73, 75, 79–83, 85, 86, 89, 90, 93–95, 97–109 ASC. See Adipose stromal cells (ASC)

CXCL12 (SDF-1)............................................................. 16 CXCR4 ............................6–9, 16, 17, 24, 31, 41, 51–53, 59, 79, 80, 140, 141, 144, 145, 160, 211, 212, 215 Cytokine ....... 6, 7, 9, 16, 24, 26, 38, 43, 80–82, 85, 111, 118, 129–130, 191, 192, 211, 222, 224, 225, 228, 232 Cytotoxic ................................................................... 80, 248

D Dimethylsulfoxide (DMSO) ....................... 98, 99, 103, 104, 109, 111, 112, 168

E

B Biomarker .................................................156, 159, 166, 248 Blood peripheral................................... 1, 2, 5, 9, 15, 31, 37, 40, 43, 49, 51, 54–59, 71, 79–81, 83, 85, 98, 102, 115, 118, 127, 130, 140, 143, 145, 157–160, 167, 191–196, 203, 207–218 umbilical cord .....................................115, 157–160, 217 Bone marrow derived progenitor cells .......................................... 1–3, 6 transplantation ............... 37, 38, 49–51, 57, 60, 121–123, 136, 137, 158, 177, 185, 236, 241, 245, 247, 248

C Cancer ................................... 43, 97–98, 165–166, 174, 181, 183, 187, 244–250 hematological ........................................................ 37, 49 Cardiovascular ................................................. 112, 159–161 CD34 positive cells................................................ 85, 97, 98 Chemokines............... 3, 6, 9, 16, 24, 26, 43, 51, 79, 117, 212 Chemotherapy ............................. 6, 7, 16, 32, 37, 38, 41, 43, 51, 70, 71, 73–75, 79–83, 98, 111, 166 Chondrocytes ........................... 140, 151, 176, 178, 180, 249 Circulation................................... 1, 4, 16, 17, 20, 21, 26, 28, 30, 85, 144, 145, 157–159, 209, 243–249 Clinical protocol .................................................... 16, 69–76 Clodronate.................118, 119, 123–125, 127, 128, 130, 136 COBE® Spectra Apheresis System™ ..................................81 Colony forming assays ........................23, 119, 128–130, 141 Combination treatment ....................................... 53, 55, 141 Cryopreservation ............................2, 98–101, 103–106, 111

Egress .............. 1–3, 6, 10, 15, 16, 32, 51, 140, 143, 244, 246 Endothelial cells .................................. 25, 51, 150, 155, 157, 158, 166, 167, 208, 222, 225–228, 236, 244 Endothelial progenitor cells (EPC) ...................... 17, 32, 69, 119, 140–150, 156–161, 166, 207–212, 214–218, 235–237, 244–246, 248 Engraftment .................................... 4, 19–20, 27–30, 38, 49, 52, 74, 98, 181, 185, 241 EPC. See Endothelial progenitor cells (EPC) Extramedullary ........................................................ 243–249

F FACS. See Fluorescence activated cell sorting (FACS) Fibroblasts ..........................................17, 191–204, 244, 246 Fibrocytes .........................................192, 196, 197, 202, 203 Fibrosis .................................................................... 191, 192 Flow cytometry............................ 17–20, 59, 71, 82, 98, 100, 101, 103, 108, 119, 125–128, 144, 165–167, 169, 170, 178, 179, 185, 186, 205, 210–212, 224, 247, 249 Fluorescence ...................................... 58, 171, 174, 176, 177, 181, 182, 185, 186, 189, 230 Fluorescence activated cell sorting (FACS) ................ 17–23, 25, 30, 58, 168, 177, 181–184, 188, 213–217, 232

G G-CSF. See Granulocyte colony-stimulating factor (G-CSF) Graft .................. 49, 56, 57, 59, 60, 71, 80, 97, 111, 112, 174 Granulocyte colony-stimulating factor (G-CSF) ................... 16, 38, 49, 70, 79, 81, 117, 119

Mikhail G. Kolonin and Paul J. Simmons (eds.), Stem Cell Mobilization: Methods and Protocols, Methods in Molecular Biology, vol. 904, DOI 10.1007/978-1-61779-943-3, © Springer Science+Business Media, LLC 2012

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STEM CELL MOBILIZATION: METHODS AND PROTOCOLS 254 Index Granulocyte-macrophage colony-stimulating factor GM-CSF......................................................... 38, 70 Growth factors ..........................................24, 41, 72, 73, 86, 176, 211, 225, 228, 246

H Hematopoiesis ............................ 16, 28–31, 37, 50, 221, 249 Hematopoietic reconstitution .........................................4, 28, 29, 56–58, 60, 130, 136, 222 stem/progenitor cells (HPC/HSPC) ................ 1–10, 15, 17, 20, 23, 25, 26, 40, 43, 51, 53, 56, 69, 79–83, 85, 97, 98, 117, 119, 139, 156–158, 166, 207, 208, 222, 225, 229, 230, 236, 243, 244, 246 Hypocalcemia ........................................................ 86, 94–95

