Developmental Vascular Biology (Current Topics in Developmental Biology)

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Series Editor Gerald P. Schatten Director, PITTSBURGH DEVELOPMENTAL CENTER Deputy Director, Magee-Women’s Research Institute Professor and Vice-Chair of Ob-Gyn-Reproductive Sci. & Cell Biol.-Physiology University of Pittsburgh School of Medicine Pittsburgh, Pennsylvania 15213

Editorial Board Peter Gru¨ss Max-Planck-Institute of Biophysical Chemistry Go¨ttingen, Germany

Philip Ingham University of Sheffield, United Kingdom

Mary Lou King University of Miami, Florida

Story C. Landis National Institutes of Health National Institute of Neurological Disorders and Stroke Bethesda, Maryland

David R. McClay Duke University, Durham, North Carolina

Yoshitaka Nagahama National Institute for Basic Biology, Okazaki, Japan

Susan Strome Indiana University, Bloomington, Indiana

Virginia Walbot Stanford University, Palo Alto, California

Founding Editors A. A. Moscona Alberto Monroy

Contributors Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Roland Baron (103), Yale University School of Medicine, New Haven, Connecticut 06520 Sonia D’Silva (49), Department of Biology, University of Rochester, Rochester, New York 14627 Susan L. Forsburg (129), Molecular and Computational Biology Section, University of Southern California, Los Angeles, California 90089 Holly V. Goodson (49), Department of Chemistry and Biochemistry, The University of Notre Dame, Notre Dame, Indiana 46556 Dave Hansen (185), Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada T2N-1N4 Michele M. Hickey (217), Department of Cell and Molecular Biology, Abramson Family Cancer Research Institute, and Howard Hughes Medical Institute, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Luca Jahreiss (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Karen Lyons (1), Department of Orthopaedic Surgery, University of California, Los Angeles, California 90095; Department of Molecular, Cell and Developmental Biology, University of California, Los Angeles, California 90095 Fiona M. Menzies (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Rita K. Miller (49), Department of Biology, University of Rochester, Rochester, New York 14627 Jeffrey K. Moore (49), Department of Biology, University of Rochester, Rochester, New York 14627

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Robert Pogue (1), Department of Orthopaedic Surgery, University of California, Los Angeles, California 90095 Brinda Ravikumar (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Georges Rawadi (103), ProStrakan Pharmaceuticals, 93230 Romainville, France SergioRoman-Roman (103), ProStrakan Pharmaceuticals, 93230 Romainville, France David C. Rubinsztein (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Shinji Saiki (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Sovan Sarkar (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom Tim Schedl (185), Department of Genetics, Washington University School of Medicine, Saint Louis, Missouri 63110 M. Celeste Simon (217), Department of Cell and Molecular Biology, Abramson Family Cancer Research Institute, and Howard Hughes Medical Institute, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Angel P. Tabancay, Jr. (129), Molecular and Computational Biology Section, University of Southern California, Los Angeles, California 90089 Andrea Williams (89), Department of Medical Genetics, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge CB2 2XY, United Kingdom

Contents

Contributors

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1 BMP Signaling in the Cartilage Growth Plate Robert Pogue and Karen Lyons I. II. III. IV. V. VI. VII. VIII. IX.

Introduction 2 Signaling Pathways Regulating Chondrogenesis 5 Overview of the BMP Signaling Pathway 8 Regulation of TGF-b/BMP Signaling 10 Expression of TGF-b Superfamily Members, Their Receptors, and Inhibitors in Precartilaginous and Cartilaginous Elements 14 TGF-bs and BMPs in Chondrogenesis 16 Downstream EVectors of BMP Signaling: Smads and p38 25 Interactions with Other Signaling Pathways 28 Perspectives 32 Acknowledgments 35 References 35

2 The CLIP-170 Orthologue Bik1p and Positioning the Mitotic Spindle in Yeast Rita K. Miller, Sonia D’Silva, Jeffrey K. Moore, and Holly V. Goodson I. II. III. IV. V. VI. VII. VIII.

Introduction 50 Structure of the Bik1 Protein 50 Bik1p Interactions with Other Proteins 60 Phenotypes and Activities 61 Role of Bik1p in Spindle Positioning 63 Models for Plus-End Binding 70 Regulation of CLIP-170 and Bik1p by Phosphorylation Bik1p at the Kinetochore 76

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Contents IX. Bik1p in Genomic Studies 76 X. Concluding Remarks and Perspective Acknowledgments 79 References 79

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3 Aggregate-Prone Proteins Are Cleared from the Cytosol by Autophagy: Therapeutic Implications Andrea Williams, Luca Jahreiss, Sovan Sarkar, Shinji Saiki, Fiona M. Menzies, Brinda Ravikumar, and David C. Rubinsztein I. Proteinopathies 90 II. Intracytoplasmic Aggregate-Prone Proteins Are Cleared by Autophagy 93 III. Autophagy Induction Has Additional Antiapoptotic Consequences 95 IV. A New mTOR-Independent Autophagy Pathway 95 V. A Protective Role for Aggregates: Autophagy Upregulation VI. Conclusions 98 Acknowledgments 98 References 99

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4 Wnt Signaling: A Key Regulator of Bone Mass Roland Baron, Georges Rawadi, and Sergio Roman-Roman I. II. III. IV. V. VI. VII. VIII. IX.

Introduction 104 The Wnt Signaling Pathway 105 LRP5 and -6: Structure and Function 108 Human Genetic Evidence for the Role of Wnt Signaling in Bone Metabolism 110 How Is the HBM Mutation Constitutively Activating LRP5 and Wnt Signaling? 111 Mouse Genetics Confirms the Essential Role of the Wnt/b-Catenin Pathway in Bone 112 How Is Wnt Signaling AVecting Bone Homeostasis? 115 EVects of Wnts on Osteoblasts 116 Is the Wnt Pathway the Final Common Pathway of Bone Morphogens? 117

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Contents X. Is Wnt Signaling the Pathway Linking Mechanosensing to Bone Remodeling? 119 XI. Pharmacological Manipulation of Wnt Signaling in Bone 120 XII. Conclusion and Perspectives 121 References 121

5 Eukaryotic DNA Replication in a Chromatin Context Angel P. Tabancay, Jr. and Susan L. Forsburg I. II. III. IV. V. VI.

Chromatin Structure Regulates Access to the DNA 130 Replication Origins and Origin-Binding Proteins 137 Assembly of the Prereplication Complex on Chromatin 144 Origin Activation and Replication 150 Replication Elongation Through Chromatin 154 Conclusions 162 Acknowledgments 162 References 163

6 The Regulatory Network Controlling the Proliferation–Meiotic Entry Decision in the Caenorhabditis elegans Germ Line Dave Hansen and Tim Schedl I. Introduction 186 II. Temporal and Spatial Pattern of Proliferation and Initiation of Meiotic Development 187 III. Summary of the Core Regulatory Pathway Controlling the Proliferation–Meiotic Development Decision 192 IV. Control of the Adult Pattern of Proliferation and Meiotic Entry V. Regulation of the Activity of the GLD-1 and GLD-2 Pathways VI. Additional Components and Pathways Regulating the Proliferation–Meiotic Entry Decision 206 VII. Conclusions 210 Acknowledgments 211 References 211

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7 Regulation of Angiogenesis by Hypoxia and Hypoxia-Inducible Factors Michele M. Hickey and M. Celeste Simon I. II. III. IV. V. VI. VII. VIII.

Introduction 218 Mechanisms of Angiogenesis 219 The Hypoxia-Inducible Factor Family 221 The Role of HIF in Developmental Angiogenesis The Role of HIF in Adult Tissues 231 HIF and Ischemic Injury 232 HIF and Cancer 234 Conclusions 240 Acknowledgments 241 References 241

Index 259 Contents of Previous Volumes

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BMP Signaling in the Cartilage Growth Plate Robert Pogue* and Karen Lyons*,{ *Department of Orthopaedic Surgery, University of California Los Angeles, California 90095 { Department of Molecular, Cell and Developmental Biology University of California, Los Angeles, California 90095

I. Introduction II. Signaling Pathways Regulating Chondrogenesis A. The IHH/PTHrP Signaling Loop B. The FGF Pathway C. The Wnt Pathway D. BMPs III. Overview of the BMP Signaling Pathway A. The TGF‐ Superfamily B. The TGF‐ and BMP Signaling Cascades IV. Regulation of TGF‐ /BMP Signaling A. Extracellular Modulators of BMP Signaling B. The Extracellular Matrix C. Intracellular Regulation V. Expression of TGF‐ Superfamily Members, Their Receptors, and Inhibitors in Precartilaginous and Cartilaginous Elements A. TGF‐ s B. BMPs VI. TGF‐ s and BMPs in Chondrogenesis A. TGF‐ s in Chondrogenesis B. BMPs in Chondrogenesis C. Targets of BMP Action During Early Chondrogenesis D. BMPs in Early Chondrocyte DiVerentiation E. BMP Actions in the Growth Plate VII. Downstream EVectors of BMP Signaling: Smads and p38 A. Smad‐Mediated Signaling in Chondrogenesis B. p38 Pathways in Chondrogenesis VIII. Interactions with Other Signaling Pathways A. Synergy Between BMP and IHH/PTHrP Pathways B. Antagonistic Interactions Between BMP and FGF Pathways C. The Balance Between TGF‐ and BMP Pathways in Osteoarthritis IX. Perspectives Acknowledgments References

Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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0070-2153/06 $35.00 DOI: 10.1016/S0070-2153(06)76001-X

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Transforming growth factor‐ (TGF‐ ) superfamily members play diverse roles in all aspects of cartilage development and maintenance. It is well established that TGF‐ s and bone morphogenetic proteins (BMPs) play distinct roles in the growth plate. This chapter discusses key experiments and experimental approaches that have revealed these roles, and progress toward the identification of previously unsuspected roles. Current understanding of the mechanisms by which diVerent TGF‐ and BMP pathways exert their functions is discussed. Finally attempts to utilize this information to promote cartilage regeneration, and important issues for future research, are outlined. ß 2006, Elsevier Inc.

I. Introduction The highly reproducible sequence of skeletal development and the stratified organization of the cartilage growth plate imply that the mechanisms controlling cell behavior during chondrogenesis must be regulated precisely. Several signaling systems, including the insulin‐like growth factor (IGF), fibroblast growth factor (FGF), Wnt, Indian hedgehog (IHH)/parathyroid hormone‐ related peptide (PTHrP), and bone morphogenetic protein (BMP) pathways act in concert to orchestrate skeletogenesis (Kronenberg, 2003; Minina et al., 2001, 2002; Pathi et al., 1999; Yoon and Lyons, 2004). Understanding how these pathways interact is essential in understanding skeletogenesis, and for the development of future treatments for the osteochondrodysplasias that arise due to genetic mutations, as well as defects that occur through trauma or aging. This chapter will focus on the BMP pathway, one of the major signaling systems known to play essential roles in the commitment of cells to the chondrogenic lineage and in their subsequent progression within the growth plate from proliferation through hypertrophic diVerentiation and apoptosis. Chondrogenesis is initiated by the specification of osteochondral progenitors from mesenchymal cells. Depending on the skeletal element, osteochondral progenitors arise from the neural crest (craniofacial bones), sclerotome of the paraxial mesoderm (axial elements), or lateral plate mesoderm (appendicular bones). Prior to condensation, cells destined to form condensations respond to positional cues, mediated in part by HOX genes, which determine the sizes and locations of these condensations. As their name implies, osteochondral progenitors have the potential to give rise to bone‐ forming osteoblasts and cartilage‐forming chondrocytes. The first overt sign of chondrogenesis commences with the condensation of osteochondral progenitors into aggregates (Fig. 1) (Kronenberg, 2003; Yoon and Lyons, 2004). Fate mapping studies have shown that these progenitors arise from mesenchymal cells expressing the transcription factor Sox9 prior to the onset of condensation (Akiyama et al., 2005). SOX proteins are Sry‐related

1. BMP Signaling in the Cartilage Growth Plate

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Figure 1 Mesenchymal condensation and growth plate structure. Limb bud outgrowth (A) arises in the lateral plate mesoderm. (B) BMP signaling here induces condensation and proliferation of mesenchymal cells and promotes survival of cells within these condensations. (C) These condensations enlarge through proliferation. A perichondrium consisting of elongated cells forms at the border of the condensation. (D) The growth plates begin to form under the influence of signals arising from the condensed cells and the perichondrium. During growth plate formation, chondrocytes exit the cell cycle and begin to undergo terminal diVerentiation to produce the distinctive zones of the growth plate: resting, proliferative, prehypertrophic, and hypertrophic. Hypertrophic chondrocytes produce factors that stimulate the formation of the periosteum, and the invasion of blood vessels and osteoblasts from the region of the periosteum into the hypertrophic zone. Oppositely oriented growth plates in the long bones separate as bones are formed.

homeobox transcription factors that induce expression of chondrocytic genes such as type II collagen in response to BMP signaling. Sox9 is clearly a major eVector of chondrogenesis; in Sox9 mutants, chondrogenic condensations do not form. Through mechanisms that are not fully understood, a further lineage restriction occurs within the condensations, leading to a population of committed chondroprogenitors located within the center of the aggregate that continue to express Sox9, and a population of osteochondral progenitors located at the periphery, known as the perichondrium (Fang and Hall, 1997; Nakashima and de Crombrugghe, 2003; Nakashima et al., 2002). A population of osteochondral progenitor cells resides in the perichondrium throughout skeletal development. These cells serve as a source of chondrocytes that are responsible, at least in part for appositional

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growth, and as a source of osteoblasts that ultimately replace the cartilage with bone. Studies have shown that the ability of these perichondrial cells to give rise to osteoblasts rather than chondroblasts requires the expression of the transcription factor osterix and is dependent on Wnt signaling (Hill et al., 2005; Nakashima et al., 2002). The cells in the centers of the condensations diVerentiate into chondroblasts. These cells proliferate and produce a specialized extracellular matrix (ECM) containing collagens II, IX, and XI, and the proteoglycan aggrecan, in addition to other components. Within the condensations that are destined to become bone, the chondroblasts organize into structures called growth plates, which ultimately are responsible for bone growth (Fig. 1). Growth plates form after chondrocytes in the centers (diaphyses) of the condensations proceed toward terminal diVerentiation. This central region is known as the primary ossification center, and is the first region of the cartilage anlage that will be replaced by bone. Within the growth plate, distinct populations of chondrocytes can be defined based on cellular morphology and expression of marker genes. Small and round chondrocytes are found at the distal ends of presumptive long bones, in an area called the resting zone (RZ). This zone serves as a source of chondrocytes, which when stimulated to divide become part of the proliferative zone (PZ). A cell shape change from round to flat is induced by interactions with the cartilage ECM, and marks the formation of the columnar zone (CZ). The highest rate of proliferation in the growth plate is found near the top of the CZ and the rate decreases as the descendent cells move down the columns (Smits et al., 2004). The entry of cells from the RZ into the PZ and their continued proliferation is dependent on the action of the PTHrP (Pthlh—Mouse Genome Informatics), which is produced by periarticular chondrocytes (Kobayashi and Kronenberg, 2005; Kronenberg, 2003). As cells within the CZ become further and further distanced from the source of PTHrP, they undergo growth arrest and diVerentiate into prehypertrophic chondrocytes at the bottom of the CZ. The prehypertrophic cells produce the secreted factor Indian hedgehog (IHH), which signals back to periarticular cells to promote expression of PTHrP. IHH thus regulates the onset of terminal diVerentiation (Kronenberg, 2003). Cell volume increases up to 20‐fold (Goldring et al., 2006) as prehypertrophic chondrocytes diVerentiate to produce the hypertrophic zone (HZ), changing their ECM from a predominantly type II collagen‐containing matrix to one containing type X collagen. It has been estimated that this increase in the size of chondrocytes as they undergo hypertrophy is responsible for 44–59% of long bone growth, and that the diVerent lengths of various long bones are related to diVerences in cell sizes (Ballock and O’Keefe, 2003; Wilsman et al., 1996). The remainder of the long bone expansion is thought to be due to proliferation and ECM synthesis.


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The final cell type in the growth plate is the late hypertrophic chondrocyte. These cells express markers of preosteoblasts, such as matrix metalloproteinase 13 (MMP13), which promotes vascular invasion and replacement of the cartilage by bone. The final fate of late hypertrophic cells is a subject of debate. The normal fate is apoptotic cell death, but cells that escape death may be able to diVerentiate into preosteoblasts (Bianco et al., 1998; Roach et al., 1995). Concurrent with the onset of hypertrophic diVerentiation, cells in the perichondrium flanking the HZ diVerentiate into osteoblasts. This region of the perichondrium becomes known as the periosteum because it ossifies and contributes osteoblasts. Blood vessels and osteoblasts from the newly formed bone invade the hypertrophic region, and replace the cartilage with bone and bone marrow (Kronenberg, 2003).

II. Signaling Pathways Regulating Chondrogenesis A number of signaling pathways regulate the survival, proliferation, and diVerentiation of chondrocytes within the growth plate. These include the IGF, FGF, Wnt, IHH/PTHrP, TGF‐ , and BMP pathways (Kronenberg, 2003; Li et al., 2005; Olney et al., 2004; Provot and Schipani, 2005; Yoon and Lyons, 2004). These signals control the balance between proliferation and diVerentiation so that the cells enter and exit the cell cycle at the appropriate time. A. The IHH/PTHrP Signaling Loop As discussed earlier, IHH and PTHrP play key roles in regulating the entry and exit of cells into and out of the columnar zone. IHH controls at least three distinct steps in the growth plate. First, IHH, produced by prehypertrophic chondrocytes, stimulates expression of PTHrP in periarticular cells, thereby regulating the onset of hypertrophic diVerentiation. PTHrP in turn signals back to proliferating chondrocytes in the CZ by binding to its receptor, preventing their apoptosis and inhibiting their diVerentiation into IHH‐expressing prehypertrophic cells. Second, IHH stimulates the entry of RZ chondrocytes into the CZ independently of PTHrP. Finally, IHH induces the ossification of the perichondrium (Kobayashi et al., 2005; Kronenberg, 2003; Minina et al., 2001, 2002).

B. The FGF Pathway Whereas IHH and PTHrP are positive signals promoting chondrocyte proliferation, FGFs provide essential inhibitory signals. The majority of FGFs

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are expressed in the perichondrium. The importance of FGF signaling in chondrogenesis was revealed by the discovery that achondroplasia (ACH), the most common genetic form of dwarfism, is caused by activating mutations in the FGF receptor gene Fgfr3 (Ornitz and Marie, 2002; Wilkie et al., 2002). Fgfr3 expression is initiated in condensing chondrocytes and is retained in proliferating growth plate chondrocytes (Ohbayashi et al., 2002; Peters et al., 1993). Mouse models of ACH (gain‐of‐function) and Fgfr3/ animals (loss‐of‐function) demonstrate that a predominant role for FGFR3 is to limit chondrocyte proliferation (Colvin et al., 1996; Deng et al., 1996; Naski et al., 1998). The eVects of FGFs are mediated by multiple pathways, two of which are known to operate in chondrocytes: signal transducer and activator of transcription (STAT) and mitogen‐activated protein kinase (MAPK) pathways (Murakami et al., 2004; Ornitz and Marie, 2002; Sahni et al., 1999). FGF induces nuclear translocation of STATs 1, 5a, and 5b by phosphorylating them (pSTAT1/5); pSTAT1 consequently induces expression of cell cycle inhibitors as well as proapoptotic signals (Laplantine et al., 2002; Meraz et al., 1996; Sahni et al., 1999). Stat1/ mice are protected from the chondrodysplasia caused by FGF2 overexpression, supporting an essential role for STAT1 in FGF‐mediated inhibition of proliferation (Sahni et al., 2001). Ihh is another key target: Fgfr3ACH mice exhibit decreased expression of Ihh and its receptor, Patched, whereas expression of these genes is upregulated in mice lacking Fgfr3 (Naski et al., 1998; Ornitz and Marie, 2002), implying that FGF signaling exerts a negative regulatory influence on the PTHrP/IHH pathway. The roles of FGF pathways in chondrocyte diVerentiation are unclear. A prevalent view holds that FGFs delay diVerentiation (Ornitz and Marie, 2002). The rate of transit of cells from the proliferative to the hypertrophic zone is delayed in Fgfr3ACH mice (Murakami et al., 2004). However, as limb culture studies indicate that Fgfr3ACH mice or wild‐type (WT) limbs exposed to FGFs express increased levels of the late hypertrophic chondrocyte markers osteopontin (OPN) and MMP13, an alternative view postulates that FGFs promote at least some aspects of hypertrophic diVerentiation (Minina et al., 2002; Sahni et al., 2001). Support for this notion comes from microarray experiments (Dailey et al., 2003); in chondrocytic cell lines, FGFs promote the expression of some (MMP13, OPN) markers associated with terminal diVerentiation, but not others (collagen X). Regardless of the interpretation, the eVects of FGFs on terminal diVerentiation appear to be mediated at least in part by MAPK pathways: overexpression of a constitutively active (CA) mutant of MAPK/ERK kinase 1 (MEK1), the kinase that phosphorylates and activates ERK1/2, leads to the same shortened zone of hypertrophy seen in Fgfr3ACH mice (Murakami et al., 2004).


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C. The Wnt Pathway As is true for FGF pathways, Wnt pathways also play multiple roles throughout chondrogenesis, and some of these are growth inhibitory. There are 19 Wnt family members, and they transduce their signals through several diVerent pathways [Nelson and Nusse (2004) and references therein]. Among these, the canonical pathway has been studied most extensively. Expression patterns of many Wnts suggest that they signal to mesenchymal condensations and regulate osteoblast and chondrocyte diVerentiation (Guo et al., 2004; Kato et al., 2002; Parr et al., 1993). The canonical Wnt‐signaling pathway is transduced by stabilizing ‐catenin protein (Logan and Nusse, 2004). Stabilized ‐catenin accumulates in the cytoplasm and translocates to the nucleus, where it activates downstream gene transcription. Forced expression of ‐catenin in prechondrogenic cells in vitro inhibits chondrocyte diVerentiation, whereas its overexpression in chick limb buds and mouse embryos accelerates conversion of proliferating chondrocytes to hypertrophic chondrocytes (Guo et al., 2004; Hartmann and Tabin, 2000; Ryu et al., 2002). Thus, Wnt signaling through ‐catenin appears to inhibit prechondrogenic cells from acquiring and/or maintaining a chondrogenic fate, but is required to allow exit of proliferating chondrocytes from the cell cycle in order to promote hypertrophy. These eVects of Wnt pathways may be mediated in part through physical and functional antagonism between Sox9 and ‐catenin (Akiyama et al., 2004). In vivo, Sox9 is expressed throughout the PZ, but is downregulated in hypertrophic chondrocytes, whereas nuclear ‐catenin is found at low levels in the PZ and higher levels in the HZ; overexpression of Sox9 and inactivation of ‐catenin in chondrocytes produce similar phenotypes in vivo (Akiyama et al., 2004; Bi et al., 1999; Enomoto‐Iwamoto et al., 2002; Guo et al., 2004). These antagonistic interactions predict that pathways that are required for expression of Sox9, such as BMP and TGF‐ pathways, discussed later, will antagonize Wnt pathways in the growth plate.

D. BMPs Bone morphogenetic proteins were initially discovered based on their ability to induce ectopic cartilage and bone (Wozney, 1989). The mechanisms underlying this unique capability have been the subject of intense investigation, but mechanistic explanations of the eVects of BMP pathways could not be articulated until the framework of the BMP signal transduction pathway had been elucidated (Derynck and Zhang, 2003; Massague et al., 2005). BMPs play important roles at multiple stages of skeletal development, from early patterning and mesenchymal cell condensation, to regulation of chondrocyte

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proliferation and maturation in the growth plate, and to joint formation and ossification (Cheng et al., 2003; Satoh et al., 2005). This chapter will focus on the roles of BMPs in the growth plate. Particular attention will be paid to the elements of their signaling cascades, regulation, timing and locations of signaling events, roles in regulating the various stages in the chondrocyte life cycle, and interactions with other signaling pathways.

III. Overview of the BMP Signaling Pathway A. The TGF‐b Superfamily The transforming growth factor‐ superfamily is a large and diverse group of structurally related secreted growth factors (Massague, 1998). The members of the superfamily can be subdivided based on structural considerations and on the signal transduction pathways that they activate. Members of the TGF‐ /activin/nodal class signal through specific receptors to activate common downstream signals, whereas members of the BMP class signal through a structurally related set of receptors to activate a diVerent set of signals.

B. The TGF‐b and BMP Signaling Cascades The transforming growth factor‐ superfamily members bind as dimers to receptor complexes consisting of heterotetrameric combinations of types I and II serine‐threonine kinase receptors (Fig. 2). At least two distinct pathways mediate TGF‐ /BMP signaling: a canonical Smad pathway and an MAPK pathway. A number of reviews discuss TGF‐ and BMP signal transduction pathways (Derynck and Zhang, 2003; Feng and Derynck, 2005; Shi and Massague, 2003); hence these will be reviewed only briefly here. Ligand binding to the receptor complex causes the type II receptor to phosphorylate the type I receptor within a glycine/serine‐rich (GS) domain. The type I receptors subsequently activate receptor Smads (R‐Smads; Smads 1, 2, 3, 5, and 8) by phosphorylating them. R‐Smads then recruit the common partner Smad (Smad4; co‐Smad). These complexes enter the nucleus, where they bind the promoters of target genes, and recruit other transcription factors (such as Hoxc‐8, FAST‐1, OAZ, Runx2, AP‐1, and STAT) (Derynck and Zhang, 2003; Massague et al., 2005). Receptor Smads bind DNA directly or in cooperation with other transcription factors (Derynck and Zhang, 2003; Massague et al., 2005). Since the binding aYnity of Smads for DNA is relatively weak, it is thought that DNA‐ binding partners are required for eYcient DNA binding in vivo (Massague et al., 2005; Miyazono et al., 2001). Transcription is activated or repressed by


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Figure 2 Schematic representation of elements of the BMP signaling pathway. The elements of TGF‐ and BMP signaling pathways have been reviewed in detail elsewhere (e.g., Massague et al., 2005; Moustakas and Heldin, 2005) and are only briefly outlined here. BMP dimers bind to heterotetramers of type I and type II receptors. The availability of ligand to the receptors is controlled by secreted antagonists such as noggin and chordin, by ECM components such as syndecan and fibrillin, which can have positive or negative eVects, and possibly by coreceptors such as DRAGON. Ligand binding causes the type II receptors to phosphorylate the type I receptors. In the canonical Smad pathway, the BMP receptors phosphorylate R‐Smads, Smads 1, 5, and 8. These subsequently complex with Smad4 (Co‐Smad) and enter the nucleus to induce gene transcription in cooperation with other coactivators. The intracellular localization of Smads is influenced by extracellular components such as hyaluronan (via CD44) and cytoskeletal components such as filamin. Negative regulation of Smad and BMP receptor stability occurs intracellularly through I‐Smads and Smurfs. The noncanonical BMP pathway employs p38 MAP kinase, which is activated through TAB/TAK1.

recruitment of general transcriptional activators (such as CBP/p300) or repressors (TGIF, Ski, and Sno) to the complex (Massague et al., 2005). For members of the TGF‐ /activin/nodal subclass, ligands bind to receptor complexes containing the type I receptors ALK4/ActR‐IB, ALK5/T R‐I, and/or ALK7, and activate Smads 2 and 3. Ligands of the BMP subclass bind to the type I receptors ALK2/ActRI, ALK3/BMPRIA, and/or ALK6/ BMPRIB, and activate Smads 1, 5, and 8. However, these distinctions are not always exact. For example, TGF‐ ligands bind the type II TGF‐ receptor (TGF‐ RII) and interact not only with ALK5, which activates Smads 2 and 3, but also with ALK1, which activates Smads 1 and 5 (Goumans

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et al., 2003). Similarly, TGF‐ ligands can bind to ALK2, which is related structurally to ALK1, when complexed with TGF‐ RII, activating Smads 1 and 5 (Lai et al., 2000). Thus, combinatorial interactions of type I and II receptors may allow diVerential signaling in response to the same ligand (Derynck and Zhang, 2003). DiVerent BMPs bind with diVerent aYnities to the three type I receptors. Thus, the activity of a TGF‐ or BMP pathway in a given tissue will depend not only on the level of active ligand itself, but also on the constellation of receptors that are expressed. For example, BMPs can be subdivided into distinct groups based on amino acid sequence comparisons. BMPs 2 and 4 are closely related to each other, as are BMPs 5, 6, and 7. Growth and diVerentiation factors (GDFs) 5, 6, and 7 form a third subgroup within the BMP family. BMP4 binds preferentially to BMPRIA and BMPRIB, whereas BMP7 binds with higher aYnity to ALK2 and BMPRIB than to BMPRIA; GDF5 interacts most eYciently to BMPRIB (Macias‐Silva et al., 1998; Nishitoh et al., 1996; ten Dijke et al., 1994; Yamashita et al., 1995). Thus, a given ligand can induce diVerent outcomes with diVerent eYciencies depending on the composition of the receptor complex. In addition to signaling through Smads, members of the TGF‐ superfamily can activate the ERK1/2, Jun kinase (JNK), and p38 MAPK pathways (Derynck and Zhang 2003; Moustakas and Heldin, 2005). For the most part, the physiological relevance of these non‐Smad pathways in mediating the eVects of TGF‐ s and BMPs in chondrocytes is not well understood. Two coreceptors for BMP signaling have been identified. RGMa, HFE2, and DRAGON are members of the repulsive guidance molecule family of genes. RGMa and DRAGON have been shown to bind to BMP2 and BMP4 (but not other family members), as well as to BMP receptors, and enhance BMP signaling (Babitt et al., 2005; Samad et al., 2005). Expression and activity of these coreceptors have not yet been demonstrated in cartilage. It is possible that the localized expression of these or similar proteins contributes to diVerential BMP signaling, making the signal stronger in certain areas of the growth plate, thereby allowing a single signal to have diVering eVects on diVerent cells, based on signal strength.

IV. Regulation of TGF‐b/BMP Signaling The potential for TGF‐ s and BMPs to aVect diverse aspects of cell behavior, even within a single organ such as cartilage, suggests that their activities must be tightly regulated. As it is not just the mere presence or absence of the TGF‐ / BMP signal, but also its strength, that influences patterns of gene expression, rate‐limiting factors that modulate the activity levels of TGF‐ /BMP cascades can have profound eVects in cartilage. Consistent with this notion, many mechanisms for negative regulation of BMP pathways have been described.


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A. Extracellular Modulators of BMP Signaling A considerable degree of regulation occurs at the level of ligand availability. A large number of secreted antagonists, including noggin, chordin, follistatin, gremlin, and DAN form high‐aYnity complexes with BMPs extracellularly, and prevent them from binding to their receptors (Balemans and van Hul, 2002; Canalis et al., 2003). Nifuji et al. (2004) showed that adenoviral overexpression of noggin in cell and organ culture models inhibits expression of chondrogenic markers including collagens and Ihh, although they were not able to show whether or not Smad1 phosphorylation was aVected. The best evidence demonstrating an essential role for secreted BMP antagonists comes from analysis of mice lacking noggin. These exhibit massive cartilage overgrowth and failure to form joints, similar to phenotypes seen with BMP overactivity. Noggin is coexpressed with, and induced by, BMPs (Kameda et al., 1999; Nifuji and Noda, 1999), resulting in a feedback loop that limits the intensity and range of the BMP signal. Chordin expression levels are inversely proportional to the degree of maturation, with the highest levels in the periarticular perichondrial region (Zhang et al., 2002). These findings led Zhang et al. (2002) to hypothesize that a decreasing chordin gradient from the RZ to the hypertrophic zone could influence the onset of hypertrophy stimulated by BMPs. Chordin‐null mice show craniofacial defects and vertebral abnormalites (Bachiller et al., 2003). While these defects do not support a major requirement for chordin in the growth plate, functional redundancy with other BMP antagonists such as noggin is likely. The actions of some antagonists, such as chordin, are negatively regulated by additional binding proteins. One of these is twisted gastrulation (Tsg; a chordin antagonist that is expressed in the developing digital rays and hind limbs) (De Robertis and Kuroda, 2004; Scott et al., 2001). Others include tolloid (a BMP1‐like metalloproteinase that cleaves and inactivates chordin) and kielin (a chordin‐like protein) that has been shown to enhance BMP responsiveness in murine fibroblasts (Lin et al., 2005). The roles of these molecules have not been investigated in depth in chondrocytes.

B. The Extracellular Matrix The ECM has also been implicated in the regulation of BMP signaling. Fisher et al. (2006) used limb bud micromass cultures to show that heparin sulfate proteoglycans (HSPGs) such as syndecan‐3 can negatively aVect BMP‐ mediated chondrogenesis, possibly by sequestering ligands like BMP2, thereby limiting their diVusion and/or availability to receptors. In contrast to these inhibitory eVects on BMP2 signaling, others have found evidence that at earlier stages of condensation, and during osteoblast diVerentiation, HSPGs

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may enhance aspects of BMP signaling (Irie et al., 2003; Seghatoleslami and Koshe, 1996). Takada et al. (2003) found that heparan sulfate, a major component of the proteoglycan ECM in cartilage, enhanced BMP2 activity in C2C12 culture. Furthermore, mice with genetrap mutations in the chondroitin‐4‐sulfotransferase 1 gene showed diminished BMP signaling, along with accelerated diVerentiation, indicating that chondroitin sulfate (CS) may have an important positive influence on BMP signaling in the hypertrophic zone (Kluppel et al., 2005). Whether this is a direct result of an interaction between BMP and CS is unknown. Other studies have shown that fibrillins sequester, and thus influence, TGF‐ and BMP signaling pathways. Fibrillins are the major structural components of microfibrils, architectural elements that are ubiquitous in connective tissues (Sakai et al., 1986). Mutations in fibrillin‐1 lead to Marfan syndrome, a heritable disorder of connective tissues aVecting the skeleton, eye, and cardiovascular system (Dietz and Pyeritz, 1995). Analyses of fibrillin‐1–deficient mice demonstrate that at least one manisfestation of Marfan syndrome, a failure in distal alveolar septation, is caused by abnormal activation of TGF‐ , demonstrating that TGF‐ is normally sequestered in an inactive form by fibrillin‐1 (Neptune et al., 2003). This sequestration is not limited to TGF‐ s, as a direct interaction between BMP7 and fibrillin‐1 has also been demonstrated (Gregory et al., 2005). Genetic evidence of a similar relationship between fibrillin‐2 and BMP7 has been revealed; mice doubly heterozygous for null alleles of fibrillin‐2 and Bmp7 exhibit syndactyly and polydactyly, skeletal features that are never seen in the corresponding single heterozygotes (Arteaga‐Solis et al., 2001). From the above‐mentioned discussion, it is clear that sequestration within the matrix can have both positive and negative eVects on pathway output. In vivo studies will be required to determine the physiological relevance and impact, positive or negative, of these interactions on BMP signaling in cartilage. Further involvement of the ECM is seen in the form of hyaluranon–cell interactions mediated through CD44 (Peterson et al., 2004). This study showed that Smad1 phosphorylation induced by BMP7 is dependent on an interaction between Smad1 and the cytoplasmic domain of CD44. Inhibition of this interaction reduced BMP7‐mediated Smad1 phosphorylation and nuclear translocation.

C. Intracellular Regulation In addition to a potentially vast number of extracellular molecules that bind BMPs, additional levels of control are operative intracellularly. These include modulation of BMP receptor and R‐Smad activity or stability via


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ubiquitin‐mediated proteolysis through the actions of inhibitor Smads (I‐Smads) 6 and 7, and Smad ubiquitination regulatory factors (Smurfs) 1 and 2. I‐Smads can inhibit TGF‐ and BMP signaling cascades both by blocking R‐Smad phosphorylation by type I receptors (Smads 6 and 7) and inhibiting interactions between R‐Smads and Smad4 (Smad6); Smad6 preferentially downregulates BMP signaling, whereas Smad7 downregulates TGF‐ , activin, and BMP signaling (Ebara and Nakayama, 2002; Hata et al., 1998; Massague et al., 2005). In a further demonstration of negative feedback, there is also evidence that BMPs can stimulate expression of Smad6 (Ishida et al., 2000). Smads are regulated by ubiquitin ligases that target them for proteosomal degradation (Datto and Wang, 2005; Murakami et al., 2003). Smurfs show Smad specificity (and therefore pathway specificity), with Smurf1 targeting Smads 1 and 5, and Smurf2 targeting Smads 2 and 3. Smad4 is regulated by a ubiquitin ligase called ectodermin (Dupont et al., 2005). In the case of Smurf1, cooperative activity with the inhibitory Smads has been shown, and these activities include targeting BMP type I receptors for degradation (Murakami et al., 2003). However, BMP receptors and Smads may not be physiological substrates of Smurf1 in skeletal tissues in vivo; no growth plate defects are seen in mice lacking Smurf1 (Yamashita et al., 2005). Rather, Smurf1‐null mice exhibit increased bone mass due to enhanced sensitivity of osteoblasts to the BMPs. However, this eVect is not mediated by increased levels of Smads 1 and 5. Rather, loss of Smurf1 leads to increased basal activity of JNK. The absence of a growth plate phenotype may reflect functional redundancy with Smurf2. Thus, whether Smurfs have a role in chondrogenesis in vivo, and whether components of Smad pathways are targets of Smurfs, remains to be determined. It has been shown that Smad2 and Smad5 interact with filamins (Sasaki et al., 2001; Tang et al., 2003). Filamins are actin cross‐linking proteins that are important for regulation of the cytoskeleton. Mutations in filamin‐b are responsible for a number of skeletal phenotypes (Bicknell et al., 2005; Krakow et al., 2004), raising the possibility that dysregulation of BMP pathways may be involved in these conditions. Protein interactions, such as that between Smads and filamins, may be used to suggest the involvement of BMPs or other pathways in disease phenotypes. Conversely, similarities in phenotypes have been used to imply interactions between proteins. For example, three proteins responsible for diVerent forms of brachydactyly were examined for interactions based on the similar phenotypes caused by mutations (Sammar et al., 2004). This study showed that GDF5 signaling through BMPRIB in chondrocytes can be modulated by the tyrosine kinase receptor Ror2. Ror2 can interact directly with BMPRIB to inhibit signaling through Smad1/5.

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V. Expression of TGF‐b Superfamily Members, Their Receptors, and Inhibitors in Precartilaginous and Cartilaginous Elements A. TGF‐bs TGF‐ s 1–3 can be detected in the perichondrium and periosteum as soon as these tissues arise, and persist there until after birth (Gatherer et al., 1990; Millan et al., 1991; Pelton et al., 1990; Sandberg et al., 1988; Serra and Chang, 2003). TGF‐ s 1–3 are coexpressed in hypertrophic cartilage in the growth plate (Ivkovic et al., 2003; Minina et al., 2005). Expression of TGF‐ s 1 and 2 persists in adult articular cartilage, suggesting a role for the TGF‐ /activin/ nodal pathway in maintenance in addition to development (Ellingsworth et al., 1986; Morales et al., 1991). Consistent with this possibility, ALK5/ T R‐I and TGF‐ RII are expressed in perichondrium in addition to proliferating and hypertrophic chondrocytes within the growth plate (Serra et al., 1999). The presence of nuclear localized Smad2 and Smad3 provides strong evidence that chondrocytes throughout the growth plate are engaged in TGF‐ /activin signaling; Smad2 is expressed most strongly in PZ chondrocytes, whereas Smad3 is most prominent in hypertrophic chondrocytes (Sakou et al., 1999). Nuclear Smad2 can be detected readily in adult articular cartilage, but the percentage of positive cells in this tissue declines significantly with age.

B. BMPs A role for BMP pathways in early stages of condensation is suggested by the finding that multiple BMPs, including BMPs 2, 4, 5, and 7, and GDF5 are expressed surrounding or within early condensing mesenchyme in limb buds and developing somites (Dudley and Robertson, 1997; Kingsley, 1994; Lyons et al., 1995; McMahon et al., 1998; Winnier et al., 1995). Once a perichondrium has formed, multiple BMPs become highly expressed there (Chang et al. 1994; Jones et al., 1991; Lyons et al., 1990; Macias et al., 1997; Minina et al., 2005; Wozney et al., 1988). Hypertrophic chondrocytes express BMPs 2 and 6 (Fig. 3) (Lyons et al., 1990; Minina et al., 2005). BMP7 is expressed in proliferating chondrocytes, particularly in regions adjacent to the perichondrium (Lyons et al., 1995; Minina et al., 2005). Finally, GDFs 5, 6, and 7 are highly expressed in regions of joint formation (Settle et al., 2003; Storm and Kingsley, 1999; Wolfman et al., 1997). Thus, overlapping expression of multiple BMP ligands is characteristic of every region of the growth plate. The presence of distinct and overlapping expression patterns in the


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Figure 3 Expression patterns of BMP pathway components in the growth plate and perichondrium.

perichondrium raises the possibility that BMPs expressed here control the development of chondrocytes in the growth plate by exposing them to a progression of signals. The type I BMP receptors also exhibit characteristic expression patterns in the growth plate. BMPRIA is highly expressed in perichondrium, proliferating, and hypertrophic chondrocytes. ALK2 is expressed primarily in resting and proliferating chondrocytes, and BMPRIB is found throughout the growth plate, including the developing articular surface and in the perichondrium (Minina et al., 2001, 2005; Sakou et al., 1999; Yi et al., 2001; Zou et al., 1997). The type II BMP receptor is also expressed throughout the growth plate. Thus, as is the case for the ligands, essentially every region of the growth plate expresses at least one type I BMP receptor and most regions express multiple type I receptors. Consistent with the expression of multiple BMP ligands and receptors, nuclear localized Smad1/5 is abundant in early condensations (Yoon et al., 2005). Similarly, expression of Smads 1, 5, and 8 can be readily detected at later stages throughout the growth plate and in developing articular cartilage (Kobayashi et al., 2005; Minina et al., 2005; Sakou et al., 1999; B. Yoon and K. Lyons, unpublished data).

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These well‐defined expression patterns of the elements of the BMP pathway suggest important functions for BMP signaling at essentially every stage of chondrogenesis. The existence of distinct and overlapping patterns for the ligands allows for diVerent combinations of ligands to meet diVerent combinations of receptors, and may help to explain how a wide range of signaling outcomes can be achieved by a limited number of receptors. These expression patterns are not static; examination of growth plates at diVerent stages of development reveals diVerent expression patterns (Minina et al., 2005). Essentially nothing is known about the regulatory mechanisms responsible for these dynamic patterns of ligand and receptor expression. Also, it is not clear whether the mechanisms by which BMP ligand and receptor expression are regulated on a transcriptional or posttranscriptional level are similar in all skeletal elements, since the majority of studies have focused on analysis of only a few appendicular elements. Kingsley and colleagues (DiLeone et al., 2000; Mortlock et al., 2003) have used a BAC transgenic approach to identify distal control elements regulating expression of Bmp5 and Gdf6 in cartilage. Through characterization of an insertional mutation, Baur et al. (2000) have identified a control region far upstream of the Bmpr1b transcription start site that serves as a tissue‐specific enhancer of Bmpr1b expression in cartilage. Further dissection of these control elements, and additional studies of the transcriptional control of other BMP ligands and receptors, will be essential for an understanding of skeletal development.

VI. TGF‐bs and BMPs in Chondrogenesis In spite of extensive investigation, the roles of the TGF‐ /activin/nodal and BMP pathways in chondrogenesis and maintenance of adult cartilage remain unclear. In vivo and in vitro systems have been used extensively to investigate the eVects of TGF‐ and BMP signaling pathways in skeletal development. The most commonly used models are cell culture, in vitro limb culture, manipulation of chick embryos, and transgenic and knockout mouse models. Many groups have exploited cell lines that are capable of undergoing chondrogenic diVerentiation following stimulation with TGF‐ s and BMPs, and/or culture in micromass systems, where high cell density promotes cell–cell interactions that encourage cartilage nodule formation.

A. TGF‐bs in Chondrogenesis TGF‐ s regulate chondrocyte diVerentiation in vitro, but have varying eVects depending on the developmental stage and culture conditions.


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In general, TGF‐ s promote and are required for chondrogenesis in cultures of early undiVerentiated mesenchyme (Kulyk et al., 1989; Pittenger et al., 1999), but inhibit terminal chondrocyte diVerentiation in diVerentiated chondrocytes and in organ culture (Ballock, et al., 1993; Ferguson, et al., 2000; Moses and Serra, 1996; Serra et al., 1999). A study has shown that Sox9 physically associates with Smad2/3 and that this association leads to synergistic enhancement of the ability of Sox9 to activate the Col2a1 promoter (Furumatsu et al., 2005b), providing additional evidence for a physiologically significant role for TGF‐ signaling pathways in chondrogenesis. Studies involving adenoviral overexpression of Smads 2 and 3, dominant‐ negative forms of these Smads, and siRNA approaches show that both the induction of chondrogenesis in mesenchymal cell lines and suppression of maturation of overt chondrocytes in response to TGF‐ appear to be mediated primarily by Smad3, with Smad2 playing a supporting role (Ferguson et al., 2000; Furumatsu et al., 2005a; Li et al., 2006; Valcourt et al., 2002). The mechanisms underlying the distinct eVects of Smads 2 and 3 are unclear, but may be related to diVerences in DNA‐binding properties, ability to interact with cooperating factors, and/or susceptibility to proteolysis (Massague et al., 2005). Roles for the TGF‐ /activin/nodal pathway in vivo are suggested by phenotypes of mice and humans with mutations in components of the TGF‐ pathway. Loss of neither Tgf 1 nor Tgf 3 is associated with obvious skeletal defects (Dickson et al., 1995; Kaartinen et al., 1995; Proetzel et al., 1995; Shull et al., 1992). However, targeted deletion of Tgf 2 results in perinatal lethality with multiple skeletal defects, including alterations in size and shape of long bones (Sanford et al., 1997). Loss of the type I TGF‐ receptor Alk5/Tgf rI, the type I activin receptor Alk4/Acvr1b, or the type II receptor Tgf r2 leads to early embryonic lethality, precluding an analysis of the roles of these receptors in skeletal development (Larsson et al., 2001; Oshima et al., 1996). However, mice bearing floxed alleles of Tgf r2, crossed to a Col2a1‐Cre transgene (TGF‐ r2;Col2‐ Cre), die perinatally and exhibit defective formation of vertebrae and intervertebral discs; development of long bones is essentially unaVected (BaY et al., 2004). Several interesting observations arise from these findings. The severe axial defects, but lack of obvious abnormalities in appendicular elements, may arise either due to fundamental diVerences in the requirement for TGF‐ signaling in axial versus appendicular elements or diVerences in the onset of Col2a1‐Cre expression in axial versus appendicular elements. The more severe axial defects may reflect a significant role for TGF‐ pathways prior to, but not after, condensation during skeletal development. Alternatively, TGF‐ signaling may play a more predominant role in the perichondrium than in growth plate chondrocytes. That TGF‐ signaling is required in the perichondrium has been suggested by the finding that

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overexpression of a dominant‐negative form of TGF‐ RII in this tissue blocks the ability of exogenous TGF‐ to inhibit chondrocyte maturation in organ culture (Alvarez et al., 2001, 2002). Finally, it is important to bear in mind that TGF‐ s can bind to TGF‐ RII, and when complexed with type I receptors such as ALK2, transduce a BMP signal (activation of Smad1/5/8) (Desgrosellier et al., 2005). Thus, the extent to which the Tgf 2/ and Tgf‐ rII;Col2‐Cre phenotypes reflect decreased signaling through TGF‐ /activin/ nodal pathways versus BMP pathways is an unknown, but important issue. Given the possibility for cross talk between TGF‐ and BMP pathways at the level of ligand and type II receptor utilization, additional studies are needed to evaluate the roles of the TGF‐ /activin/nodal pathway in chondrogenesis in vivo. Investigations of the roles of TGF‐ –specific Smads, Smads 2 and 3, provide insight into the role of the canonical TGF‐ / activin/nodal pathway in chondrocytes. As discussed earlier, in vitro studies indicate that Smads 2 and 3 have direct eVects in prechondrogenic mesenchyme and in growth plate chondrocytes. Evidence that these findings are physiologically significant comes from the observation that Smad3/ chondrocytes exhibit enhanced maturation in vitro (Li et al., 2006). Overexpression of dominant‐negative forms of Smads 2 and 3 in the perichondrium of metatarsal rudiments demonstrates that these Smads mediate the ability of exogenous TGF‐ to inhibit chondrocyte proliferation and maturation. However, no diVerences in growth and diVerentiation were noted in the absence of TGF‐ , raising the possibility that perichondrial signaling through these Smads is required for responsiveness to exogenous TGF‐ , but does not play a major role in normal development. Moreover, the phenotypes of Smad3/ mice and mice overexpressing a dominant‐negative form of TGF‐ RII indicate roles for TGF‐ /activin/ nodal canonical signaling pathways in postnatal, but not developing, articular and growth plate chondrocytes. These mutants develop severe progressive osteoarthritis; the hypertrophic zone is enlarged and the PZ is correspondingly reduced in some adult Smad3/ mutants (Yang et al., 2001). The extent to which Smad2/3 pathways are active in chondrogenesis during development remains uncertain, because the possibility of functional redundancy has not yet been investigated in vivo. The finding that Smad2þ/ mice exhibit defective development of Meckel’s cartilage in organ culture strongly supports the possibility of functional redundancy elsewhere in chondrogenesis (Ito et al., 2002). Resolution of these issues will require generation of TGF‐ R1‐conditional null mice, generation of Smad2/3 double mutants, and deletion of TGF‐ RII in perichondrium. Clearly, resolution of these issues will require ablation of TGF‐ RI in cartilage, as well as ablation of TGF‐ RII in limb mesenchyme prior to condensation, and in perichondrium.