Myeloablative ...................................................... 97, 98, 222

N Neonatal ...........................................155, 159, 160, 235–241 Neovascularization.....................................32, 140, 156–158, 160, 161, 235, 244 Niche ................................................... 3–5, 7, 9, 50–52, 119, 207, 221–232, 245, 247 Non-hematopoietic ........................................ 166, 208, 209

O Osteoblasts ............................................ 5, 16, 30, 31, 50, 51, 117–119, 140, 151, 176, 178, 180

P

Lentivirus ................................................................ 227–228 Leukapheresis .............................................38, 39, 69, 73, 76 Leukocytes ..... 15, 17, 20, 23, 85, 86, 93, 111, 122–132, 135, 136, 152 Liposomes ....................................................... 123, 124, 128

PBL. See Peripheral blood, leukocytes (PBL) PBMC. See Peripheral blood, mononuclear cells (PBMC) PBSC. See Peripheral blood, stem cells (PBSC) Perfusion.....................................30, 139–141, 143–145, 159 leukocytes (PBL) ........................... 17, 20, 23, 85, 86, 93, 111, 122–132, 135, 136, 152 mononuclear cells (PBMC) ....................................... 192 stem cells (PBSC) ............................... 36, 43, 49, 79, 111 Plasma ...................... 7, 16, 26, 27, 29, 52, 56, 74, 85, 87, 90, 93, 97–104, 107, 109, 118, 191, 192, 195, 197, 202 Plerixafor ...................7, 43, 50, 51, 69–71, 75, 76, 79–82, 85 Processing ................... 97–109, 111–115, 120, 135, 169, 176

M

R

Macrophages ....................................... 23, 38, 39, 50, 56, 70, 118, 147, 150, 157, 188, 249 Mesenchymal stromal/stem cells (MSC).................... 17, 31, 32, 69, 119, 140–145, 147–149, 151–152, 173–189, 207–212, 214–218, 244–249 Microenvironment..............15–17, 20, 21, 25, 26, 30, 31, 41, 50, 174, 181, 182, 187, 221, 222, 224, 245, 247 Migration .......................................... 3, 8, 17–19, 24–25, 32, 174, 181, 245, 247–249 Mobilization endothelial progenitors ............ 17, 69, 70, 119, 140, 149, 156–157, 245 hematopoietic progenitors .........................18, 22, 25, 26, 51, 53, 69, 85, 86, 243 mesenchymal progenitors ..........................................244 Monocytes .................................... 31, 56, 150, 157, 244, 249 Mouse model ........................................3, 159, 160, 235–241 MSC. See Mesenchymal stromal/stem cells (MSC) Multispectral imaging ..............................174, 177, 185, 188 Murine .................................... 17, 18, 20–26, 28, 30, 31, 58, 141–145, 174, 175, 178–180, 183, 187, 192, 194, 197–200, 208, 209, 236, 237

Red blood cells .............................. 83, 97, 99, 108, 146, 152, 167, 171, 209, 212, 213 Regeneration ..................................... 69, 139, 140, 158, 161, 207, 208, 221, 222, 225, 243, 246

I Inflammation .........15, 20, 143, 173–189, 191, 209, 245, 246 Infusion ............................... 75, 83, 86, 94, 98, 99, 106–108, 111–115, 121, 141, 142, 145, 239

L

S Serum amyloid P ............................................. 191–193, 197 Side effects ................................. 41, 42, 72, 75, 76, 113, 166 Stem/progenitor cell collection .......................................................................2 hematopoietic (HPC/HSC) .................................... 1–10 Sysmex XE-2100L automated hematology analyzer ............................................................ 79, 82

T Targeted delivery .............................................................173 Thawing ....................................... 97, 98, 100, 106, 109, 111 Therapy ..................................................................... 74, 160 Tissue/wound repair ........................................ 181, 191, 245 Trafficking ...................................... 4, 7, 28, 51, 55, 243, 244 Transgenic ........................ 118, 173, 174, 177, 181–183, 187

STEM CELL MOBILIZATION: METHODS AND PROTOCOLS 255 Index Transplantation allogeneic ..... 2, 37–39, 43, 49–51, 71, 81, 82, 97, 98, 111 autologous.............................2, 15, 24, 25, 37, 38, 43, 50, 51, 69, 71–73, 79–83, 97, 98, 101, 107, 236 Tumorigenesis .................................................................155

V Vascular endothelial growth factor (VEGF)............ 211, 246 Vascular progenitor cells ...................157–160, 235–241, 245 Vasculature ........143–145, 155, 158, 160, 165, 166, 245, 246

Vasovagal ............................................................... 85, 93, 95 VEGF. See Vascular endothelial growth factor (VEGF) Very small embryonic-like stem cells (VSEL) ......... 207–218

W White adipose tissue........................................................244

X Xenograft........................................................... 28, 174, 187