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B. BMPs in Chondrogenesis The ability of BMPs to induce ectopic chondrogenesis strongly suggests that these factors play essential roles in commitment and condensation. In vitro systems have been particularly informative for investigating the roles of BMP pathways at these early stages. BMPs induce chondrogenic diVerentiation in pluripotent mesenchymal cell lines (Denker et al., 1999; Haas and Tuan 1999; Ju et al., 2000; Kramer et al., 2000; Majumdar et al., 2001; Nifuji et al., 2004). Cell–cell interactions are crucial in these cultures, and a key event in BMP2‐mediated cartilaginous nodule formation is upregulation of N‐cadherin; the requirement for homophilic interactions mediated by N‐cadherin was demonstrated by the finding that N‐cadherin inhibitors neutralize the eVects of BMPs (Haas and Tuan, 1999). A central role for BMPs in early stages of chondrogenesis in vivo was first demonstrated in chick limbs. The use of the secreted inhibitor noggin has been a particularly informative approach; noggin is specific for BMP ligands and its application permits antagonism of endogenous BMP pathways while avoiding artifacts that might arise from dominant‐negative receptors or exposure to physiologically irrelevant levels of BMPs. Noggin blocks condensation, leading to a complete absence of cartilage; in complementary studies, overexpression of constitutively active BMP receptors promotes expansion of cartilage at the expense of other tissues (Capdevila and Johnson, 1998; Pizette and Niswander, 2000). These and related studies establish that BMP signaling is required, and acts as an instructive signal, for the commitment of mesenchymal cells to a skeletal fate. In these experiments, while constitutively active forms of either BMPRIA or BMPRIB promote chondrogenesis, only the overexpression of dominant negative (DN)‐BMPRIB, and not DN‐BMPRIA or DN‐ALK2, blocks these events, leading to the suggestion that BMPRIB is the major transducer of BMP signals during condensation (Enomoto‐Iwamoto et al., 1998; Fujii et al., 1999; Kawakami et al., 1996; Zou et al., 1997). However, Bmpr1b‐null mice exhibit a restricted skeletal phenotype, with major defects confined to phalangeal elements, challenging this view and suggesting either overlapping functions or a more prominent role for BMPRIA during condensation in mice (Baur et al., 2000; Yi et al., 2000). The finding that early stages of chondrogenic condensation and diVerentiation proceed relatively normally in mice lacking either Bmpr1a or Bmpr1b, whereas simultaneous ablation of both receptors leads to a nearly complete loss of condensations, demonstrates extensive functional redundancy between these receptors (Yoon et al., 2005). C. Targets of BMP Action During Early Chondrogenesis Identifying the downstream targets that mediate the ability of BMPs to initiate chondrogenesis is clearly an area of importance. Key insights have

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come from examining the relationship between BMP pathway activity and Sox9 expression. As discussed, Sox9 is the earliest marker for cells committed to the osteochondroprogenitor lineage, and in mice lacking Sox9, cartilage condensations do not form (Bi et al., 1999). Sox9 is expressed continuously in chondrocytes up to the stage of hypertrophic diVerentiation, and in addition to its essential role in mediating condensation Sox9 is required along with the related transcription factors L‐Sox5 and Sox6 for maintenance of the chondrocyte phenotype (Bi et al., 1999; Smits et al., 2001). A number of studies have examined the relationship between BMP signaling and expression of L‐Sox5, Sox 6, and Sox9 (Chimal‐Monroy et al., 2003; Fernandez‐Lloris et al., 2003; Zehentner et al., 1999). BMPs promote expression of Sox9 in mesenchymal cell lines undergoing chondrogenesis, suggesting that BMPs act upstream of Sox9. However, antisense Sox9 blocks the ability of BMPs to induce chondrogenic diVerentiation, demonstrating that Sox9 is essential for BMP‐induced chondrogenesis and suggesting that BMPs and Sox9 participate in a positive feedback loop (Fernandez‐Lloris et al., 2003; Zehentner et al., 1999). The relationship between BMP action and Sox9 expression has also been investigated in vivo (Chimal‐Monroy et al., 2003). Implantation of BMP2‐ soaked beads near condensed cartilage leads to upregulation of Sox9 in condensations, whereas noggin beads elicit downregulation of Sox9. These studies indicate that BMPs are not suYcient to initiate Sox9 expression in uncondensed limb mesenchyme, as BMP beads can only induce Sox genes in condensed cells. However, in this system, Sox genes are induced by TGF‐ in uncondensed mesenchyme (Chimal‐Monroy et al., 2003). In the absence of genetic evidence linking TGF‐ /activin/nodal pathways to early stages of chondrogenesis (discussed previously), the physiological relevance of this activity of TGF‐ is unclear at present. In vivo evidence that BMPs are essential for maintenance of Sox9 expression in early condensations comes from analysis of mice in which a floxed allele of Bmpr1a, the Col2a1‐Cre transgene, and a null allele for Bmpr1b have been utilized to ablate both receptors in cartilage (Yoon et al., 2005). In this system, Bmpr1a function is ablated shortly after the formation of prechondrogenic condensations in appendicular elements. Analysis of the rudimentary condensations in these mice reveals a complete absence of expression of Sox9 (Yoon et al., 2005). Taken together, these results indicate that BMP signaling is not suYcient to initiate Sox9 expression prior to condensation, at least in appendicular elements, but is required for the maintenance of Sox9 expression in condensations. The molecular mechanisms underlying the ability of BMP signaling to maintain Sox9 expression remain elusive, but must be clearly identified to understand how BMPs induce and maintain commitment of cells to the chondrogenic lineage.


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Studies suggest that the gene for the transcription factor Barx2 is a downstream target of BMP signaling that is required to initiate chondrogenic condensation (Meech et al., 2005). Further exploration of the mechanisms by which BMPs control the expression of this gene prior to and during condensation is thus perhaps be very informative.

D. BMPs in Early Chondrocyte Differentiation The requirement for BMPs to maintain Sox gene expression is consistent with the results of numerous studies that demonstrate a continuous role for BMPs in chondrocytes. This aspect of BMP action has been studied extensively in ATDC5 chondrosarcoma cells. Treatment of these cells with BMPs results in acquisition of a cellular morphology characteristic of chondrocytes and upregulation of type II collagen production (Shukunami et al., 1998, 2000). Conversely, the overexpression of a DN‐type I BMP receptor or treatment with noggin reduces cartilage formation and type II collagen production (Ito et al., 1999; Shukunami et al., 2000). In vivo evidence demonstrating a persistent role for BMPs in chondrogenesis came initially from studies in chick limbs. Chondrocytes do not arise in precartilaginous cells in condensations in the absence of BMP signaling (Pizette and Niswander, 2000). Analyses of mice deficient in individual BMP ligands have not been particularly illuminating regarding their roles in skeletal development. As discussed earlier, there is a considerable degree of overlap in the expression of BMP ligands, very likely obscuring the full range of activities in which these ligands participate during skeletal development. Moreover, the expression of many BMPs is initiated at the onset of germ layer formation, and embryonic lethality at early stages has precluded analyses of their roles in the skeleton. However, some important findings have emerged from studies of mice and humans deficient in BMP ligands. Studies of the phenotypes of mice lacking Gdf5 and Bmpr1b reveal that GDF5 is essential for early stages of chondrogenesis in vivo, and that many of its eVects are transduced through BMPRIB. The brachypodism (bp) mutation in the mouse is a null allele for Gdf5. The primary phenotypic features are absence of phalangeal elements, shortening of other appendicular elements, and loss of interphalangeal joints (Storm et al., 1994). Although GDF5 is expressed in the region of developing joints and has been proposed to stimulate joint formation, it is also expressed in early condensations, is a potent stimulator of chondrogenesis in vitro (Erlacher et al., 1998), and leads to ectopic cartilage, tendon, and ligament formation when implanted subcutaneously (Wolfman et al., 1997). Moreover, rather than inducing ectopic joint formation, transgenic mice overexpressing Gdf5 exhibit cartilage overgrowth, consequent joint loss, and accelerated chondrocyte diVerentiation

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(Tsumaki et al., 1999, 2002). In the Gdf5 mouse, the loss of phalangeal elements appears to be due to apoptosis of mesenchymal cells in condensations, resulting in an insuYcient number of cells in the developing digital rays (Takahara et al., 2004). Hence, consistent with its expression in chondrogenic condensations, GDF5 appears to stimulate survival of chondroprogenitor cells, and to accelerate maturation of overt chondrocytes. Analyses of human chondrodysplasias have demonstrated that many of these functions, and the relationship between GDF5 action and BMPRIB signaling, are conserved. To date, loss‐of‐function and/or dominant‐negative mutations in Gdf5 have been found in patients with the Hunter‐Thompson form of acromesomelic chondrodysplasia (MIM 201250), Grebe‐type chondrodysplasia (MIM 200700), Du Pan syndrome (MIM 228900), autosomal dominant brachydactyly type C (MIM 113100), and brachydactyly type A2 (MIM 112600) (Faiyaz‐Ul‐Haque et al., 2002; Polinkovsky et al., 1997; Schwabe et al., 2004; Thomas et al., 1996). The first three of these syndromes are autosomal recessive, and the aVected allele leads to a loss of function. The phenotypes include extreme shortening of digits, and in more severe conditions, (e.g., Grebe‐type chondrodysplasia), profound shortening of long bones. Brachydactyly type C is less severe and can be either recessive or dominant. These varying phenotypes correlate well with the level of activity of the mutated GDF5 protein. Supporting a conserved relationship between GDF5 and BMPRIB, mutations in Bmpr1b cause the autosomal dominant syndrome brachydactyly type A2 in humans (Lehmann et al., 2003). In this case, the mutated receptor acts in a dominant‐negative manner. Mutations in Bmpr1b are also responsible for a case of autosomal recessive acromesomelic chondrodysplasia with genital anomalies (MIM 609441; Demirhan et al., 2005). In this case, the aVected allele contains a frameshift mutation leading to a truncated protein that is expected to result in a loss of function. This phenotype closely resembles that seen for mice lacking Bmpr1b (Baur et al., 2000; Yi et al., 2000). Thus, in both the mouse and human, loss of Bmpr1b has less severe consequences than does loss of Gdf5. This indicates that while BMPRIB is an essential receptor for GDF5, this ligand utilizes additional type I receptors to transduce its eVects in vivo. This type of cross talk, whereby a given BMP ligand transduces its eVects through multiple receptors, is also suggested by the phenotype of mice doubly deficient in Bmp7 and Bmpr1b. Only minor skeletal abnormalities are manifested in Bmp7/ mice, the most prevalent of which is polydactyly. However, the double‐mutant animals exhibit severe generalized chondrodysplasia, and complete loss of some skeletal elements (Yi et al., 2000). A mutation in Gdf5 has been described as the basis for proximal symphalangism (MIM 185800; Seemann et al., 2005). This phenotype resembles that phenotype seen in transgenic mice overexpressing Gdf5 in that joints are


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fused, and is thus distinct from those encountered in individuals harboring the loss‐of‐function and dominant‐negative Gdf5 mutations. Functional studies reveal that the Gdf5 allele underlying proximal symphalangism encodes a form of GDF5 that has an aYnity for BMPRIA that is equal to its aYnity for BMPRIB (Seemann et al., 2005); normally, GDF5 has approximately 12 times lower aYnity for BMPRIA. In summary, these findings demonstrate that GDF5 and BMPRIB play essential and conserved roles in early stages of chondrogenesis, regulating survival and proliferation. They also show that diVerential utilization of BMP receptors is required for normal skeletal development.

E. BMP Actions in the Growth Plate As previously discussed, numerous lines of evidence demonstrate that BMP signaling pathways are required for commitment to the chondrogenic lineage, and for survival and proliferation of condensed chondroprogenitors. In addition to these roles in early chondrogenesis, BMPs have important functions at later stages in the growth plate. As discussed, BMP ligands and receptors are expressed throughout the growth plate. That BMPs promote proliferation in the growth plate was first demonstrated by the observation that noggin‐null mice have overgrown skeletal elements (Brunet et al., 1998). Because noggin is an antagonist specific to the BMP pathway, this finding established that chondrocytes exhibit a proliferative response to endogenous BMPs in vivo. It also demonstrated that the correct balance between positive and negative controls on this response is essential. In accordance, overexpression of noggin results in smaller growth plates and reduced proliferation, while application of BMPs increases proliferation rates and growth plate size (De Luca et al., 2001; Minina et al., 2001; Pathi et al., 1999; Tsumaki et al., 2002). Noggin is highly expressed in regions of joint formation, suggesting that it is required there not only to prevent cartilage overgrowth but possibly also to induce a reversion of cells from a chondrogenic fate to a fibroblast‐like behavior, and to permit cell death leading to joint cavitation. This speculation is based on the findings that loss of BMP signaling in chondrocytes leads to loss of both Sox9 expression and the chondrocyte phenotype, increased apoptosis, and acquisition of expression of joint markers (Baur et al., 2000; Yi et al., 2000; Yoon et al., 2005). Consistent with this interpretation, in humans, heterozygous loss‐of‐function mutations in NOGGIN cause a form of proximal symphalangism (OMIM 185800), carpal–tarsal coalition syndrome (OMIM 186570), and multiple synostosis syndrome (OMIM 186500) (Gong et al., 1999; Seemann et al., 2005). All three conditions are characterized by the absence of joint cartilage and consequent fusions of the aVected elements.

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In addition to roles in regulation of proliferation and type II collagen expression, a major activity of BMPs is to promote hypertrophic diVerentiation. In ATDC5 cells and chick chondrocytes, BMPs increase the expression of type X collagen, the major ECM marker for hypertrophic chondrocytes (Grimsrud et al., 1999; Ito et al., 1999; Shukunami et al., 1998). This eVect is direct because BMPs upregulate type X collagen promoter activity (Volk et al., 1998). As previously discussed, BMPs 2 and 6 are expressed in the hypertrophic zone, and it has often been proposed that these BMPs regulate terminal diVerentiation of chondrocytes. To test this hypothesis, Kugimiya et al. (2005) created mice with the genotype Bmp2þ/;Bmp6/ (Bmp2/ mice die during the early embryonic phase and Bmp6/ mice exhibit no significant skeletal phenotype). Fetal Bmp2þ/;Bmp6/ mice have smaller growth plates, but there were no abnormalities in the progression of chondrocyte diVerentiation, or in expression of hypertrophic chondrocyte markers. However, the mice exhibit reduced bone formation. This suggests that the prominent expression of these BMPs in hypertrophic cells reflects a paracrine function on the osteoblast rather than an autocrine role in the hypertrophic chondrocyte. It is likely, however, that a role in terminal diVerentiation of hypertrophic chondrocytes might be revealed by analyses of mice completely conditionally deficient in both ligands. The roles of BMP pathways in hypertrophic diVerentiation appear to be complex in vivo. Exposure of metatarsal rudiments to noggin reduces Ihh expression and the length of the hypertrophic zone, suggesting that BMPs prevent premature exit from the cell cycle (De Luca et al., 2001; Minina et al., 2001; Naski et al., 1998; Pathi et al., 1999). On the other hand, Kobayashi et al. (2005) showed that overexpression of a constitutively active form of BMPRIA also leads to shortened columnar stacks of proliferating chondrocytes, with concomitant upregulation of maturational markers, suggesting that excessive BMP signaling causes premature onset of hypertrophic diVerentiation. In organ culture, the antagonist noggin also increased the expression of the maturation marker OPN, a marker for the most‐diVerentiated cells in the growth plate (Minina et al., 2001). The observation that both gain and loss of BMP activity can lead to upregulation of maturation markers probably reflects diVerent eVects of manipulating the BMP pathway at diVerent stages of proliferation and maturation. Thus, BMPs may promote the onset of diVerentiation, but their role in the most terminal stages of hypertrophy remains unclear. Mice lacking Bmpr1a and Bmpr1b in cartilage do not exhibit a complete arrest in diVerentiation; although the process is delayed, hypertrophic chondrocytes are clearly present along the edges of the rudimentary cartilaginous nodules (Yoon et al., 2005). At present it is unclear whether there is residual signaling through ALK2, the remaining type I BMP receptor; experiments


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are underway to test this hypothesis. Preliminary studies are consistent with this possibility; loss of Alk2 exacerbates the eVects of loss of either Bmpr1a or Bmpr1b, suggesting that ALK2 has at least some functions that overlap those of BMPRIA and BMPRIB (R. Pogue, S. Brugger, and K. Lyons, unpublished observations). The finding that chondrogenesis and hypertrophic diVerentiation are restricted to regions adjacent to the perichondrium in the highly abnormal cartilage in these mutants is intriguing. While the perichondrium is a rich source of BMPs, it is also the major source of other growth factors, such as Wnts, FGFs, and TGF‐ s that might promote alternative pathways leading to chondrocyte diVerentiation. Additional experiments involving ablation of BMP pathways in defined zones of the growth plate will certainly be needed to clarify the role of BMP pathways at distinct stages of chondrocyte diVerentiation.

VII. Downstream Effectors of BMP Signaling: Smads and p38 As discussed previously, BMPs mediate their eVects through Smad‐mediated and non‐Smad–mediated pathways (Zhang et al., 2003). Thus, determining the extent to which these alternative pathways are utilized in specific aspects of chondrogenesis is a vital issue.

A. Smad‐Mediated Signaling in Chondrogenesis While the potential importance of canonical Smad1/5/8 and p38 pathways in chondrogenesis is established, the roles that these pathways play in regulating specific aspects of chondrogenesis remain unclear. Neither the role of the p38 pathway in cartilage nor the extent to which p38 is activated by BMP signaling in vivo are known. Even the roles of canonical Smad pathways in vivo in chondrogenesis are unknown. Phosphorylated (activated) Smad1/ 5/8 is abundant in early prechondrogenic condensations, and is absent when BMP receptors are ablated (Yoon et al., 2005). However, overexpression of BMP‐specific Smads does not promote eYcient cartilage formation in C3H10T1/2 and ATDC5 cells, but does so in MC615 chondroprogenitor cells (Fujii et al., 1999; Hatakeyama et al., 2003; Ju et al., 2000). The extent to which these discordant findings reflect intrinsic diVerences in the diVerent cell lines, a requirement for noncanonical pathways downstream of BMPs in addition to Smad1/5/8, sensitivity to the BMP signal, or technical diYculties inherent in overexpressing Smads, remains unknown. Loss‐of‐function approaches using mice carrying floxed alleles of Smads 1 and 5 are ongoing to address these issues.

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Smad4 (also known as common partner Smad or co‐Smad) has been found complexed to R‐Smads in all Smad target gene promoters analyzed by chromatin immunoprecipitation to date (Massague et al., 2005). However, whether Smad4 is required for all Smad‐mediated signaling events is unclear. Smad4‐deficient cells retain some responsiveness to TGF‐ (Levy and Hill, 2005; Sirard et al., 1998; Subramanian et al., 2004; Wisotzkey et al., 1998), but the level of responsiveness varies with the cell type. Moreover, the phenotype of mice lacking Smad2 is more severe than that of mice lacking Smad4, and loss of Smad4 in the mouse epiblast demonstrated that only a subset of TGF‐ /activin/nodal responses require Smad4 (Chu et al., 2004; Nomura and Li, 1998; Sirard et al., 1998). The phenotypes of mice lacking the BMP Smads, Smad1/5/8 are less severe than those of Smad4/ mice (Chang et al., 1999; Hayashi et al., 2002; Lechleider et al., 2001; Tremblay et al., 2001), but this may be a consequence of functional redundancies among these R‐Smads. Zhang et al. (2005), utilizing Col2a1‐Cre and a floxed allele of Smad4, found that Smad4 is essential for normal organization of the growth plate. In the cartilage‐specific Smad4 mutants, the growth plate becomes severely disorganized during postnatal stages. Irregular patches of chondrocytes occasionally give rise to columnar zones whose orientation bears little relationship to the long axis of the skeletal element in these mice. This phenotype does not resemble that of Tgf 2/ mice nor of mice lacking Tgf r2 in cartilage (Alvarez and Serra, 2004). The Smad4;Col2a1‐Cre phenotype is also in striking contrast to that of mice lacking Bmpr1a and Bmpr1b in cartilage, where chondrogenesis is essentially arrested prior to and during the early condensation stage (Yoon et al., 2005). These findings raise the possibility that BMP‐mediated signaling through p38 might explain the more profound eVects seen in Bmpr1a;Bmpr1b mutants. Alternatively, BMP Smads might transduce some eVects in early condensations independently of Smad4. Analysis of mice harboring cartilage‐specific loss of pathway‐restricted R‐Smads will be needed to discriminate among these possibilities. Some of the best evidence for involvement of canonical BMP Smad pathways in aspects of chondrocyte diVerentiation comes from analysis of the chicken type X collagen promoter (Volk et al., 1998). Type X collagen is expressed in prehypertrophic and hypertrophic chondrocytes and is an essential constituent of the matrix in this region of the growth plate. Runx2 and Runx3, members of the Runt‐related transcription factor family, are also expressed in prehypertrophic and hypertrophic chondrocytes (Kim et al., 1999; Yoshida et al., 2004). A requirement for these factors in terminal diVerentiation is demonstrated by the finding that Runx2/; Runx3/ mice exhibit a complete absence of prehypertrophic and hypertrophic chondrocytes (Yoshida et al., 2004). Runx2 (a transcriptional coactivator for Smads) and Smad1 bind directly to the type X collagen promoter and their


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combined activities are required for its expression (Drissi et al., 2003; Zheng et al., 2003). The Ihh gene represents a second example of a potential direct target of BMP Smad pathways in chondrocyte diVerentiation. As previously discussed, Ihh is expressed in prehypertrophic chondrocytes and it regulates the onset of terminal diVerentiation by controlling levels of PTHrP expression (Kronenberg, 2003). Runx2 binds directly to and activates the Ihh promoter in vitro (Yoshida et al., 2004). A study using a ChIP‐based cloning method demonstrated that the Ihh gene is also a direct target of BMPs, and that the promoter region contains multiple motifs that bind to Smad4 and are required for BMP‐dependent activation (Seki and Hata, 2004). A microarray analysis of BMP‐induced gene expression in chondrocytes identified GADD45 as an immediate early target of BMP signaling; BMP‐ induced Smad1/Runx2 activity was shown to upregulate expression of GADD45 , which is found in the nuclei of late hypertrophic chondrocytes, and is an activator of expression of Mmp13. Inhibition of GADD45 delays terminal chondrocyte diVerentiation (Goldring et al., 2006; Ijiri et al., 2005). These findings suggest that GADD45 may be a key target through which BMPs induce terminal diVerentiation of hypertrophic chondrocytes. Taken together, these findings suggest that coordinated regulation of a battery of genes by BMP‐Smads and Runx2 may be a characteristic feature of hypertrophic diVerentiation.

B. p38 Pathways in Chondrogenesis As discussed previously, in addition to activation of Smad‐mediated pathways, members of the TGF‐ superfamily can activate alternative pathways (Moustakas and Heldin, 2005). In chondrocytes, the most extensively studied of these is the p38 pathway. BMPs can aVect chondrocytes by activating p38 MAPK through the TGF‐ –activated kinase, TAK1 (Hatakeyama et al., 2003; Ju et al., 2000; Nakamura et al., 1999; Qiao et al., 2005; Seto et al., 2004). Transgenic mice expressing constitutively active MKK6, which activates p38, exhibit dwarfism with delayed endochondral ossification, reduced chondrocyte proliferation, and inhibition of hypertrophy (Zhang et al., 2006). Although it is unclear how, if at all, this result relates to p38 mediation of BMP signaling in the growth plate, it is clear that activated p38 has the potential to greatly influence chondrocyte development. Furthermore, the study by Qiao et al. (2005) indicated that signaling through TAK1 could increase levels of type II collagen but that this did not require increased transcription of the Col2a1 gene. This demonstrates that as well as influencing gene expression, BMP and TGF‐ signaling also exerts considerable eVects in the growth plate at the posttranscriptional level.

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In ATDC5 cells and micromass cultures, BMP treatment leads to sustained phosphorylation of p38. In some studies, the ability of BMPs to promote chondrogenesis may require p38, as p38 inhibitors strongly suppress induction of type II collagen and chondrogenic diVerentiation (Nakamura et al., 1999; Zuzarte‐Luis et al., 2004). Reporter assays employing the type X collagen promoter in chondrocytes also suggest a requirement for p38 in BMP responses; p38 induces activity through the same region that had previously been shown to be a BMP‐responsive element. They suggested that p38 acts with Smads as a coactivator of the ColX promoter (Reilly et al., 2005). At present, while these studies support an essential role for the p38 in chondrogenesis, and in at least some activities mediated by BMPs, it is as yet unclear whether p38 acts in parallel or downstream of Smads. In vivo studies will be required to elucidate the extent to which these pathways mediate distinct aspects of chondrogenesis and the extent to which they interact.

VIII. Interactions with Other Signaling Pathways An important aspect of the control of chondrocyte proliferation and maturation in the growth plate involves interactions among signaling pathways (Fig. 4). This cross talk integrates the signals that promote and inhibit proliferation, hypertrophy, and apoptosis (Goldring et al., 2006; Kronenberg 2003; Yoon and Lyons, 2004).

A. Synergy Between BMP and IHH/PTHrP Pathways As previously discussed, Ihh is expressed by prehypertrophic chondrocytes, and loss of Ihh leads to reduced chondrocyte proliferation and premature diVerentiation (Kronenberg, 2003). This phenotype arises because IHH is required for PTHrP expression in the periarticular region of the growth plate. BMPs interact with the IHH/PTHrP pathway by promoting Ihh expression. Exposure of cartilage to noggin in organ culture reduces Ihh expression (Minina et al., 2001; Pathi et al., 1999). The demonstration that the Ihh promoter contains multiple Smad‐binding motifs and is activated by BMP treatment suggests that this regulation is direct (Seki and Hata, 2004). In turn, IHH maintains BMP levels, indicating the existence of a positive feedback loop (Minina et al., 2001; Pathi et al., 1999). This eVect may be direct because the downstream mediators of IHH signaling, Gli transcription factors, directly upregulate Bmp4 and Bmp7 promoter activity (Kawai and Sugiura, 2001), and a key role for IHH is to counteract the activity of repressive forms of Gli proteins (Hilton et al., 2005; Koziel et al., 2005).


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Figure 4 Interactions between BMPs and other signaling pathways in the growth plate. BMPs promote chondrocyte proliferation, and onset of hypertrophy, but may inhibit terminal diVerentiation. BMP promotes proliferation through direct eVects on chondrocytes as well as by upregulating Ihh expression, which in turn, induces PTHrP. IHH induces proliferation indirectly through upregulation of PTHrP as well as directly on proliferating chondrocytes. BMPs and IHH mutually upregulate each other’s expression. IHH works in synergy with BMP to promote hypertrophy, at least in part through regulation of expression of markers of chondrocyte terminal diVerentiation, such as Runx2 and osteocalcin. However, the role of BMP pathways in clearance of late hypertrophic chondrocytes is unclear. FGFs antagonize BMPs by multiple mechanisms. FGF and BMP have opposing eVects on proliferation, survival, and Ihh expression through unknown mechanisms, some of which may be direct eVects on Smad activity, and some of which may be indirect via regulation of expression of ligands, receptors, and signaling components of these pathways. Whether BMPs and FGFs have opposing and/or cooperative eVects on terminal hypertrophic diVerentiation is as yet unclear. FGFs promote some aspects of terminal diVerentiation, such as osteocalcin expression, but inhibit others, such as IHH expression.

In addition to this positive feedback loop, BMPs and IHH act independently of each other to regulate chondrocyte proliferation, and IHH regulates PTHrP expression independently of BMP signaling (Minina et al., 2001). There is also evidence for synergy between BMP and IHH signaling pathways. In a limb bud cell line, although IHH on its own cannot induce Runx2 and osteocalcin expression, treatment with a blocking antibody against IHH inhibits BMP‐induced expression of these genes (Long et al., 2004). Finally, immunoprecipitation studies show that Smad1 directly

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associates with truncated (activator) forms of Gli3 proteins, suggesting a possible mechanism for this synergy (Liu et al., 1998).

B. Antagonistic Interactions Between BMP and FGF Pathways As discussed previously, many of the eVects of FGFs in the growth plate are opposite to those of BMP signaling, in that excessive FGF signaling inhibits proliferation and enhances the rate of hypertrophy, whereas excessive BMP activity generally promotes proliferation and delays the onset of terminal diVerentiation (Kronenberg 2003, Minina et al., 2002). Functional antagonism between BMP and FGF signaling pathways is confirmed in limb culture studies. BMP treatment rescues the phenotype of FGF‐treated growth plates, and FGF treatment neutralizes the eVects of BMPs (Minina et al., 2002). The mechanisms underlying this antagonism are not understood. Several studies indicate that FGFs and BMPs mutually regulate the expression of signaling components in the opposing pathway. However, whether this regulation is positive or negative is unclear. Overexpression of activated FGFR3 inhibits Bmp4 expression in postnatal mouse growth plates (Naski et al., 1998) but induces Bmp4 and Bmp7 in embryonic growth plates, where BMP treatment also induces Fgf18 expression (Minina et al., 2002). These opposing results could be due to diVerences in the pathways as a function of age diVerences in the mice, as suggested by the researchers (Minina et al., 2002), or to diVerences in the methodologies used to alter the levels of pathway activation (e.g., chronic eVects due to genetic alterations versus short‐term eVects to acute exposure to nonphysiological levels of growth factors). Understanding the basis for these diVerent results will perhaps lead to insights regarding the relationship of these two pathways to each other. In principle, many of the opposing activities of BMPs and FGFs can be explained by the finding that BMPs promote Ihh expression, whereas FGFs inhibit it (Minina et al., 2002; Naski et al., 1998). As previously discussed, Ihh is a direct target of BMP Smads, although whether this is the case in chondrocytes has not been demonstrated (Seki and Hata, 2004). The mechanisms by which FGF pathways antagonize BMP‐induced Ihh expression are unknown. One potential mechanism is suggested by the discovery that the linker region of R‐Smads contains multiple consensus sites for phosphorylation by ERK1/2, a known component of the FGF‐induced growth arrest in cartilage (Aikawa et al., 2001; Kretzschmar et al., 1997; Murakami et al., 2004; Raucci et al., 2004). When these consensus sites are phosphorylated, Smad activity is inhibited. These data are in agreement with the results of Reilly et al. (2005), who found that inhibition of ERK1/2 increased BMP‐ stimulated activation of the ColX promoter. However, to date, a role for


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linker phosphorylation in regulating BMP output in vivo has only been demonstrated in the context of neural induction in Xenopus (Delaune et al., 2005; Kuroda et al., 2005; Pera et al., 2003) and the relevance of these studies to mammals has been called into question (Aubin et al., 2004). Aubin et al. (2004) generated mice that are resistant to phosphorylation in the linker region of Smad1. These mice do not exhibit chondrodysplasia, suggesting that phosphorylation of the linker region is not required for normal chondrogenesis. However, the mutated allele behaved as a hypomorph, most likely as a result of mistargeting of the mutated protein (Aubin et al., 2004). This mislocalization could conceivably negate any increased activity of Smad1L that would have been expected by its resistance to ERK1/2‐mediated inhibition of transcriptional activity. These observations are consistent with several interpretations. First, contrary to the findings of others (Kretzschmar et al., 1997; Kuroda et al., 2005; Pera et al., 2003), ERK1/2‐mediated linker phosphorylation of Smad1 may be irrelevant in most physiological contexts, including chondrogenesis. According to this scenario, functional diVerences between WT and linker mutant forms of Smad1 can only be seen in cases where these forms of Smad1 are overexpressed. Second, it is possible that the eVects of linker phosphorylation are most relevant in vivo in contexts where ERK1/2 activity is abnormally elevated, as is the case in achondroplastic dwarfism caused by activating mutations in Fgfr3. Additional in vivo studies are clearly needed to evaluate the role this mechanism plays in FGF‐mediated antagonism of BMP pathways. How might BMP pathways inhibit FGF activity? In addition to regulating expression of FGFs and their receptors (Minina et al., 2002; Naski et al., 1998), the finding that FGF eVects in the growth plate are counteracted by C‐type natriuretic peptide (CNP) (Krejci et al., 2005) invites speculation that BMPs may be involved. Although this has not been directly investigated in relation to chondrocyte proliferation and diVerentiation, Yeh et al. (2006) showed that CNP can enhance BMP7‐induced Smad5 phosphorylation in osteoblast diVerentiation.

C. The Balance Between TGF‐b and BMP Pathways in Osteoarthritis There is also strong evidence to show that the balance between the BMP and TGF‐ signaling pathways is important. Sternal chondrocytes were examined from Smad3/ mice (Li et al., 2006). In these cells, higher levels of endogenous BMP signaling activity were observed, correlated with increased levels of Bmp2 and Bmp6 expression, elevated basal levels of phosphorylated Smads 1 and 5, and concurrent upregulation of ColX expression, indicating inappropriate maturation of the articular chondrocytes. Such unchecked

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BMP signaling in the articular region could contribute to diseases such as osteoarthritis by promoting diVerentiation of articular chondrocytes. It has been shown that in osteoarthritic articular chondrocytes, there is upregulation of Smurf2, which targets Smads 2 and 3, again indicating a shift of equilibrium toward BMPs and diVerentiation (Zuscik et al., 2004). The potential importance of this balance between BMP and TGF‐ pathways with respect to osteoarthritis is reviewed by Drissi et al. (2005). Understanding how this antagonistic relationship is achieved on a mechanistic level, whether it is through competition for common signaling components (e.g., type II receptors, Smad4, and so on), through activation of opposing downstream pathways and transcriptional targets, and/or through interactions of TGF‐ and BMP receptors on the cell surface, is an issue of importance. This understanding may permit the manipulation of these pathways to eVect repair of articular cartilage.

IX. Perspectives The BMP signaling cascade has been shown in a wide range of studies to be crucial to the proper patterning and growth of the skeleton. Through precisely regulated signaling events, involving cross talk with other pathways, BMPs regulate proliferation and maturation of chondrocytes, and this ultimately determines the size and shape of endochondral skeletal elements. Emerging evidence also points to a role for the TGF‐ and BMP pathways in maintenance of articular cartilage in adults. There are many questions regarding BMP function in all stages of the development and function of the growth plate that remain to be addressed. As mesenchymal condensations mature, osteochondroblasts at the edges of the condensations are directed toward a perichondrial fate as opposed to that of a growth plate chondrocyte. The mechanisms that trigger this diVerential development are not understood. Hence, whether BMP and/or TGF / activin/nodal pathways contribute to this process is unknown. The expression of abundant ligands for BMP and TGF‐ pathways make the perichondrium a potential source of morphogen gradients that may contribute to the organization and orientation of chondrocyte columns within the growth plate. The mechanisms guiding cell shape changes as chondrocytes progress through the growth plate are also poorly understood. The ECM is known to play a major role, but whether signaling factors such as BMPs are also involved is unknown. As discussed, proliferating chondrocytes near the top of the CZ display the highest rate of proliferation, and the rate decreases as the columnar chondrocytes approach the prehypertrophic zone. It is possible that the change in proliferation rate may be linked to the cell shape changes


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that mark the transition from round proliferating chondrocytes to flattened columnar proliferating chondrocytes. The trigger for the cell shape change may involve the actions of BMPs. A related issue is the control of chondrogenesis along the lateral (closest to the perichondrium) to medial (interior of the growth plate) axis. As discussed previously, analyses of several mouse mutants have revealed that loss of BMPs aVects chondrogenic cores to a greater extent than chondrogenesis closest to the perichondrium (Baur et al., 2000; Yi et al., 2001; Yoon et al., 2005), and Minina et al. (2005) have shown that Bmp7 is expressed in a medial to lateral gradient. Understanding the mechanistic basis for these findings may reveal diVerences in signaling pathways and/or their interactions along the lateral‐medial axis. To address such a possibility, it will be necessary to determine more finely the patterns of expression of BMPs and their receptors in the perichondrium, and to determine the extent to which these factors diVuse from their sites of production, thereby potentially forming lateral to medial signaling gradients. The distinct expression patterns of the BMPs and their receptors imply that their induction must be tightly controlled. A number of the animal model studies described here indicate that BMPs and their receptors are capable of compensating for each other to a considerable degree. However, the mechanisms by which the cells sense the deficiency of one protein, and employ another to compensate are not understood. These issues need to be examined through analyses of the control regions of the genes for BMPs and their receptors. This will require further use of methods such as the BAC‐ transgenic approach employed by DiLeone et al. (2000) and Mortlock et al. (2003). While several key targets of BMP signaling pathways in the growth plate, such as Ihh, have been identified, there is little information regarding those genes whose expression is regulated directly by BMPs in the growth plate and is critical to the biological activities of BMPs. In particular, identification of additional targets of the BMP pathway will shed light on the mechanisms by which BMPs control terminal diVerentiation and apoptosis of hypertrophic chondrocytes. It is becoming increasingly clear that a large number of proteins, many of which function in multiple signaling pathways, interact with BMP pathways in synergistic and antagonistic manners. For example, as discussed, extracellular matrix components can exert both positive and negative influences on BMP signaling. An important area of investigation will be exploring the extent to which ECM components regulate ligand availability and diVusion from the perichondrium versus regulation of intracellular mediators of the BMP pathway. That latter possibility is exemplified by Peterson et al. (2004), who demonstrated that the ECM can directly influence intracellular elements of the BMP pathway, as was shown by the regulation of Smads by

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hyaluronan via CD44. It is very likely that other extracellular molecules are directly connected to intracellular elements of BMP pathways. As discussed, it has been hypothesized that the diVerences in lengths of long bones are related to diVerences in cell sizes between growth plates (Ballock and O’Keefe, 2003; Wilsman et al., 1996). While BMPs are known to be involved in the onset of hypertrophy, their role in controlling the extent to which cell size increases is unknown. It is likely that, as is the case elsewhere in the growth plate, this process requires cooperation and reciprocal modulation between BMP and other signaling cascades. Less is known about the extent to which pathway usage varies between diVerent bones. It is widely assumed that BMPs and other signaling pathways play identical roles in all skeletal elements. However, the fact that growth plates in these elements exhibit diVerent rates of proliferation and diVerentiation implies that there are at least some diVerences in pathway utilization. Identifying the basis for these diVerences will be essential to understanding the evolution of skeletal form. Additional issues that require further attention concern the degree to which BMPs employ canonical Smad versus noncanonical pathways in the growth plate, as well as the circumstances under which these pathways act in synergy or antagonistically. Answers to these questions will require complimentary genetic analyses employing in vitro and in vivo models for each pathway, and will help to address the mechanisms by which the BMP ligands and receptors achieve diversity in the results of signaling events. The recent report by Qiao et al. (2005) stating that BMPs and TGF‐ were capable of acting through TAK1 to increase type II collagen levels, but by a method that did not appear to require de novo transcription of the Col2a1 gene raises new questions regarding virtually unexplored posttranscriptional modes of action by BMPs and TGF‐ . How is information on BMP and TGF‐ signaling in the growth plate being applied to treatment of human trauma and disease, in particular osteoarthritis? Many studies are addressing the potential role of manipulating the BMP pathways in orthopedic clinical treatments. Methods of BMP‐ based gene and cell therapy are actively being investigated in relationship to repair of cartilage (Drissi et al., 2005; Luyten, 2004; Wu et al., 2003). Some of these involve the use of BMPs to treat mesenchymal stem cells, inducing chondrogenesis before implantation. It is widely assumed that growing knowledge regarding the targets of BMP pathways will be paired with stem cell technology to produce eVective therapies that will set new standards for tissue repair and regeneration. For example, a study has shown that the antiepileptic drug phenytoin, which has already been known to aVect bone formation, can induce cartilage nodule formation in ATDC5 cells, and that this induction is accompanied by upregulation of Tgf 1, Bmp4, Ihh, and the IHH receptor, Ptc (Okada et al., 2005).


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The Tgf 1upregulation may be key, since as discussed earlier, levels of TGF‐ signaling, as measured by nuclear Smad2 levels in articular cartilage, decline with age, potentially leading to a loss of the protective TGF‐ influence. This and many similar studies form the basis for discovery of drugs that can upregulate the correct combination of signaling molecules for cartilage and/or bone repair (for reviews of drug and growth factor treatment strategies for cartilage see Holland and Mikos, 2003; Seeherman and Wozney, 2005; Verbruggen, 2006).

Acknowledgments The authors would like to thank Jessica Chen and Bonny Lee for assistance with literature searches.

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Zhang, D., Ferguson, C. M., O’Keefe, R. J., Puzas, J. E., Rosier, R. N., and Reynolds, P. R. (2002). A role for the BMP antagonist chordin in endochondral ossification. J. Bone Miner. Res. 17, 293–300. Zhang, D., Schwarz, E. M., Rosier, R. N., Zuscik, M. J., Puzas, J. E., and O’Keefe, R. J. (2003). ALK2 functions as a BMP type I receptor and induces Indian hedgehog in chondrocytes during skeletal development. J. Bone Miner. Res. 18, 1593–1604. Zhang, J., Tan, X., Li, W., Wang, Y., Wang, J., Cheng, X., and Yang, X. (2005). Smad4 is required for the normal organization of the cartilage growth plate. Dev. Biol. 284, 311–322. Zhang, R., Murakami, S., Coustry, F., Wang, Y., and de Crombrugghe, B. (2006). Constitutive activation of MKK6 in chondrocytes of transgenic mice inhibits proliferation and delays endochondral bone formation. Proc. Natl. Acad. Sci. USA 103, 365–370. Zheng, Q., Zhou, G., Morello, R., Chen, Y., Garcia‐Rojas, X., and Lee, B. (2003). Type X collagen gene regulation by Runx2 contributes directly to its hypertrophic chondrocyte‐ specific expression in‐vivo. J. Cell Biol. 162, 833–842. Zou, H., Wieser, R., Massague, J., and Niswander, L. (1997). Distinct roles of type I bone morphogenetic protein receptors in the formation and diVerentiation of cartilage. Genes Dev. 11, 2191–2203. Zuscik, M. J., Baden, J. F., Wu, Q., Sheu, T. J., Schwarz, E. M., Drissi, H., O’Keefe, R. J., Puzas, J. E., and Rosier, R. N. (2004). 5‐Azacytidine alters TGF‐beta and BMP signaling and induces maturation in articular chondrocytes. J. Cell. Biochem. 92, 316–331. Zuzarte‐Luis, V., Montero, J. A., Rodriguez‐Leon, J., Merino, R., Rodriguez‐Rey, J. C., and Hurle, J. M. (2004). A new role for BMP5 during limb development acting through the synergic activation of Smad and MAPK pathways. Dev. Biol. 272, 39–52.

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The CLIP‐170 Orthologue Bik1p and Positioning the Mitotic Spindle in Yeast Rita K. Miller,* Sonia D’Silva,* Jeffrey K. Moore,* and Holly V. Goodson{ *Department of Biology, University of Rochester Rochester, New York 14627 { Department of Chemistry and Biochemistry The University of Notre Dame Notre Dame, Indiana 46556

I. Introduction II. Structure of the Bik1 Protein A. Head Domain B. Coiled‐Coil Domain C. Cargo‐Binding Domain III. Bik1p Interactions with Other Proteins IV. Phenotypes and Activities V. Role of Bik1p in Spindle Positioning A. Genetics of Spindle Positioning B. Bik1p in the Dynein Pathway C. Bik1p and the Kar9p Pathway D. Bik1p in Mating Cells VI. Models for Plus‐End Binding A. Microtubule Plus‐End Tracking (þTIP) Behavior B. Mechanisms of Microtubule Plus‐End Tracking Behavior C. Interactions Between EB1 and CLIP‐170 Versus Bim1p and Bik1p VII. Regulation of CLIP‐170 and Bik1p by Phosphorylation VIII. Bik1p at the Kinetochore IX. Bik1p in Genomic Studies A. Synthetic Lethal Interactions B. Gene Expression Levels C. Quantitation of Bik1p Molecules in the Cell X. Concluding Remarks and Perspective Acknowledgments References

Bik1p is the yeast Saccharomyces cerevisiae representative of the CLIP‐ 170 family of microtubule plus‐end tracking proteins. Bik1p shares a number of similarities with its mammalian counterpart CLIP‐170, including an important role in dynein function. However, Bik1p and CLIP‐170 diVer in several significant ways, including the mechanisms utilized to track Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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0070-2153/06 $35.00 DOI: 10.1016/S0070-2153(06)76002-1

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microtubule plus ends. In addition to presenting functional comparisons between Bik1p and CLIP‐170, we provide sequence analyses that reveal previously unrecognized similarities between Bik1p and its animal counterparts. We examine in detail what is known about the functions of Bik1p and consider the various roles that Bik1p plays in positioning the yeast mitotic spindle. This chapter also highlights several recent findings, including the contribution of Bik1p to the yeast mating pathway. ß 2006, Elsevier Inc.

I. Introduction Bik1p has captivated researchers for nearly 20 years. It was originally identified from its bilateral karyogamy phenotype, displaying a defect in the nuclear fusion step of the mating pathway in the budding yeast, Saccharomyces cerevisiae (Berlin et al., 1990; Kurihara et al., 1994; Trueheart et al., 1987). Early experiments indicated that Bik1p plays a critical role in the regulation of microtubule dynamics (Berlin et al., 1990), microtubules being one of the fundamental cytoskeletal polymers within the cell. This involvement in microtubule dynamics is conserved in orthologues of Bik1p from other organisms, including mammalian CLIP‐170 and S. pombe Tip1p. Like CLIP‐170 and Tip1p, Bik1p is a ‘‘microtubule plus‐end tracking protein’’ (þTIP); it dynamically tracks the growing plus ends of microtubules (Carvalho et al., 2004). With these common attributes, Bik1p serves as a model for study of þTIP function and mechanism (Carvalho et al., 2003; Schuyler and Pellman, 2001). Like its CLIP‐170 counterpart, Bik1p also interacts with the dynein machinery (Dujardin et al., 1998; Goodson et al., 2003; Sheeman et al., 2003; Tai et al., 2002; Vaughan et al., 1999). It has become clear that Bik1p interacts with a number of additional proteins including Bim1p/EB1 (Wolyniak et al., 2006; Miller, unpublished observations), the MAP215 homologue Stu2p (Chen et al., 1998; Lin et al., 2001), Kar9p (Moore et al., 2006), and the TOR kinases (Choi et al., 2000). Thus, Bik1p has a number of intriguing facets to its biology.

II. Structure of the Bik1 Protein The Bik1p protein (51 kDa) is comprised of three domains, an N‐terminal head domain which contains a CAP‐Gly motif and a serine‐rich region, a central coiled‐coil region, and a C‐terminal region sometimes called the ‘‘cargo‐binding’’ domain (Fig. 1). On the basis of this combination of domains, Bik1p is considered to be the yeast orthologue of the mammalian CLIP‐170 protein, although there are some significant diVerences between the two proteins, as described below.

2. Bik1p, the Yeast CLIP‐170

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Figure 1 Bik1p belongs to a conserved family of CAP‐Gly proteins. Bik1p contains four distinct domains: the CAP‐Gly domain, the serine‐rich domain, the central coiled‐coil, and a C‐terminal cargo‐binding domain. Adapted from Brunner and Nurse, 2000.

A. Head Domain 1. CAP‐Gly Domain The CAP‐Gly domain (Fig. 2) is so named because it is found in cytoskeletal associated proteins and is glycine rich (Riehemann and Sorg, 1993). CAP‐Gly domains are conserved throughout evolution in organisms spanning from yeast to humans. Their existence can be documented in all fully sequenced eukaryotic genomes presently in GENBANK (Goodson, unpublished analysis). In addition to being found in CLIP‐170–related proteins, CAP‐Gly domains are found in several other highly conserved proteins including the p150 subunit of the dynactin complex (also known as dynactin, which regulates the dynein motor protein), and the tubulin folding factors cofactor B and E, which are specific for alpha‐tubulin (Grynberg et al., 2003; Li et al., 2002; Lopez‐Fanarraga et al., 2001; Lytle et al., 2004; Vadlamudi et al., 2005). In S. cerevisiae, Nip100p is the orthologue of p150Glued (Kahana et al., 1998). Alf1p is the orthologue of cofactor B (Feierbach et al., 1999). Pac2p is the orthologue of cofactor E (Fleming et al., 2000; Hoyt et al., 1997). Tubulin cofactors are thought to act by binding to tubulin monomers and assisting in the final stages of folding necessary to bring about heterodimer formation. CAP‐Gly motifs are also found in more phylogenetically restricted proteins, including Drosophila kinesin‐73 (Li et al., 1997) and CYLD, the human familial cylindromatosis tumor suppressor protein CYLD (Saito et al., 2004). Another CAP‐Gly containing protein is the CLIPR59 protein (Perez et al., 2002), which in contrast to other CLIPs, is not found along microtubules but is instead localized on Golgi‐associated vesicles. Its localization there is mediated by palmitoylation at its C‐terminus (Lallemand‐Breitenbach et al., 2004; Perez et al., 2002).

Figure 2 Alignment of the CAP‐Gly domain in Bik1p from S. cerevisiae with its fungal orthologues. The sequences of the two CAP‐Gly motifs of human CLIP‐170 and the single CAP‐Gly of human p150 are provided for comparison. Genbank sequences are abbreviated by the first four letters of the genus and two letters of the species, followed by the gene name or NCBI gi number. Sequences starting with MIT or WashU were obtained from the Saccharomyces Genome Database (SGD). The sequences of the two CAP‐Gly motifs of human CLIP‐170 and the single CAP‐Gly of human p150 are provided for comparison. The alignment was performed using ClustalX (Thompson et al., 1997) and shaded using the EMBNet Boxshade server (http://www.ch.embnet.org/software/BOX_form.html), both using default settings.


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CAP‐Gly domains are an important characteristic of proteins of the CLIP‐ 170 family and thus are found in all discernable CLIP‐170 relatives, including Tip1p of S. pombe (Brunner and Nurse, 2000); CLIP‐190 of Drosophila melanogaster (Dzhindzhev et al., 2005); and the mammalian CLIP‐115 protein (Akhmanova et al., 2001; Hoogenraad et al., 2000). Restin is a splice isoform of CLIP‐170, which contains a 35 amino acid insert (Sahin et al. 2002). Restin is found at high levels in the Reed–Sternberg cells of Hodgkin’s lymphomas (Bilbe et al., 1992). The CAP‐Gly sequence is generally considered to be a microtubule‐binding domain. Alteration or deletion of the CAP‐Gly domain interferes with the microtubule‐binding ability of both p150 and CLIP‐170 (Pierre et al., 1992, 1994; Puls et al., 2003; Waterman‐Storer et al., 1995). However, it is important to note that this domain binds to tubulin in cofactors B and E (Feierbach et al., 1999; Tian et al., 1999), and it is implicated in the high‐aYnity binding of CLIP‐170 to tubulin (Folker et al., 2005; Folker and Goodson, unpublished observations). Based on these characterizations, the CAP‐Gly domain in Bik1p is hypothesized also to be both a microtubule and tubulin‐binding domain, but this has not yet been demonstrated using purified Bik1p in vitro. Whereas CLIP‐170 has two CAP‐Gly domains, Bik1p has only one. The significance of this diVerence, particularly in terms of polymer and tubulin dimer‐binding aYnities, remains to be determined (see below). A chimeric Bik1p in which the mammalian CAP‐Gly domain of CLIP‐170 replaced the CAP‐Gly domain of Bik1p was found to be functional, localizing like the native yeast protein (Carvalho et al., 2004). Whether the converse chimera will be similarly functional is not known. These findings suggest that there is functional conservation of this domain across evolution. In the sequence alignment of Fig. 2, it is clear that the glycine residues of the Bik1p CAP‐Gly domain are well conserved. However, examination of this alignment and more in‐depth phylogenetic analysis of the family of CAP‐Gly motifs have not been able to establish whether the single CAP‐Gly domain found in Bik1p is more like the first or second CAP‐Gly found in CLIP‐170 (Fig. 2; Gregoretti and Goodson, unpublished analysis). Three independent structures now exist for the CAP‐Gly motif, one for the Caenorhabditis elegans cofactor B (Li et al., 2002), one for human p150 complexed with EB1 (Hayashi et al., 2005), and one for the much more divergent third CAP‐Gly of the human CylD protein (Saito et al., 2004). These structures contain a novel tertiary structure consisting of one alpha‐ helix and three beta sheets (Fig. 3) (Li et al., 2002). The striking similarity between the cofactor B and p150 structures (Fig. 3A and B) suggests that the CAP‐Gly motifs of Bik1p and other CLIP‐170 relatives will be very similar. Examination of these structures shows that the highly conserved GKNDG sequence lies on the surface of the molecule (Fig. 3C; Li et al., 2002). Point mutations within this region result in the mislocalization of Bik1p and

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Figure 3 Structure and conservation of the CAP‐Gly motif. (A, B) X‐ray structures of the CAP‐Gly motifs of proteins tubulin folding cofactor B (A) and dynactin p150 (B). The striking similarity between these proteins suggests that the CAP‐Gly motifs of Bik1p and its orthologues will have similar structures. (C) Conservation of the CAP‐Gly motifs of fungal Bik1p homologues mapped onto the p150 structure shown as in ribbon (left) or space full (right) format. The numbers of selected S. cerevisiae Bik1p residues are given for orientation: those in black are diVerent from the p150 residues shown. The orientation of all structures is the same (the addition of side chains changes the appearance of the structure greatly). Rendering and conservation mapping were performed with Protein Explorer (http:/molvis.sdsc.edu/protexpl/).

CLIP‐170 (Lin et al., 2001; Pierre et al., 1992) and disrupt the interactions of Alf1p/cofactor B with alpha‐tubulin (Feierbach et al., 1999). Mapping the sequence conservation between yeast Bik1p relatives onto the mammalian p150 crystal structure shows that this motif lies near a highly conserved central core region (Fig. 3C). Whether these conserved amino acids play a strictly structural role or possess other functions remain to be determined, but the conservation of surface residues suggests a role in protein– protein interactions. Mutations in at least one CAP‐Gly domain containing protein have been linked to human disease (Puls et al., 2003). Using human pedigrees, a mutation within the CAP‐Gly domain of the p150 dynactin subunit has been linked to an autosomal dominant form of a lower motor neuron disease. This is presumed to disrupt the essential function of dynactin in the dynein‐mediated retrograde transport of vesicles and organelles along microtubule tracks within


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axons. In the North American geneology studied, the disease presented with a relatively late age of onset, at about 35 years. This single amino acid substitution of glycine to serine at postion 59 (G59S) in p150 resulted in a reduced, but not abolished, ability of the protein to bind microtubules in vitro (Lansbergen et al., 2004; Puls et al., 2003). A stronger disruption of microtubule binding would probably have caused a more severe phenotype for the disease (Puls et al., 2003). This mutation also abolished the ability of p150 to bind to the C‐terminus of CLIP‐170 (Lansbergen et al., 2004). 2. Serine‐Rich Region Following the CAP‐Gly domain is a serine‐rich region of approximately 50 amino acids that is also found in other CLIP‐170–related proteins, including p150. This region is highly conserved at a primary sequence level within the Saccharomyces genus, but as organisms become more divergent, only the preponderance of serines and threonines remains obvious (Fig. 4). This lack of primary sequence conservation initially argues against a specific functional significance for this region. In CLIP‐170, perhaps it is just a poorly structured ‘‘molecular spring’’ connecting the two microtubule‐binding motifs. However, such a dismissal is likely premature. The serine‐rich region of CLIP‐115 is required for microtubule localization in vivo and microtubule binding in vitro (Hoogenraad et al., 2000), suggesting that this region, which is also often basic, is involved directly in microtubule binding. The large number of amino acids that can be phosphorylated hint at a possible regulatory role for this region. Addressing the function of this region will be an important focus for future eVorts.

B. Coiled‐Coil Domain Bik1p and all of its recognized orthologues contain an extended coiled‐coil domain which (at least in biochemically characterized proteins) allows the formation of parallel dimers (Hoogenraad et al., 2000; Lansbergen et al., 2004; Pierre et al., 1992; Scheel et al., 1999). In Bik1p and many of its fungal relatives, the coiled‐coil region is relatively short in comparison to that of CLIP‐170, consisting of only 207 amino acids located at residues 190–397 (Fig. 5). This is in contrast to the approximately 1000 amino acids commonly present in the mammalian proteins. The coiled‐coil domains are also binding interfaces for other proteins, including the CLIP‐associating proteins known as CLASPs, which are conserved members of the MAST/ ORBIT family (Galjart, 2005; Mimori‐Kiyosue et al., 2005). It is interesting to note that the coiled‐coil region in most CLIP‐170 relatives contains a break after approximately 110 amino acids (Fig. 5).

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Figure 4 Serine‐rich alignment. The alignment of serine‐rich regions of selected fungal Bik1p homologues is shown with human CLIP‐170 and p150 proteins for comparison. Sequence names are abbreviated as in Fig. 2. Alignment begins at the last conserved G of the CAP‐Gly motif and ends at the first amino acid before the coiled‐coil begins (predicted for each protein by the COILS server at http://www.ch.embnet.org/software/COILS_form.html). The alignment emphasizes the weak nature of primary sequence conservation in this region. The alignment was performed by ClustalW using default parameters.

C. Cargo‐Binding Domain At the C‐terminus of Bik1p and its relatives is a region sometimes called the ‘‘cargo‐binding domain’’ owing to an early conception that it bound vesicles (Pierre et al., 1992). Whether this region is involved in interactions with


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Figure 5 Coils plot. The prediction of the coiled‐coil structure in S. cerevisiae is shown for Bik1p, its fungal relatives, and human CLIP‐170. Analyses were performed by Coils server at http://www.ch.embnet.org/software/COILS_form.html using the 28 amino acid window. Plots were aligned at the beginning of the coiled coil. Some proteins have been truncated. Most sequences, including human CLIP‐170, have a significant predicted break approximately 110 amino acids after the beginning of the coiled coil. The conservation of this break is noteworthy given the otherwise poor conservation in this region. The significance of this break is not known, but it is interesting to speculate that it is related to intramolecular folding (Lansbergen et al., 2004) or other protein–protein interactions. The distance on the x‐axis is plotted in amino acids.

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membranes is unclear, but it is now known that this domain mediates a series of protein–protein interactions. This region contains two diVerent sequence motifs of note. First, there is a metal‐binding motif known as a ‘‘zinc knuckle,’’ and second, there is a conserved set of acidic and aromatic residues at the very C‐terminus, called the ‘‘Acidic‐Aromatic’’ motif. The zinc knuckle is a short motif consisting of critical histidine and cysteine residues in the sequence pattern C‐X2‐C‐X3‐GH‐X4‐C. In Bik1p, this is located at amino acids 416–429 (Fig. 6) (Pierre et al., 1992; Riehemann and Sorg, 1993). In other contexts, zinc knuckles have been shown to be involved in protein–nucleic acid interactions (Arning et al., 1996; Katz and Jentoft, 1989; Rein, 1994), but thus far the Bik1p and CLIP‐170 knuckles are known to bind only protein. Whereas Bik1p and its close relatives contain only one knuckle, the animal proteins and some fungal proteins contain two (Fig. 6). The single zinc knuckle motif in Bik1p appears to be most similar to the second of two zinc knuckles in the mammalian CLIP‐170 sequence (Goodson et al., 2003).

Figure 6 Alignment of metal‐binding motifs or ‘‘zinc knuckles.’’ Some proteins have two motifs, which are indicated by ‘‘K1’’ and ‘‘K2’’ after the sequence name. The gi numbers are given in Fig. 1. Human CLIP‐170 sequences are provided for comparison.


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As discussed in more detail later, the first zinc knuckle of CLIP‐170 is involved in the head to tail interaction of CLIP‐170 with itself (Goodson et al., 2003; Lansbergen et al., 2004). The second zinc knuckle mediates interactions with other proteins including p150 and LIS1 (Coquelle et al., 2002; Goodson et al., 2003; Lansbergen et al., 2004; Tai et al., 2002). There may be some competition between the two knuckles for binding to various ligands, as the binding sites on CLIP‐170 for the N‐terminus of CLIP‐170 and p150 may overlap (Lansbergen et al., 2004). It is not clear whether the single zinc knuckle of the Bik1p and its close relatives accomplishes the same job as the two motifs of the animal proteins, or whether the extra motif in the animal CLIP‐170 relatives mediates additional functions. The second sequence motif of note is located at the very C‐terminus. It is characterized by an aromatic amino acid, often preceded by a series of acidic residues. This sequence, which we term here the ‘‘Acidic‐Aromatic’’ motif, has not been previously noted in published work and is not obvious except when examining an alignment of several Bik1p/CLIP‐170 proteins (Fig. 7). The function of this motif is not known, but is nearly universally conserved in the Bik1p/CLIP‐170 family. This motif is strikingly similar to the one present at the C‐terminus of proteins in the EB1 family and the C‐terminus of alpha‐ tubulin (Badin‐Larcon et al., 2004; Hayashi et al., 2005; Komarova et al., 2005). In S. cerevisiae, the absence of the C‐terminal hydrophobic phenylalanine in the alpha‐tubulin Tub1p results in a decrease of Bik1p‐GFP localization at the plus end of cytoplasmic microtubules (Badin‐Larcon et al., 2004).

Figure 7 A comparison of the C‐terminal Acidic‐Aromatic motif in fungal and metazoan CLIP‐170 members. Organisms are abbreviated by the first four letters of the genus and two letters of the species. Proteins are given published names or the gi numbers. Sequences start with the last conserved cysteine of the zinc kunckle and end with the final amino acids of the proteins.

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We predict that this motif in Bik1p and CLIP‐170 will also have significant functions.

III. Bik1p Interactions with Other Proteins Like most proteins, Bik1p does not function in isolation within the cell, interacting instead with a series of other proteins. The function of these interactions—the when, where, why, and how—are only now beginning to be understood. Several proteins have been identified as interacting with Bik1p by two‐hybrid analysis and/or physical methods. These include proteins that function in positioning the mitotic spindle such as the EB1 homologue Bim1p (Ito et al., 2001; Moore and Miller, unpublished two hybrid observations); the MAP215 homologue Stu2p (Chen et al., 1998; Ito et al., 2001; Kosco et al., 2001; Lin et al., 2001; van Breugel et al., 2003); the kinesin Kip2p (Carvalho et al., 2004); the actin to microtubule linking protein Kar9p (Moore et al., 2006); and the LIS1 homologue Pac1p (Sheeman et al., 2003). Bik1p also interacts with the ‘‘target of rapamycin’’ kinases TOR1 and ‐2 (Choi et al., 2000). Lap4p and Spp382p were also identified in genome‐wide screens for Bik1p interacting proteins, but the basis for these connections remains obscure (Hazbun et al., 2003; Ho et al., 2002). The function(s) of Bik1p’s interaction with many of these proteins has been particularly diYcult to dissect out because several (Bim1p, Stu2p, and Kip2p) are themselves microtubule‐binding proteins (Korinek et al., 2000; Lee et al., 2000; Wang and HuVaker, 1997). In addition, several of these proteins display multiple interactions with each other (Wolyniak et al., 2006). For instance, Bim1p binds to Kar9p and is required for the localization of Kar9p to microtubules. This is somewhat analogous to the binding of mammalian EB1 to the adenamatous polyposis coli (APC) protein (Mimori‐Kiyosue et al., 2000; Wen et al., 2004), which is mutated in approximately 80% of human colon cancers (Kinzler and Vogelstein, 1996; Sieber et al., 2002). Bim1p also binds to Stu2p (Chen et al., 1998). However, inferences drawn from other systems have shed some light on various possible functions for some of these interactions. In fission yeast, the EB1 homologue is involved in plus‐end localization of the CLIP‐170 homologue (Busch and Brunner, 2004). EB1 binds the CAP‐Gly domain found in CLIP‐190 of Drosophila (Dzhindzhev et al., 2005) and p150Glued of mammalian cells (Askham et al., 2002; Hayashi et al., 2005; Ligon et al., 2003). Therefore, it is possible that Bim1p also binds the CAP‐Gly domain of Bik1p in S. cerevisiae. Indeed recently, two‐hybrid analysis has shown that Bim1p interacts with a segment of Bik1p containing both the CAP‐Gly domain and serine‐rich region (Wolyniak et al., 2006). Considering that Bik1p promotes microtubule stability and that Bim1p promotes the dynamicity of microtubules, enhancing their ability to


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switch between growing and shrinking phases (Tirnauer et al., 1999), it will not be surprising to learn that they modulate or even enhance each other’s activity in some way. Bik1p also interacts with the alpha‐tubulin, Tub1p, by two‐hybrid analysis (Schwartz et al., 1997). In a two‐hybrid analysis of a battery of TUB1 alanine‐scanning mutations, BIK1 interacts with several alleles that fail to interact with both BIM1 and ALF1 (Feierbach et al., 1999; Richards et al., 2000; Schwartz et al., 1997). The disruption profiles of BIM1 and ALF1 display several similarities, indicating that Bim1p and Alf1p may share an overlapping binding site (Feierbach et al., 1999). In contrast, the disruption profile of BIK1 is quite diVerent, suggesting that its binding site may have limited (if any) overlap with that of ALF1. However, further work will be required to determine whether Bik1p has a unique binding site on the alpha‐tubulin molecule. Information about possible Bik1p interactions with beta‐ and gamma‐tubulin is incomplete. Consistent with the presence of a long central coiled‐coil motif, several lines of evidence including EM analysis, sedimentation, and scanning force microscopy suggest that CLIP‐170 forms long parallel dimers (Lansbergen et al., 2004; Pierre et al., 1992; Scheel et al., 1999). Similarly, Bik1p interacts with itself through the coiled‐coil region by two‐hybrid analysis (Wolyniak et al., 2006; D’Silva and Miller, unpublished observations). In addition to long rods, scanning force microscopy experiments and fluorescence resonance energy transfer (FRET) experiments suggest a model in which CLIP‐ 170 is a flexible molecule that permits an autoinhibitory intramolecular interaction between the N‐ and C‐terminal domains that reduces its aYnity for microtubules (Lansbergen et al., 2004). Phosphorylation is postulated to release the inhibition, allowing it to bind to microtubules (Lansbergen et al., 2004). This autoinhibition seems to be mediated by interaction of the CAP‐Gly motifs with the first metal‐binding motif (Lansbergen et al., 2004). The absence of this motif in Bik1p suggests that similar autoinhibition may not occur in Bik1p, but it is also possible that the single zinc knuckle in Bik1p mediates many or all of the interactions partitioned between the two CLIP‐170 knuckles.

IV. Phenotypes and Activities Bik1p is a microtubule‐stabilizing factor, and yeast cells lacking Bik1p have very short microtubules (Berlin et al., 1990). Consistent with these characteristics, Bik1p localizes with the plus ends of microtubules. It is also found at the spindle pole body (SPB), which is the microtubule‐organizing center in yeast (Fig. 8). Several mechanisms for this microtubule stabilization are possible. Bik1p might act to either inhibit catastrophes or promote rescue events. CLIP‐170 has been shown to promote microtubule rescue in mammalian

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Figure 8 Bik1p localizes to SPBs and the plus end of microtubules. Bik1p was localized in living S. cerevisiae cells using Bik1p fused to 3X‐GFP (a gift from Dr. David Pellman). Microtubules were visualized with CFP‐Tub1p.

systems (Arnal et al., 2004; Komarova et al., 2002). In S. pombe, Tip1p is thought to act as an anticatastrophe factor (Brunner and Nurse, 2000). The microtubule‐stabilizing activity of Bik1p is likely to be mediated through its CAP‐Gly domain. This prediction is based on the observation that the conserved CAP‐Gly domain is necessary for microtubule‐binding activity in CLIP‐170, CLIP‐115, and p150Glued (Hoogenraad et al., 2000; Pierre et al., 1992; Waterman‐Storer et al., 1995). This requirement has not been shown directly for Bik1p, but several pieces of data are consistent with this model. Deletion of the CAP‐Gly domain or mutagenesis of conserved residues within it (N43A, G45A, K46E, and G49A found in Bik1p‐MTP mutant) abolish the localization of the Bik1p protein with microtubules (Lin et al., 2001). Several questions remain concerning the microtubule‐binding activity of Bik1p. As mentioned earlier, Bik1p has only one CAP‐Gly domain, whereas CLIP‐170 has two; raising the issue of the relative contributions of multiple CAP‐Gly domains to microtubule binding. Direct microtubule‐binding studies using CLIP‐170 fragments did not test whether a single CAP‐Gly domain was suYcient for microtubule binding (Pierre et al., 1992), although one binding unit is suYcient for p150Glued (Waterman‐Storer et al., 1995). Neither of the two CAP‐Gly domains in CLIP‐115 bind to microtubules with high aYnity in vitro, instead requiring contributions from the serine‐ rich region (Hoogenraad et al., 2000). Future investigations will show whether the serine‐rich region of Bik1p is similarly required. Determining whether Bik1p actually binds to tubulin dimers is an area ripe for investigation. Studies in this area should shed light on whether


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treadmilling mechanisms contribute to Bik1p behavior in yeast (see below). Unfortunately, Bik1p‐his6 purified from bacteria has not proven to be functional in in vitro microtubule‐binding assays, so more involved approaches will be required (Miller, D’Silva, Folker, and Goodson, unpublished observations).

V. Role of Bik1p in Spindle Positioning Bik1p associates with both the intranuclear and extranuclear (or cytoplasmic) microtubules of yeast. Its role in positioning the mitotic spindle is derived from its eVects on cytoplasmic microtubules.

A. Genetics of Spindle Positioning Two major systems termed the Kar9p pathway and the dynein pathway control spindle positioning in yeast (Miller and Rose, 1998). These two systems are partially redundant with each other, meaning that either one alone is suYcient for overall cell viability. Mutations in just one pathway result in only very mild growth defects (Li et al., 1993; Miller and Rose, 1998; Yeh et al., 1995). However, when both pathways are defective, the cell is extremely sick or dead, a condition called ‘‘synthetic lethality’’ (Miller and Rose, 1998; Muhua et al., 1994). The Kar9p pathway functions in preanaphase to move the spindle up to the bud neck and align it along the mother‐bud axis (Kusch et al., 2002; Miller and Rose, 1998; Miller et al., 2000). The spindle in kar9D mutants is positioned farther away from the bud neck at the beginning of anaphase than in wild type (Yeh et al., 2000). The dynein pathway is so named for its key player, the minus end‐directed microtubule motor, dynein. Dynein is responsible for moving the spindle across the bud neck at or just before anaphase (Bloom, 2001; Kahana et al., 1995; Yeh et al., 1995). In dynein mutants, the spindle instead frequently elongates entirely within the mother cell, leaving the bud temporarily without a nucleus. The spindle continues to elongate by a ‘‘telescoping’’ mechanism that extends the spindle through the bud neck, continuing until the nuclear masses are separated at the distal ends of the mother and bud (Yeh et al., 1995). Finally, cytokinesis ensues and the cells enter G1. For both pathways, the spindle movements are controlled by the interaction of cytoplasmic microtubules with the bud cortex and bud neck, in combination with microtubule dynamics and microtubule motor activities (Gundersen and Bretscher, 2003; Kusch et al., 2002; Pearson and Bloom, 2004; reviewed in Bloom, 2001).

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The Kar9p pathway is sometimes referred to as the ‘‘early pathway’’ and the dynein pathway referred to as the ‘‘late pathway.’’ However, the exact nature of the temporal overlap between the two pathways has not been carefully defined to date. While the Kar9p and dynein pathways play distinctive roles in spindle positioning, it is likely be an oversimplification of the actual cell biology to just say they are the ‘‘early’’ and ‘‘late’’ pathways. Live cell analysis of dynein mutants showed that the ‘‘bouncing around’’ or ‘‘jostling’’ of the spindle that normally occurs in a wild‐type mother cell prior to anaphase is greatly diminished in the dynein mutant (Yeh et al., 1995, 2000). This suggests that dynein is active prior to pulling the spindle across the bud neck. How late in the cell cycle Kar9p acts also remains to be determined. The relatively mild growth defect seen for mutations in either the Kar9p or dynein pathways is the result of a checkpoint mechanism that monitors the position of the spindle. This delays the cell cycle until the spindle can be repositioned across the neck. Thus, a cell is prevented from undergoing cytokinesis when two nuclei reside within the mother cell and the daughter has none (Bloecher et al., 2000; Daum et al., 2000; Yeh et al., 1995). The spindle‐positioning checkpoint involves many of the genes important for regulating exit from mitosis, also known as the mitotic exit network (MEN). These include BUB2, BFA1, TEM1, LTE1, and KIN4 (Bardin et al., 2000; Bloecher et al., 2000; Pereira and Schiebel, 2005; reviewed in D’Amours and Amon, 2004; Pereira et al., 2000, 2001). With many important details of this signaling network emerging rapidly, it is abundantly clear that the cell has good sensors to carefully monitor the position of the SPBs relative to the plane of cytokinesis, located at the bud neck in yeast.

B. Bik1p in the Dynein Pathway In yeast, dynein most likely exerts its force from the cortex, where it powers microtubule sliding events that pull the nucleus across the bud neck (Adames and Cooper, 2000) (For a review of dynein function at the cortex in mammalian systems see Dujardin and Vallee, 2002). The predominant model describing how dynein gets to the cortex is the ‘‘oV‐loading’’ model (Lee et al., 2005). In this model, dynein is transported to the cortex by riding out on the dynamic plus end of cytoplasmic microtubules. It is then transferred or ‘‘oV‐loaded’’ to the cortex where it becomes activated to pull on the cytoplasmic microtubules. Thus, the mechanisms that target dynein to the plus end are critically important for regulating dynein function. Genetic analysis first suggested that Bik1p was likely to play a role in the dynein pathway (Miller and Rose, 1998). bik1D mutants are synthetically lethal with a number of mutants in the Kar9p pathway, but not with mutations in dynein or the dynactin complex. Thus, Bik1p has an essential


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function in the dynein pathway (Miller and Rose, 1998). The role of Bik1p in dynein function is most likely the basis for its identification as a pac mutant (perish in the absence of cin8, pac14). Cin8p is a kinesin with a role in the mitotic spindle that is also synthetically lethal with mutations in dynein (Geiser et al., 1997). The molecular basis for these genetic observations was elucidated by an elegant series of cell biological studies that showed that Bik1p helps recruit dynein to the plus end of the microtubule (Lee et al., 2003, 2005; Li et al., 2005; Sheeman et al., 2003). Bik1p is actually one of at least four proteins (Bik1p, Kip2p, Pac1p/LIS1, and Ndl1p) whose function in the dynein pathway is based in part on their ability to recruit dynein to the plus end of the microtubule (Li et al., 2005; Sheeman et al., 2003). Ndl1p is required for Pac1p localization to microtubules but is not required for the localization of Bik1p (Li et al., 2005). Based on this hierarchy of localizations, these proteins appear to constitute a two‐part system for the recruitment of dynein, Ndl1p/Pac1p comprising one component and Bik1p the other (Li et al., 2005). Experiments defining how these two modules interact with other elements of the dynactin complex will continue to bring new excitement to the dynein field (Efimov, 2003; Efimov and Morris, 2000). In mammals, two Ndl1p/nudE homologues, Ndel1 and Nde1, have been identified. Ndel1 has been implicated in the recruitment of the p60 subunit of microtubule severing katanin (Toyo‐oka et al., 2005), but a similar activity has not been identified in yeast. In both CLIP‐170 and Bik1p, the C‐terminal cargo‐binding domain is involved in binding LIS1 and its yeast orthologue, Pac1p, respectively. Mutations in the human LIS1 gene are causal for the severe developmental brain disease, type 1 lissencephaly, which results from incomplete neuronal migration (Kato and Dobyns, 2003; Sapir et al., 1999; reviewed in Tsai et al., 2005; Tsai and Gleeson, 2005; Vallee et al., 2001). LIS1 itself interacts with three diVerent subunits of the dynein and dynactin complex (Coquelle et al., 2002; Tai et al., 2002; Tsai et al., 2005), an idea that was originally established in the filamentous fungi, Aspergillus nidulans (Morris et al., 1998; Xiang et al., 1995; see Xiang, 2003 for a review). In S. cerevisiae, the last 134 amino acids of Bik1p are necessary and suYcient for its two‐hybrid interaction with Pac1p (Geiser et al., 1997; Sheeman et al., 2003). In pac1D and bik1 mutants lacking the cargo‐binding domain, Dyn1p‐3GFP localization is largely lost at the microtubule plus end (Lee et al., 2003; Sheeman et al., 2003). In mammalian systems, the C‐terminal domain of CLIP‐170 also interacts directly with p150Glued (Lansbergen, et al., 2004). This interaction in vitro requires that the second zinc knuckle of CLIP‐170 be intact (Coquelle et al., 2002; Goodson et al., 2003; Lansbergen et al., 2004; Valetti et al., 1999). In vitro binding studies suggest that LIS1 and p150Glued compete for binding at this site (Lansbergen et al., 2004). Considering the significant conservation of function between mammalian and fungal forms of LIS1, it

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will be interesting—but perhaps not unexpected—if future experimentation shows that the single zinc knuckle motif in Bik1p is also required for its binding to Pac1p.

C. Bik1p and the Kar9p Pathway A defining feature of the Kar9p pathway is that it orients cytoplasmic microtubules by linking the actin and microtubule cytoskeletons. This linkage occurs at the distal ends of cytoplasmic microtubules, and consists of a Kar9p bridge between the type V myosin Myo2p and the microtubule‐binding protein, Bim1p. With this linkage established, Myo2p walks along actin cables steering Kar9p‐attached microtubules from the mother cell toward the bud neck and into the bud (Beach et al., 2000; Hwang et al., 2003; Yin et al., 2000) (Fig. 9). Accordingly, mutations in either Bim1p or Kar9p result in misoriented microtubules (Gundersen et al., 2004; Huisman and Segal, 2005; Korinek et al., 2000;

Figure 9 Model for Bik1p in spindle positioning during mitosis. Bik1p is delivered to the plus end of the microtubule by the kinesin Kip2p. In the presence of Bik1p, Kar9p is localized to the daughter‐bound SPB. But in the absence of Bik1p, Kar9p is present on both the mother‐ and daugher‐bound SPBs.


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Lee et al., 2000; Miller et al. 2000; reviewed in Gundersen and Bretscher, 2003). Current models depict Kar9p as interacting with only one microtubule‐binding protein, Bim1p, at the plus end. Cytoplasmic microtubules emanate from both the original (or old) SPB and the newly duplicated (or new) SPB. Of the two SPBs, the old SPB is the one inherited by the daughter cell. The new SPB is retained in the mother cell (Pereira et al., 2001). A key element in the myosin‐guidance model is that Kar9p is associated with only the old SPB and the microtubules emanating from it. This ensures that only this set of microtubules has the capacity to be carried into the bud by myosin. Therefore, the mechanisms that direct Kar9p to the old SPB are vital for high fidelity inheritance of this SPB. Recently, Kar9p was shown to physically interact with Bik1p (Moore et al., 2006). The role that Bik1p plays with Kar9p for spindle positioning appears to be diVerent than that played by Bim1p (Moore et al., 2006). Bim1p is essential for recruiting Kar9p to the old SPB and microtubules (Liakopoulos et al., 2003; Miller and Rose, 1998). In contrast, Bik1p is important for restricting Kar9p to the old, daughter‐bound SPB. In cells deleted for Bik1p, Kar9p localization is found on both SPBs rather than just the SPB destined for the bud (Fig. 10) (Moore et al., 2006). As a result, bik1D mutants have an increased number of cells in which the new SPB is incorrectly inherited by the daughter cell (Moore et al., 2006). The mechanism by which Bik1p restricts Kar9p to one SPB involves the phosphorylation of Kar9p (Moore et al., 2006). Two sites of phosphorylation have been identified on Kar9p, serines 197 and 496 (Liakopoulos et al., 2003), although a third is likely to exist (Moore et al., 2006). Consistent with this, at least three isoforms of Kar9p can be observed in whole cell extracts, resulting from phosphorylation at either one or both sites. In bik1D extracts, one of these isoforms is greatly diminished, suggesting that Bik1p promotes one of the phosphorylation events on Kar9p (Fig. 11) (Moore et al., 2006). Moore et al. (2006) postulate that Bik1p contributes to the phosphorylation at serine 496 and that this is likely to involve the B‐type cyclin Clb5p, which acts in conjunction with the cyclin‐dependent kinase, Cdc28p (Moore et al., 2006). Clb5p interacts with Kar9p in a region that overlaps serine 496 (Moore et al., 2006). This is a segment that is conserved between the APC protein and Kar9p (Bienz, 2001; Liakopoulos et al., 2003). In support of this hypothesis, Bik1p is required for the two‐hybrid interaction of Clb5p with Kar9p. One model explaining these results is that Bik1p binds to Kar9p and induces a conformational change. This allows the Clb5p/Cdc28p kinase access to Kar9p for the phosphorylation event (Moore et al., 2006). This model predicts that pseudo‐phosphorylation at serine 496 should suppress the mislocalization of Kar9p in the bik1D mutant. However, this model does not define the identity of the actual asymmetrically‐localized selector protein that restricts Kar9p to one SPB.

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Figure 10 Bik1p acts in the restriction of Kar9p to one SPB. In wild‐type cells with short preanaphase spindles, SPBs were visualized by labeling the SPB component Spc110p with CFP. Kar9p‐3GFP localizes to one of the two poles and can be seen on cytoplasmic microtubules emanating from that pole. In bik1D cells, Kar9p‐3GFP is commonly detected at both poles and both sets of cytoplasmic microtubules.

The mechanism by which Bik1p alters Kar9p localization is novel. The possibility that Bik1p alters Kar9p localization by altering its phosphorylation status (rather than through a simple stoichiometric binding mechanism) is quite remarkable. Prior to this report, MAPs were not known to play a role in altering the phosphorylation of other MAPs. Furthermore, the identification of Bik1p as a protein common to both the Kar9p and dynein pathways suggests that mechanisms may be found in the future for coordinating these two pathways through this common element.


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Figure 11 Bik1p promotes the phosphorylation of Kar9p. Protein extracts from wild‐type and bik1D cells expressing TAP‐tagged Kar9p were treated with lambda phosphatase or mock treated with no enzyme (Moore et al., 2006). A Western blot of these extracts was probed with HA antibody to detect the HA epitope of the TAP tag. In extracts from wild‐type cells, a triplet of Kar9p‐tap bands is apparent. Treatment with phosphatase collapses this triplet to primarily a single major band, indicating that the two slower‐migrating bands represent phosphorylated species of Kar9p‐tap. In extracts from bik1D cells, only a doublet of Kar9‐tap is visible, suggesting that at least one phospho‐isoform of Kar9p‐tap is absent or greatly diminished in bik1D.

Because bik1D mutants have short cytoplasmic microtubules, one might initially wonder whether they are long enough reach the bud cortex and make a contribution to spindle positioning. One explanation is that the microtubules may really only need to be long enough to reach an actin cable where the Myo2p‐Kar9p‐Bim1p bridge connects the actin and microtubule cytoskeletal networks. Since actin cables extend from the bud neck into the mother cell, this distance may not actually be that great.

D. Bik1p in Mating Cells Like mitotic cells, Bik1p plays an important role in mating cells, where it acts to maintain microtubule length (Berlin et al., 1990; Kurihara et al., 1994). Studies suggest that Bik1p also makes additional contributions to the mating process (Molk et al., 2006). In mating cells (also called shmoos), cytoplasmic microtubules extend from the SPB to the tip of the mating projection. Here they maintain persistent yet dynamic attachments, undergoing alternating phases of both growth and shortening, all the while remaining in contact with the cell surface (Maddox et al., 1999). Live cell analyses suggest that Bik1p plays an important role in this cortical attachment. When the microtubule enters the depolymerization phase in bik1D shmoos, it detaches from the cortex rather than staying in contact with the cortex as it does in wild type (Molk et al., 2006). These experiments show that Bik1p is necessary to maintain persistent microtubule contacts with the cortex prior to cell fusion (Molk et al., 2006). A similar role was shown previously for the Kar3p kinesin in linking the plus end to the shmoo cortex (Maddox et al., 2003).

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Both microtubule depolymerization and polymerization occur at the plus end in yeast (Maddox et al., 1999, 2000). In the shmoo, these alternating cycles result in corresponding oscillations of the attached SPB toward and away from the shmoo tip (Maddox et al., 1999). During microtubule depolymerization, the fluorescence intensity representing Bik1p‐GFP localization at the microtubule tip increases. Conversely during the polymerization phase, Bik1p‐GFP localization decreases (Molk et al., 2006). This localization profile is similar to that seen for the kinesin Kar3p‐GFP, but diVerent than that seen with Bim1p‐GFP (Maddox et al., 2003; Molk et al., 2006). In addition to its minus end‐directed motor activity, Kar3p has been shown to display depolymerizing activity at both the microtubule minus end (Endow et al., 1994) and at the plus end (Sproul et al., 2005). Therefore, the increased localization of Kar3p‐GFP at the microtubule plus end is postulated to tether the depolymerizing microtubule to the cortex (Maddox et al., 2003). In contrast, Bik1p acts in microtubule stabilization. Therefore, its increased localization at the plus end of the depolymerizing microtubule seems somewhat paradoxical because it is being concentrated at a site during a time when its microtubule stabilizing activity should be minimal. One model that reconciles these observations suggests that Bik1p together with Kar3p is part of a sliding microtubule collar or ‘‘sleeve’’ that links microtubule dynamics to the cortex (Molk et al., 2006). Thus, this model emphasizes the microtubule binding activity of Bik1p rather than its microtubule stabilizing activity. After cell fusion in mating, the two haploid nuclei congress to the center of the zygote for nuclear fusion. Early models attributed the generation of the motive force to overlapping microtubules of opposite orientation and the minus end‐directed motor activity of the Kar3p kinesin (Meluh and Rose, 1990). Live cell imaging of nuclear congression suggests a new model in which little or no overlap is present between oppositely oriented microtubules (Molk et al., 2006). Bik1p‐GFP marks the plus end of the oppositely oriented microtubules and during congression forms one dot rather than two, suggesting that Bik1 helps maintain plus end linkages during nuclear congression (Molk et al., 2006). These analyses raise intriguing new questions and provide important new insights into the mechanism by which microtubule depolymerization at the plus end can be coupled to work.

VI. Models for Plus‐End Binding A. Microtubule Plus‐End Tracking (þTIP) Behavior Because Bik1p and CLIP‐170 were two of the first plus‐end tracking proteins identified, they serve as the premier models for how plus‐end tracking proteins are maintained at the dynamic plus end of the microtubule. The reader is


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referred to several excellent in‐depth reviews for additional reading on this topic (Carvalho et al., 2003; Schuyler and Pellman, 2001). In mammalian cells, CLIP‐170 visualized with a GFP fusion is seen in a comet‐like pattern following the growing, but not the shrinking, ends of cytoplasmic microtubules (Perez et al., 1999). These CLIP‐170 comets move from the centrosomal area of the cell out toward the cell periphery. Movies of plus‐end tracking in live cells really are one of the seven wonders of the biology world. Readers are encouraged to view these movies in the supplemental data of any of a variety of manuscripts including ones for CLIP‐170 (Folker et al., 2005; Kandere‐ Grzybowska et al., 2005; Perez et al., 1999); dynein in A. nidulans (Zhang et al., 2003); p150Glued in tissue culture (Vaughan et al., 2002); EB1 and EB3 in tissue culture cells and neurons (Mimori‐Kiyosue et al., 2000; Stepanova et al., 2003); CLASP (Wittmann and Waterman‐Storer, 2005); and Tea1p in fission yeast (Feierbach et al., 2004). While both CLIP‐170 and Bik1p are plus‐end tracking proteins, there is a key diVerence that enhances the interest in these two proteins. CLIP‐170 is seen only on the growing end of microtubules, whereas Bik1p is also found on the shrinking ends, a phenomenon referred to as ‘‘backtracking’’ (Lin et al., 2001). The basis for this diVerence is not currently understood, but it suggests, along with the experiments described later, that the mechanisms of how Bik1p and CLIP‐170 associate with the plus end may be fundamentally diVerent.

B. Mechanisms of Microtubule Plus‐End Tracking Behavior Three mechanisms have been proposed for how þTIPs dynamically track microtubule plus ends. These models, which are not mutually exclusive, can be described as ‘‘motor‐based transport,’’ ‘‘end‐loading,’’ and ‘‘surfing’’ (Fig. 12). In a ‘‘motor‐based transport’’ mechanism, the þTIP moves to the plus end by riding on a plus‐end directed molecular motor (Fig. 12A). In such a mechanism, the motor requires a means of remaining attached to the microtubule once the end is reached so that the þTIP can become concentrated at the plus end. In an ‘‘end‐loading’’ mechanism (also called ‘‘treadmilling’’), there are actually two separable phases—binding specifically at the tip, followed by dissociation from ‘‘older’’ areas of polymer. Tip binding could occur because the þTIP has higher aYnity for some end‐specific conformation (such as the GTP cap), or because the þTIP binds to unpolymerized tubulin and copolymerizes with it. Dissociation from ‘‘older’’ polymer could occur by a variety of mechanisms, including intrinsically weak aYnity for polymer or regulation. ‘‘Surfing’’ is a term often applied to any example of þTIP behavior, but we use it here to refer specifically to a mechanism in which a þTIP diVuses along the microtubule lattice in the anterograde direction, concentrating at the end because it has higher aYnity

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Figure 12 Potential mechanisms of microtubule plus‐end tracking. (A) Motor‐driven transport—CLIP‐170 (red) is physically moved toward the microtubule tip by motors (teal). Fungal CLIP‐170 homologues Bik1p and Tip1p appear to track by this mechanism. (B) Surfing— CLIP‐170 (red bars) diVuses (‘‘surfs’’) along a tip‐specific conformation or another plus‐end tracking protein (bright green). Note that CLIP‐170 is represented by bars rather than circles because this model likely requires multiple binding sites. (C) End‐loading—CLIP‐170 (red) binds specifically at the microtubule plus end then dissociates shortly thereafter. CLIP‐170 is stationary on the microtubule and would not be observed to move toward either the plus or minus end of the microtubule. Loading at the tip could occur by preferential binding of CLIP‐170 to the cap (‘‘cap‐recognition’’), or by preassociation with unpolymerized tubulin followed by copolymerization. Dissociation could occur by CLIP‐170 having an intrinsically weak aYnity for the ‘‘older’’ GDP polymer, or be induced by a regulatory event that reduces the aYnity of CLIP‐170 for microtubules. Figure and legend reprinted from Folker et al., 2005 with permission.


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for an end‐specific conformation. We call this surfing because the þTIP rides on the potential energy ‘‘wave’’ created by the constantly shifting position of the end‐specific conformation. Of course, proteins could also tip‐track by binding to other proteins that are tip‐tracking. This mechanism is sometimes called ‘‘hitchhiking.’’ As indicated in Fig. 12, one way to experimentally distinguish between these mechanisms is to determine whether the þTIP molecules move relative to the microtubule lattice: in the ‘‘motor‐driven’’ and ‘‘surfing’’ mechanisms, the þTIP molecules do move, but in treadmilling mechanisms, the movement is an illusion—the repeated cycles of binding at the plus end and releasing from older areas of the polymer create a shifting gradient of þTIP concentration that is similar to what would happen if one could visualize the GTP cap. Data suggest that Bik1p utilizes primarily a motor‐based transport mechanism, whereas CLIP‐170 uses mostly a copolymerization‐based treadmilling mechanism (Carvalho et al., 2004; Folker et al., 2005; Perez et al., 1999). The evidence that Bik1p utilizes a motor‐based transport mechanism comes from several observations. First, the localization of Bik1p to plus ends is highly dependent on the kinesin motor, Kip2p; in a strain lacking Kip2p, the amount of Bik1p at the plus end of the microtubule is decreased dramatically (Carvalho et al., 2004). Concomitantly, the amount of Bik1p present at the SPB is increased. Overexpression of Kip2p drives additional Bik1p to the plus end (Carvalho et al., 2004). In further support of this model, dual channel microscopy demonstrates that Bik1p‐3CFP and Kip2‐ 3YFP comigrate along the cytoplasmic microtubule toward the plus end (Carvalho et al., 2004). Moreover, Bik1p and Kip2p physically interact in a complex, as shown by immunoprecipitation (Carvalho et al., 2004). The S. pombe, CLIP‐170 homologue Tip1p and kinesin homologue Tea2p interact in similar ways (Busch et al., 2004), suggesting that this mechanism may be conserved, at least among fungi. This evidence indicates that Kip2p plays an important role in controlling the amount of Bik1p at the plus end. However, the fact that residual amounts of Bik1p are found at the plus end of the microtubule in kip2D strains (Carvalho et al., 2004) suggests that there are also Kip2p‐independent mechanisms for localizing Bik1p to the plus end. It is also interesting to note that when yeast expressing a temperature sensitive allele of STU2 (a MAP215 homologue that binds to Bik1p) are shifted to the nonpermissive temperature, Bik1p is lost from the SPB, but not from the plus end. This observation supports the idea that Bik1p can be recruited to the plus end from the cytoplasm, and not only loaded at the SPB (Carvalho et al., 2004). In contrast, CLIP‐170 appears to track plus ends primarily by a copolymerization‐based treadmilling mechanism. This conclusion is based on two types of data. First, movies and kymographs of CLIP‐170 comets indicate that the pattern of CLIP‐170 distribution on the microtubule does

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not change significantly with time, suggesting that CLIP‐170 molecules do not move relative to the microtubule lattice (Folker et al., 2005; Perez et al., 1999). This lack of movement excludes motor‐based transport and surfing, leaving treadmilling as the likely mechanism. Second, biochemical analysis shows that CLIP‐170 binds tightly and stably to unpolymerized tubulin (Folker et al., 2005) and promotes formation of tubulin oligomers (Arnal et al., 2004). This strong association with tubulin indicates that CLIP‐170 should be saturated by tubulin when the proteins are at physiological concentrations and suggests that CLIP‐170 tracks microtubule plus ends by a copolymerization mechanism (Folker et al., 2005). Why would CLIP‐170 and Bik1p, which seem otherwise so similar, track plus ends by fundamentally diVerent mechanisms? Two answers seem apparent. First, the plus‐end tracking mechanisms of Bik1p and CLIP‐170 may not be as diVerent as initially thought. Given the array of protein–protein interactions mediated by þTIPs, it seems likely that many þTIPs can get to microtubule plus ends by more than one mechanism. One could imagine a situation where the set of interactions mediated by orthologous þTIPs is conserved, but the predominant interaction diVers from one system to another. An analogy can be found in the relative functions of the actin and tubulin cytoskeletons in yeast and animal cells. This consideration suggests that CLIP‐170 will be found to interact with members of the kinesin family. Second, it should be noted that the properties of yeast tubulin and animal tubulin diVer significantly. It is possible that these diVerent properties (slower GTPase, slower growth) confer diVerences in the mechanisms used by þTIPs to track microtubule plus ends (Folker et al., 2005). C. Interactions Between EB1 and CLIP‐170 Versus Bim1p and Bik1p EB1 proteins are also a very highly conserved family of microtubule plus‐end binding proteins. Thus, the role that their interactions play with CLIP‐170 has aroused keen interest. Three EB family members have been identified in mammals, EB1, EB2, and EB3 (Juwana et al., 1999; Mimori‐Kiyosue et al., 2000, 2005; Morrison et al., 1998; Nakagawa et al., 2000; Stepanova et al., 2003; Su et al., 1995). EB1 proteins contain a calponin homology domain which is responsible for its microtubule‐binding activity (Hayashi and Ikura, 2003), and a coiled‐coil domain which is responsible for dimerization (Honnappa et al., 2005; Slep et al., 2005). While studies concerning the interactions between CLIP‐170 family members and EB1 proteins are ongoing in several laboratories, reports suggest that intriguing diVerences exist between EB family members (Komarova et al., 2005). CLIP‐170 binds all three EB proteins through its N‐terminal domain, although its aYnity for EB1 and EB3 is higher than it is for EB2


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(Komarova et al., 2005). Experiments depleting both EB1 and EB3 expression through RNAi knockdown resulted in a quicker release of CLIP‐170 from the microtubule ends. EB2 did not appear to have a strong eVect (Komarova et al., 2005). In the converse experiment, overexpression of EB1 resulted in a slower rate of CLIP‐170 release from microtubule ends. In vitro binding studies also suggest that the interaction of CLIP‐170 with EB1 is influenced by the presence of the C‐terminal tail of EB1 (Komarova et al., 2005). In budding yeast, preliminary studies show that Bim1p is not required for the localization of Bik1p at the plus end (Carvalho et al., 2004). In addition, strains lacking BIM1 may have an increased amount of Bik1p at the plus ends (Carvalho et al., 2004). From these studies, it appears that EB1 proteins regulate the release of CLIP‐170 proteins, but not necessarily their assembly onto microtubule ends. Additional details of this regulation remain to be elucidated in other systems and exciting experiments should be expected in the future.

VII. Regulation of CLIP‐170 and Bik1p by Phosphorylation Relatively little is known about the regulation of CLIP‐170 and Bik1p. The association of CLIP‐170 to microtubules is regulated by phosphorylation. First, phosphorylation inhibits its binding to microtubules (Rickard and Kreis, 1991). The kinase, which remains unidentified, acts primarily on serine residues and appears to copurify with microtubules. Depolymerization of microtubules with nocodazole induces a rapid dephosphorylation of CLIP‐170 (Rickard and Kreis, 1991). Phosphorylation by protein kinase A (PKA) of p150 near the CAP‐Gly domain reduces its binding to microtubules (Vaughan et al., 2002). These findings raise the possibility that other MAPs within this family may be regulated similarly. While the phosphorylation identified by Rickard and colleagues inhibits CLIP‐170 binding to microtubules, recent work has provided evidence that phosphorylation of CLIP‐170 by the rapamycin‐sensitive kinase mTOR promotes microtubule binding (Choi et al., 2002). The mechanism by which this phosphorylation stimulates microtubule binding is not yet clear, but it is interesting to speculate that it might inhibit formation of the closed, autoinhibited conformation of CLIP‐170, thus increasing CLIP‐170 microtubule‐ binding activity (Lansbergen et al., 2004). Even less is known about how Bik1p activity is regulated. However, there does appear to be a conserved interaction of Bik1p with TOR (the yeast orthologue of mTOR). In fact, treatment of yeast cells with rapamycin causes phenotypes similar to those of a bik1D mutant, suggesting that phosphorylation of Bik1p by TOR may be necessary for Bik1p function (Choi et al. 2000). Phosphorylation by other kinases is also certainly possible.

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VIII. Bik1p at the Kinetochore In a series of complementary experiments, the interactions of Bik1p at the kinetochore mirror those of the mammalian CLIP‐170 (Dujardin et al., 1998; Lin et al. 2001; Tai et al., 2002). Using chromatin immunoprecipitation, or ChIP analysis, Bik1p is found associated with centromeric DNA (Lin et al., 2001). For both Bik1p and CLIP‐170, the cargo‐binding domain is required for interaction with the kinetochore, while the CAP‐Gly domain is not (Dujardin et al., 1998; Lin et al., 2001). For CLIP‐170, the cargo‐ binding domain alone can be targeted to the kinetochore (Dujardin et al., 1998). Thus, these proteins are likely to bind kinetochores independently of their ability to bind microtubules, supporting the notion that these proteins may be actual elements of the kinetochore. Experiments measuring the dynamics of microtubule turnover within the spindle and anaphase kinetics suggest that the cargo‐binding domain of Bik1p is not involved in the assembly of the microtubules within the spindle. Bim1p and Stu2p have also been shown to be components of the kinetochore (He et al., 2001; Lin et al., 2001), but little is known about how or whether these elements function together with Bik1p at the kinetochore. For CLIP‐170, its association with the kinetochore is transient as it interacts at the kinetochores of unattached but not attached chromosomes (Dujardin et al., 1998). However, this may not be the case for Bik1p (Lin et al., 2001). Bik1p at the kinetochore is important for maintaining the attachment between the kinetochore and spindle microtubules. This function was elucidated using polyploid yeast, where the necessity of having extra chromosomes attached to the mitotic spindle may tax microtubule functions within the nucleus. In triploid cells lacking Bik1p, separation of preanaphase kinetochores is drastically reduced (Lin et al., 2001). Consistent with this model, BIK1 is essential in tetraploid cells, but bik1D mutants are viable in a haploid or diploid context (Lin et al., 2001). These findings have important implications for cancer biology in which cells often display increased ploidy (Lin et al., 2001).

IX. Bik1p in Genomic Studies In the last 5 years, many studies have been carried out in S. cerevisiae on a genome‐wide basis. In this section, we highlight several of these projects from which information on Bik1p (or other yeast proteins) can be obtained through simple computer searches. While individual pieces of data from genomics projects often need to be taken with ‘‘a grain of salt,’’ this information can be extremely useful when formulating new hypotheses, especially when new interactions have been identified.


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A. Synthetic Lethal Interactions In addition to being synthetically lethal with mutations in the Kar9p pathway, a genome‐wide synthetic lethal screen identified a long list of additional genes as being either synthetically lethal or synthetically sick in combination with bik1D mutations (Tong et al., 2004). This information, as well as links to other genomic projects, can be conveniently viewed online at the Saccharomyces Genome Database (SGD) (http://www.yeastgenome.org). A quick search on ‘‘BIK1’’ will bring the reader to a page containing a treasure trove of information on BIK1 (or another favorite gene). Most of the genes identified in this and previous synthetic lethal screens (Miller and Rose, 1998; Pellman et al., 1995) can be grouped into several functional categories described as follows. 1. Mitotic exit network and/or spindle assembly checkpoint genes. Mutations in DBF2, MAD2, BFA1, MAD1, BUB1, and BUB3 were found to be synthetically lethal with bik1D (Schwartz, et al. 1997; Tong et al., 2004). 2. Microtubule‐associated genes. BIM1, KAR9, CIN8, ASE1, and alleles of TUB1 and TUB2. cin8D was also identified previously as synthetically lethal with mutations in BIK1 in a PAC screen to identify genes that perish in the absence of cin8, a screen which also identified several other genes in the dynein pathway (Geiser et al., 1997). Mutations in BIM1 were previously identified as being synthetically lethal with BIK1 mutations, based on their respective roles in the Kar9p and dynein pathways (Miller and Rose, 1998; Schwartz et al., 1997). Ase1p is a microtubule bundling protein that plays an important function in elongation of the mitotic spindle and localizes at the spindle midzone (Schuyler et al., 2003). As Bik1p is also localized to the spindle microtubules, this synthetic phenotype is likely to be based on an overlap of function there (Pellman et al., 1995). 3. Factors and chaperones involved in alpha‐ and beta‐tubulin folding. Mutations in RBL2, PAC2, CIN1, CIN4, GIM4, GIM5, PAC10/GIM2, and YKE2/ GIM1 are synthetically lethal with bik1D (Archer et al., 1995; Bhamidipati et al., 2000; Geissler et al., 1998; Pellman et al., 1995; Stearns et al., 1990; Vainberg et al., 1998; You et al., 2004). (See Lopez‐Fanarraga et al., 2001; Szymanski, 2002 for reviews on tubulin folding.) These genetic interactions raise the possibility that Bik1p may also play an important role in the regulation of the levels or folding of tubulin monomer. 4. RNA polymerase, RNA splicing, and ribosomal functions. MSL1, RPA34, RIC1, RPL19b, PAT1, RTT103, RPL24A, and KEM1. While the basis for these synthetic lethal interactions is uncertain, the interactions with PAT1, RTT103, and KEM1 are especially notable because these three genes display synthetic interactions with several other microtubule‐associated proteins, in addition to BIK1.

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5. Transcription factors. SHC1 and HCM1. Shc1p is involved in chitin synthesis during sporulation (Hong et al., 1999; Sanz et al., 2002). Hcm1p is a member of the forkhead family of transcription factors. It is involved in the transcription of SPC110. Spc110p is an essential component of the SPB, a site to which Bik1p also localizes (Zhu and Davis, 1998; Zhu et al., 1993). 6. Other. Fab1p is a phosphatidylinositol 3‐phosphate 5‐kinase 1. The rationale for bik1D’s synthetic lethality with this kinase is unclear (Cooke et al., 1998; Yamamoto et al., 1995). B. Gene Expression Levels Patterns of gene expression are often useful in understanding the function of a protein. In S. cerevisiae, these patterns have been analyzed under a wide ranging set of diVerent conditions for the entire genome. As such, the expression levels of Bik1p have been reported for such conditions as release from various cell cycle arrests, treatment with the yeast mating pheromone, alpha factor; the microtubule‐dependent process of sporulation; drug and metal treatments; as well as many others. An updated index of these studies with hyperlinks is available at the ‘‘Expression Connection’’ page of the SGD (http://www.yeastgenome.org).

C. Quantitation of Bik1p Molecules in the Cell How many molecules of Bik1p are present in the yeast cell? Using a genomics approach, the answer to this question has been estimated to be 300 (Ghaemmaghami et al., 2003). For this analysis, a tandem aYnity purification or ‘‘TAP tag’’ was incorporated at the C‐terminus of each open reading frame at its endogenous genomic loci. This was then followed by quantitative Western blot analysis using antibodies to the tag. Because this was done on a genome‐ wide basis, estimates for each of Bik1p’s interacting binding partners can also be obtained at the website supporting this paper (http://yeastgfp.ucsf.edu/). Similar to mammalian cells, there is approximately ten times as many Bim1p molecules as Bik1p (3600 Bim1p vs. 300 Bik1p) (Tirnauer et al., 2002; Weiss and Goodson, unpublished). In yeast, there is twice as much Kar9p as Bik1p (656 vs. 300). The significance of these stoichiometries will undoubtedly come into play as models become more refined.

X. Concluding Remarks and Perspective In only the last few years, the phenomenon of microtubule plus‐end tracking has changed from a curiosity specific to one protein (CLIP‐170) to an activity that appears to have a basic role in the cell biology of many proteins.


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Recent work has begun to dissect the mechanisms used by these diVerent proteins to track plus ends, as well as the remarkable web of interactions between plus‐end tracking proteins. However, the exact function of many of these proteins has been diYcult to elucidate. Comparison between fungal and mammalian orthologues is a powerful and proven method for gaining insight into both universal and organism‐specific functions of a protein family. With this in mind, we have examined plus‐end tracking proteins from the perspective of Bik1p and its orthologue CLIP‐170. Although Bik1p and CLIP‐170 diVer in several ways, the parallels between them are likely to reflect fundamental functional similarities. It seems that whenever someone postulates that yeasts really ‘‘are diVerent’’ in a particular system, greater understanding reveals the remarkably common nature of fungal and mammalian cell biology. The immediate challenge for the future will be to decipher which of these diVerences are truly significant, and what these diVerences can tell us. Other key venues for research include establishing the functional significance of the web of interactions between plus‐end tracking proteins. The microtubule plus end has become a very crowded place! And while it is likely to get more so, at least in the near term, eventually the structure of the complex society at the microtubule plus end should come into clearer focus.

Acknowledgments Work in the Miller laboratory is supported by the National Science Foundation (#MCB‐ 0414768). The Goodson laboratory is supported by the National Institutes of Health grants (GM065420) and the American Heart Association through a Scientist Development Award.

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Aggregate‐Prone Proteins Are Cleared from the Cytosol by Autophagy: Therapeutic Implications Andrea Williams, Luca Jahreiss, Sovan Sarkar, Shinji Saiki, Fiona M. Menzies, Brinda Ravikumar, and David C. Rubinsztein Department of Medical Genetics Cambridge Institute for Medical Research Addenbrooke’s Hospital, Cambridge CB2 2XY United Kingdom

I. Proteinopathies A. Autophagy B. The Ubiquitin‐Proteasome Pathway II. III. IV. V. VI.

Intracytoplasmic Aggregate‐Prone Proteins Are Cleared by Autophagy Autophagy Induction Has Additional Antiapoptotic Consequences A New mTOR‐Independent Autophagy Pathway A Protective Role for Aggregates: Autophagy Upregulation Conclusions Acknowledgments References

Intracellular protein misfolding/aggregation are features of many late‐ onset neurodegenerative diseases, called proteinopathies. These include Alzheimer’s disease, Parkinson’s disease, tauopathies, and polyglutamine expansion diseases [e.g., Huntington’s disease; and various spinocerebellar ataxias (SCAs), like SCA3]. There are no eVective strategies to slow or prevent the neurodegeneration resulting from these diseases in humans. The mutations causing many proteinopathies (e.g., polyglutamine diseases and tauopathies) confer novel toxic functions on the specific protein, and disease severity frequently correlates with the expression levels of the protein. Thus, the factors regulating the synthesis and clearance of these aggregate‐prone proteins are putative therapeutic targets. The proteasome and autophagy‐ lysosomal pathways are the major routes for mutant huntingtin fragment clearance. While the narrow proteasome barrel precludes entry of oligomers/ aggregates of mutant huntingtin (or other aggregate‐prone intracellular proteins), such substrates can be degraded by macroautophagy (which we will call autophagy). We showed that the autophagy inducer rapamycin reduced the levels of soluble and aggregated huntingtin and attenuated its toxicity in cells, and in transgenic Drosophila and mouse models. Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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We extended the range of intracellular proteinopathy substrates that are cleared by autophagy to a wide range of other targets, including proteins mutated in certain SCAs, forms of ‐synuclein mutated in familial forms of Parkinson’s disease, and tau mutants that cause frontotemporal dementia/ tauopathy. In this chapter, we consider the therapeutic potential of autophagy upregulation for various proteinopathies, and describe how this strategy may act both by removing the primary toxin (the misfolded/aggregate‐prone protein) and by reducing susceptibility to apoptotic insults. ß 2006, Elsevier Inc.

I. Proteinopathies Intracellular protein misfolding and aggregation are features of many late‐onset neurodegenerative diseases, called proteinopathies. These include Alzheimer’s disease, Parkinson’s disease, tauopathies, and various diseases caused by abnormally expanded tracts of the amino acid glutamine, like Huntington’s disease (HD). HD is an autosomal‐dominant neurodegenerative disorder caused by a CAG trinucleotide repeat expansion, which results in an abnormally long polyglutamine (polyQ) tract in the N‐terminus of huntingtin protein (Huntington’s Disease Collaborative Research Group, 1993). Asymptomatic individuals have 35 or fewer CAG repeats, while HD is caused by expansions of 36 or more repeats (Rubinsztein et al., 1996). HD belongs to a group of human neurodegenerative disorders caused by CAG repeat/polyQ expansions, which includes spinocerebellar ataxias (SCAs) type 1, 2, 3, 6, 7, and 17, spinobulbar muscular dystrophy (SBMA), and dentatorubral pallidoluysian atrophy (DRPLA) (Nakamura et al., 2001; Zoghbi and Orr, 2000). The exact mechanism(s) of cellular toxicity caused by these proteins remains to be elucidated. However, it is generally accepted that a toxic gain‐of‐function is involved. Loss‐of‐function mutations often show phenotypes distinct from those seen in the disease, if any at all, while disease severity appears to correlate with expression level of the mutant protein. The toxicity of mutant huntingtin is thought to be exposed after it is cleaved to form N‐terminal fragments consisting of the first 100–150 residues containing the expanded polyQ tract, which are also the toxic species found in aggregates (Lunkes et al., 2002; Luo et al., 2005; Wellington et al., 2002). Thus, HD pathogenesis is frequently modelled with exon 1 fragments containing expanded polyQ repeats, which cause aggregate formation and toxicity in cell models and in vivo. All the known polyglutamine diseases are associated with the formation of intraneuronal inclusions (also known as aggregates) by the mutant proteins. The role of aggregates in these diseases is controversial and they have been suggested to be both toxic and protective in the cell. Aggregate toxicity may

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result from blockage of axonal transport or from the sequestration of transcription factors leading to transcriptional dysregulation (McCampbell et al., 2000; Nucifora et al., 2001). Equally, aggregates may protect the cell by sequestering putatively harmful soluble species of the mutant protein (Arrasate et al., 2004). However, no studies have shown the aggregates to be as beneficial as the wild‐type protein. Thus, even when aggregates appear to be less toxic than the soluble mutant protein, they may be deleterious relative to the wild‐type protein. Regardless of which of these scenarios is correct, reducing the load of toxic proteins in the cell is likely to be beneficial. Mechanisms by which this can be achieved are therefore of great interest from a therapeutic standpoint.

A. Autophagy In this chapter, we will review the ubiquitin‐proteasome and autophagy‐ lysosome routes for protein degradation and discuss how they may be involved in the clearance of aggregate‐prone proteins. We will use data with mutant huntingtin exon 1 as a starting point and then extend our observations to other proteinopathies. Two major hydrolytic systems mediate intracellular protein breakdown in mammalian cells—autophagy and the ubiquitin‐proteasome pathway. In addition to their importance in cellular homeostasis, these protein’s degradative pathways play a vital role in cellular quality control by degrading misfolded, damaged, or unassembled proteins (Reggiori and Klionsky, 2002). This is of significant importance in the context of neurodegenerative diseases that are characterized by the formation of aggregated proteins. Autophagy is usually defined as a nonspecific vacuolar degradative pathway that is ubiquitous in all eukaryotic cells. There are three primary forms of autophagy: macroautophagy, microautophagy, and chaperone‐mediated autophagy. Chaperone‐mediated autophagy is more selective than the other two pathways and involves the direct translocation of cytosolic proteins containing a pentapeptide‐targeting motif to the lysosome (Massey et al., 2004). Microautophagy is the least well‐characterized process and involves invaginations of the lysosomal membrane that sequester the portions of the cytoplasm to be degraded (Kim and Klionsky, 2000). Macroautophagy in contrast, is the most common form (from here on referred to as autophagy). Following an induction signal, autophagy starts when a flat membrane cistern wraps around a portion of cytoplasm, forming a double‐membrane‐bound vacuole. This vacuole is called the autophagosome, and is at this stage devoid of all lysosomal proteins. Fusion of the autophagosome with the lysosome results in the delivery of an inner vesicle into the lumen of the degradative compartment. Breakdown of the

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Figure 1 Outline of macroautophagy in mammalian cells. Cytoplasm is enclosed by the preautophagosome isolation membrane resulting in the formation of an autophagosome. The outer membrane of the autophagosome then fuses with the lysosome where the cytoplasm‐ derived materials are degraded.

inner vesicle membrane allows the degradation of the cytosolic cargo by lysosomal hydrolases and the subsequent delivery of the end products back to the cytoplasm (Fig. 1) (Klionsky and Ohsumi, 1999; Mizushima, 2004; Stromhaug and Klionsky, 2001). Induction of the autophagic pathway occurs as a cellular response to stress conditions such as nutrient starvation and hypoxia. Starvation‐ induced autophagy is in general believed to be nonselective with the aim of generating the building blocks needed by cells to survive starvation. Autophagy can, for example, allow haploid yeast cells to survive severe starvation for weeks, whereas, they would normally die within 2 days under the same conditions (Stromhaug and Klionsky, 2001). In mammalian cells, autophagy is tightly regulated both developmentally and nutritionally through several signaling pathways, the best characterized of which is the mammalian target of Rapamycin (mTOR) pathway. The mTOR is a negative regulator of autophagy that can sense nutrient and metabolic and hormonal signals through an upstream regulatory cascade via the Class 1 phosphatidylinositol‐3 kinase (Yorimitsu and Klionsky, 2005). Under nutrient‐rich conditions, mTOR is activated and autophagy is inhibited. However, under nutrient‐limiting conditions or when mTOR is inactivated by treatment with rapamycin, autophagy is activated. The details of the regulatory mechanisms and machinery required for mammalian autophagy are still poorly understood. However, much work has been focused in yeast, where the regulatory system is simpler. Genetic studies in yeast have identified several genes, termed atg, that are involved in the autophagic processes. Many of these genes have corresponding orthologues in higher eukaryotes (Levine and Klionsky, 2004).


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B. The Ubiquitin‐Proteasome Pathway The ubiquitin‐proteasome pathway, by comparison with autophagy, is highly selective and precisely regulated with diverse proteins being degraded at vastly diVerent rates ranging from several minutes to several days (Goldberg, 2003). Degradation through this route essentially involves a number of successive steps. In the first step, the substrate is targeted for degradation by the covalent attachment of multiple ubiquitin moieties to produce a polyubiquitin chain that acts as the proteolytic signal. Ubiquitin is a 76‐amino acid protein that is unusual in its ability to form stable chemical bonds with other proteins (Hicke and Dunn, 2003). Conjugation of ubiquitin to the substrate is a three‐step process requiring the sequential activity of three enzymes. In the first step, the ubiquitin‐activating enzyme, E1, activates ubiquitin in an ATP‐dependent manner to produce a high‐energy thioester bond between ubiquitin and an internal cysteine of E1. Ubiquitin can then be passed to the next enzyme, E2, an ubiquitin‐conjugating enzyme, by transthiolation. E2 normally operates in conjunction with the third enzyme, E3, an ubiquitin ligase, to transfer the ubiquitin to a lysine side chain residue on the substrate protein. The E3 enzymes, of which there exist several distinct families, are key to the conjugation process as they regulate the timing and selection of substrate. Subsequent rounds of ubiquitin conjugation can produce a chain of four or more ubiquitin molecules attached to the substrate (Hershko and Ciechanover, 1998). In the next step, the polyubquitin chain serves as a recognition signal allowing the substrate to be transported via a series of escort proteins to the 26S proteasome where it is degraded into peptides. The 26S proteasome is a 200‐kD complex which, in mammalian cells, contains a 20S proteasome as the key proteolytic component and two flanking 19S complexes, which contain several ATPases and a binding site for ubiquitin chains. The polyubiquitinated chain on the substrate protein is recognized by the 19S subunit, which unfolds the polypeptide and feed it into the proteolytic core of the 20S subunit. Inside, the polyubiquitin chains are cleaved from the substrate protein and subsequently recycled into monomeric ubiquitin units by deubiqitinating enzymes, while the substrate protein is degraded into small peptides. These peptides are further degraded into amino acids by cytosolic peptidases (Goldberg, 2003).

II. Intracytoplasmic Aggregate‐Prone Proteins Are Cleared by Autophagy Our initial studies used either an exon 1 fragment of huntingtin with 74 glutamines or 19 alanine repeats fused to GFP as model aggregate‐prone proteins and demonstrated that they can be cleared by both the proteasome

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and autophagy in cell culture (Ravikumar et al., 2002). Autophagy may be the preferential route of clearance of these proteins as the proteasome is unable to cleave within the polyglutamine tract (Holmberg et al., 2004; Venkatraman et al., 2004) and the narrow proteasome barrel cannot admit and degrade oligomeric species of mutant huntingtin (Verhoef et al., 2002). Conversely, autophagosomes can engulf oligomeric structures and even organelles like mitochondria. In addition, longer forms of mutant huntingtin, including the full‐length forms, have been shown to be autophagy substrates, while wild‐type full‐length mutant huntingtin appears not to have any significant dependence on autophagy for its clearance (Qin et al., 2003; Shibata et al., 2006). The observation that the clearance of these proteins can be increased by autophagy led us to test if one could decrease levels of these toxic proteins by upregulating autophagy. We initiated these studies using rapamycin, an inhibitor of mTOR, which is designed for long‐term clinical use in humans (for preventing kidney transplant rejection) and is in phase 3 trials for gliomas (Bjornsti and Houghton, 2004; Galanis et al., 2005; Lee and Chapman, 2005). Rapamycin or the rapamycin ester, CCI‐779, reduced levels of both soluble and aggregated mutant huntingtin and attenuated toxicity in cell, Drosophila, and mouse models of HD (Ravikumar et al., 2004). Thus, it was important to establish if this protective eVect was confined to huntingtin or could be more widely applied to aggregate‐prone proteins. Our data demonstrate that rapamycin treatment may indeed be eVective in the clearance of a diverse range of intracytosolic aggregate‐prone proteins (Berger et al., 2006). We have shown that not only can rapamycin induce the clearance of a range of polyglutamine‐ or polyalanine‐containing proteins (including proteins mutated in certain SCAs) and forms of ‐synuclein mutated in familial forms of Parkinson’s disease but also the clearance of mutant tau associated with frontotemporal dementia/tauopathy. Furthermore, we have shown that rapamycin reduces toxicity of many of these aggregate‐prone proteins in Drosophila (Berger et al., 2006). These findings suggest a much broader potential for rapamycin therapy, beyond HD. It is important to stress that enhancing autophagy results in more rapid clearance of both soluble and aggregated species of mutant huntingtin— thus, it does not matter if one believes aggregates are more or less toxic than the soluble forms. Our data with ‐synuclein mutants suggest that autophagy does eVectively remove soluble species, as these proteins do not form aggregates in our cell lines (Webb et al., 2003). While the removal of soluble huntingtin species would be suYcient to account for the reduction in aggregate levels (as the soluble species include aggregate‐precursors), we cannot exclude whether aggregates may also be cleared by autophagy. However, polyglutamine aggregates have not been generally observed to be membrane‐bound.


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III. Autophagy Induction Has Additional Antiapoptotic Consequences In addition to the protective eVect of rapamycin via enhanced clearance of aggregate‐prone proteins, our studies suggest that rapamycin can have other cytoprotective eVects by protecting cells and Drosophila against subsequent diverse apoptotic insults (Ravikumar et al., 2006). This protective eVect is, however, lost when autophagy is inhibited. There are two major apoptotic cascades in a cell, namely the intrinsic and extrinsic pathways. The intrinsic pathway requires mitochondrial‐dependent cytochrome c release for activation of downstream caspases, and is induced by insults like reactive oxygen species or UV irradiation. The extrinsic or death‐receptor‐mediated pathway is triggered by ligands binding to cell surface death receptors and in certain cell types the resultant caspase 8 activation leads to apoptosis via a pathway that is independent of mitochondria (and cytochrome c release). We found that pretreatment with rapamycin protected against apoptosis that occurred via the mitochondrial pathway, suggesting the involvement of mitochondria in this protective mechanism. No protective eVect was seen in a death‐receptor cell‐ death paradigm that was independent of mitochondria. Consistent with these data, the additional protective eVect of rapamycin was lost when cytochrome c release from mitochondria was blocked. We provided a plausible mechanism for the protective eVect exerted by rapamycin. We observed decreased levels of several mitochondrial proteins on rapamycin pretreatment, suggesting enhanced clearance of mitochondria, which are degraded via autophagy. After proapoptotic insults, we observed decreased levels of cytosolic cytochrome c and activated caspases in cells treated with rapamycin, consistent with reduced mitochondrial loads. We propose that the protective eVect of rapamycin can be accounted for by enhanced clearance of mitochondria by autophagy, thereby reducing cytosolic cytochrome c release and downstream caspase activation after proapoptotic insults (Ravikumar et al., 2006). These data suggest that proautophagic treatments may thus be useful in certain disease conditions, like some neurodegenerative diseases, where a slow but increased rate of apoptosis is evident. Autophagy induction may have two distinct beneficial eVects in protein conformational diseases. First, it enhances removal of the primary toxin causing these diseases—the aggregate‐prone protein. Second, it may attenuate apoptotic responses to various insults caused by such toxins, for example, excitotoxicity in HD.

IV. A New mTOR‐Independent Autophagy Pathway While rapamycin is the most specific kinase inhibitor known, its target, mTOR, controls many processes independent of autophagy (Wullschleger

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et al., 2006) and the way that mTOR regulates autophagy is still unclear. This results in side eVects with long‐term rapamycin therapy, including poor wound healing and some immunosuppression, although we are not aware of side eVects that have been attributed to enhanced autophagy in people taking rapamycin. Thus, the identification of more specific targets or non‐mTOR pathways for upregulating autophagy may be of great therapeutic value. We identified a novel mTOR‐independent autophagy‐enhancing pathway (Sarkar et al., 2005). Mammalian autophagy was shown to be regulated by the inositol signaling pathway, where the autophagic activity was increased by lowering intracellular inositol and myo‐inositol‐1,4,5‐triphosphate (IP3) levels (Fig. 2). Since inositol monophosphatase (IMPase) catalyzes the hydrolysis of inositol monophosphate (IP1) into free inositol (Maeda and Eisenberg, 1980), autophagy was induced by lithium specifically via its ability to inhibit IMPase, which results in depletion of free inositol and reduced IP3 levels (Berridge et al., 1989; Sarkar et al., 2005). Induction of autophagy by lithium enhanced the clearance of soluble and aggregated mutant huntingtin, and the A53T and A30P mutants of ‐synuclein. The increased clearance of mutant huntingtin by lithium correlated with a reduction in its aggregation and toxicity. Inositol depletion is a common mechanism of action of mood‐stabilizing drugs like lithium, carbamazepine (CBZ) and valproic acid (VPA), which all act in the brain (Williams et al., 2002). Consistent with a role for inositol depletion in autophagy regulation, CBZ and VPA also enhanced the clearance aggregate‐prone proteins like mutant huntingtin and ‐synuclein (Sarkar et al., 2005). This implicates that drugs depleting intracellular inositol levels may be potential therapeutic targets in HD and related neurodegenerative disorders. This novel autophagy pathway was independent of the well‐known mTOR pathway, since IMPase inhibition by lithium or the pharmacological inhibitor L‐690,330 did not aVect mTOR activity (Sarkar et al., 2005). Furthermore, raised intracellular inositol or IP3 levels did not prevent induction of autophagy by rapamycin; moreover, lithium could exert its protective eVects even when mTOR was activated. These data suggested that the regulation of autophagy by the inositol and mTOR signaling pathways were independent of each other. When autophagy was simultaneously induced by inhibition of mTOR (with saturating doses of rapamycin) and IMPase (by lithium or L‐690,330), there was greater clearance of mutant aggregate‐prone proteins and protection against their aggregation and toxicity, compared to either pathway alone (Sarkar et al., 2005). Therefore, autophagy can be upregulated to a greater extent by an additive eVect from two independent pathways, compared to maximal stimulation by a single pathway alone. This novel pharmacological strategy for enhancing autophagy may be beneficial for the treatment of various neurodegenerative diseases, where the toxic proteins are autophagy substrates (Berger et al., 2006;


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Figure 2 mTOR‐dependent and ‐independent pathways of enhancing autophagy. Autophagy is stimulated by rapamycin (rap) by inhibiting mTOR (indicated by arrow), which is downstream of the phosphatidylinositol‐3 kinase (PI‐3K) pathway. Stimulation of the PI‐3K pathway activates Akt, which then inhibits the tuberous sclerosis complex (TSC 1/2), thereby alleviating the inhibition on mTOR. Activation of mTOR inhibits autophagy, which retards the clearance of mutant aggregate‐prone proteins and increases their toxicity. Inhibition of inositol monophosphatase (IMPase) by lithium (LiCl) or a specific inhibitor L‐690,330 in the phosphoinositol signaling pathway also induce autophagy independent of mTOR activity. Inhibition of IMPase decreases inositol (Ins) and subsequently the myo‐inositol‐1,4,5‐triphosphate (IP3) levels, which stimulates autophagy. Induction of autophagy by either pathway enhances the clearance of mutant aggregate‐prone proteins and reduces their toxicity (indicated by arrows). Mood‐stabilizing drugs like carbamazepine (CBZ) and valproic acid (VPA), which deplete intracellular inositol levels, also facilitate clearance of mutant proteins. IP3 is generated by G‐protein–coupled receptor‐mediated stimulation of phospholipase C (PLC), which hydrolyzes PIP2 into IP3 and diacylglycerol (DAG). Increase in the levels of inositol (by myo‐inositol) or IP3 (by prolyl endopeptidase inhibitor) inhibits autophagy (see text). Simultaneous inhibition of mTOR (by rapamycin) and phosphoinositol signaling (by LiCl or L‐690,330) pathways have an additive eVect on autophagy and the clearance of aggregate‐prone proteins, compared to the eVect of a single pathway alone.

Iwata et al., 2005; Ravikumar et al., 2002, 2004; Sarkar et al., 2005; Shibata et al., 2006; Webb et al., 2003).

V. A Protective Role for Aggregates: Autophagy Upregulation Increased numbers of autophagosome‐like structures have also been reported in the brains of HD patients (Sapp et al., 1997). We confirmed in cell models of HD that there is an increase in the numbers of autophagosomes—however,

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this appeared to be a feature specifically of the cells with visible aggregates. We observed the same phenomenon with other mutant polyglutamine proteins like mutant ataxin‐1 (responsible for SCA type‐I). Our studies suggest that the increased autophagic activity in these contexts was the result of the sequestration of mTOR into the polyglutamine aggregates in cell and mouse models and in HD patients. We showed that this resulted in decreased soluble levels of mTOR in cells and in mouse models of HD, resulting in decreased mTOR activity (Ravikumar et al., 2004). As mTOR is a negative regulator of autophagy, these data can explain the enhanced numbers of autophagosomes seen in the cells with aggregates. Thus, increased autophagy can be associated with nontoxic consequences in HD and other polyglutamine diseases. This also suggests a probable protective role of aggregates, as they serve to induce autophagy, an important clearance route for mutant huntingtin. Of course, this is just one consequence of aggregates, and we cannot use these data to infer if aggregates have an overall protective or deleterious role.

VI. Conclusions Our data suggest that aggregate‐prone intracytosolic proteins are autophagy substrates and that their clearance can be enhanced by upregulating this clearance pathway. In fly and mouse models of HD, autophagy upregulation is associated with beneficial eVects. Ideally, we would like to initiate such treatment strategies in HD patients as early as possible with the aim of delaying onset of disease. If one can delay the symptoms of a disease that has a median onset of 40 years until after 90 years of age, then one has eVectively prevented onset. In monogenic diseases like HD, it is at least feasible to identify asymptomatic mutation carriers using genetic tests, as most will have a family history of disease. The challenge for the future will be to find the safest ways of inducing autophagy for decades in people, and to understand more about the presymptomatic changes caused by the HD and related mutations so that one can test the eYcacy of autophagy induction (or any other preventative strategy) in apparently unaVected mutation carriers.

Acknowledgments The work in our laboratory has been funded by a Wellcome Trust Senior Fellowship in Clinical Science, an MRC Program Grant, Wyeth and EU Framework VI (EUROSCA).


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Wnt Signaling: A Key Regulator of Bone Mass Roland Baron,* Georges Rawadi,{ and Sergio Roman‐Roman{ *Yale University School of Medicine New Haven, Connecticut 06520 { ProStrakan Pharmaceuticals 93230 Romainville, France

I. II. III. IV. V. VI.

VII. VIII. IX. X. XI. XII.

Introduction The Wnt Signaling Pathway LRP5 and ‐6: Structure and Function Human Genetic Evidence for the Role of Wnt Signaling in Bone Metabolism How Is the HBM Mutation Constitutively Activating LRP5 and Wnt Signaling? Mouse Genetics Confirms the Essential Role of the Wnt/ ‐Catenin Pathway in Bone A. Gain and Loss of Function of the LRP5/6 Receptors B. Genetic Alterations of Secreted Wnt Antagonists C. Genetic Alterations of Intracellular Components of Wnt Signaling How Is Wnt Signaling AVecting Bone Homeostasis? EVects of Wnts on Osteoblasts Is the Wnt Pathway the Final Common Pathway of Bone Morphogens? Is Wnt Signaling the Pathway Linking Mechanosensing to Bone Remodeling? Pharmacological Manipulation of Wnt Signaling in Bone Conclusion and Perspectives References

The identification of a link between bone mass in humans and gain‐ [high bone mass (HBM) trait] or loss‐of‐function [osteoporosis pseudoglioma (OPPG) syndrome] mutations in the Wnt coreceptor lipoprotein receptor‐ related protein (LRP)5 or in the Wnt antagonist sclerostin (sclerosteosis, Van Buchem syndrome) has called the attention of academic and industry scientists and clinicians to the importance of this signaling pathway in skeletal biology and disease. Multiple genetic and pharmacological manipulations of Wnt signaling in mice have since then confirmed the central role of this pathway in both the establishment of peak bone mass and its maintenance throughout life. Wnt signaling appears to be located downstream of bone morphogenetic proteins (BMPs), itself induced by Hedgehog (Hh) signaling, suggesting that it is the successive recruitment of these three intracellular signaling cascades that allow the full expression of the genetic patterns that characterize the osteoblast, the cell responsible for the formation of bone. ß 2006, Elsevier Inc. Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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I. Introduction Understanding the mechanisms by which bone formation is regulated has become one of the highest priorities in both academic and pharmaceutical research on the skeleton and a new frontier in the treatment of osteoporosis. Besides the academic challenge of unraveling processes, which govern not only the embryogenesis of the skeletal system but also bone growth, bone repair, and bone remodeling, is the medical need to reconstruct bone after injuries and, even more critical for the aging population, the medical need to not only stop bone loss with antiresorptive treatments in osteoporosis but also to rebuild bone mass and architecture to levels where the biomechanical function of the skeleton can safely be performed. The exploration of the mechanisms by which the formation of bone is regulated, whether during development or during the continuous remodeling of bone that occurs throughout life, has clearly established a role for members of the Hedgehog (Hh) family of proteins and their receptors (smothened and patch) as well as for bone morphogenetic proteins (BMPs) and their receptors (Chen et al., 2004; Chung et al., 2001; Long et al., 2004; Wan and Cao, 2005). Key downstream transcription factors required for the regulation of the expression of the genes that characterize the osteoblast lineage have also been identified, including Gli and Smads. In all instances it is the proliferation and the stepwise progression of mesenchymal precursors into progressively more committed osteoblasts precursors to become bone matrix‐secreting osteoblasts that ultimately determine the overall level of bone formation. Several marker genes of the osteoblast lineage are now identified and the picture that emerges is that of a cascade of signaling pathways each of which activates the expression of a few genes characteristic of the osteoblast lineage as well as the expression of the ligand for the following signaling cascade, ultimately leading to the expression of the full set of genes characteristic of a mature, bone matrix‐secreting osteoblast. In several instances, loss‐of‐ function mutations in humans lead to a characteristic genetic skeletal syndrome mimicked by gene deletion in mice. Deletion of runt‐related transcription factor 2 (Runx2) leads to a complete lack of bone formation in mice and to craniofacial dysostosis in heterozygous humans (Ducy et al., 1997; Mundlos et al., 1997; Otto et al., 1997); deletion of Osterix leads to an arrest along the osteoblast diVerentiation pathway and several skeletal abnormalities in mice (Nakashima et al., 2002). Although in humans Runx2 mutations are clearly linked to cleidocranial dysplasia (Mundlos, 1999; Mundlos et al., 1997) the link between these genes and the regulation of bone mass and/or the susceptibility to osteoporosis is still unclear at that point, despite some reports (Doecke et al., 2006; Vaughan et al., 2002). In contrast, and in what constitutes the most important breakthrough in this field in recent years, a clear link has now been established between

4. Wnt Signaling and Bone Mass

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lipoprotein receptor‐related protein (LRP)5, a coreceptor for Wnts, and bone mass in humans and in mice. Loss of function in LRP5 leads to the osteoporosis pseudoglioma (OPPG) syndrome, with extremely low bone mass, whereas gain of function leads to the high bone mass (HBM) phenotype in humans (Boyden et al., 2002; Gong et al., 1996; Little et al., 2002). In addition, deletion mutations in the gene encoding sclerostin (Sost), another endogenous inhibitor of the Wnt pathway, also lead to osteosclerotic phenotypes (osteosclerosis, Van Buchem syndrome) (Balemans et al., 2001, 2002; Brunkow et al., 2001; Staehling‐Hampton et al., 2002). These findings have opened a whole new field of investigation both in terms of understanding the mechanism that regulate osteoblasts and their bone matrix‐secreting activity and in terms of drug discovery in the hope to target one component of the Wnt signaling pathway and thereby increase bone mass in osteoporotic patients. The goal of this chapter is to briefly discuss our current understanding of Wnt signaling in bone and how it may aVect drug discovery in future years.

II. The Wnt Signaling Pathway The Wnt family includes a number of secreted, cysteine‐rich, glycoproteins involved in the regulation of a variety of cellular activities, including cell fate determination, proliferation, diVerentiation, migration, and apoptosis (Moon and Shah, 2002). Wnt signaling proceeds through two separate pathways. One is the ‐catenin or canonical pathway, and the other is the Ca2þ or noncanonical pathway. In the absence of Wnt (Fig. 1) ‐catenin is found in a large cytoplasmic complex comprising other proteins that promote its inactivation by phosphorylation and its proteasomal degradation. This large protein complex includes, besides ‐catenin, adenamotous polyposis coli (APC), glycogen synthase kinase (GSK)‐3 , and Axin. ‐Catenin degradation being phosphorylation dependent, activation of the canonical signaling (Fig. 2) allows ‐catenin to escape from the degradation process by inhibiting GSK‐3 , decreasing ‐catenin phosphorylation, and disrupting its binding to the inactivating complex. Increases in ‐catenin stability leads to a progressive increase in its cytoplasmic concentration, resulting in its nuclear translocation, where it interacts with T cell factor (Tcf)/lymphoid enhancing factor (Lef) transcription factors to regulate the expression of target genes (Moon and Shah, 2002; Moon et al., 2002; Staal and Clevers, 2000). At the cell membrane, Wnts need to bind two diVerent families of receptors to transduce the canonical signal. The first are low‐density LRP5 or ‐6 and the second are Frizzled (Fz) receptor family members (Bejsovec, 2000; Huelsken and Birchmeier, 2001; Mao et al., 2001a; Pinson et al., 2000; Tamai et al., 2004). LRP5/6 acts synergistically with Fzs to bind Wnt and

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Figure 1

Wnt signaling pathway: Inactive.

Figure 2

Wnt signaling pathway: Canonical pathway activated.


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activate downstream signaling (Holmen et al., 2002, 2005a). In contrast, activation of the Ca2þ signaling pathway by Wnts requires only binding to Frizzled receptors. The fact that both OPPG and the HBM phenotypes are LRP5 dependent therefore suggests that activation of the canonical pathway is both necessary and suYcient to regulate bone formation and bone mass. LRP5 and ‐6 are transmembrane proteins belonging to the low‐density lipoprotein receptor (LDLR) family. LRP5 and ‐6, as their homologue in Drosophila (Arrow), are required for Wnt‐induced ‐catenin signaling (Mao et al., 2001a; Pinson et al., 2000; Tamai et al., 2000). Frizzleds are seven‐ transmembrane proteins that display structural homology to G‐protein– coupled receptor (GPCR), but there is no clear evidence that Fzs transduce canonical signal through G‐proteins (Bhanot et al., 1996). At the N‐terminus of the Fz proteins, a highly conserved cysteine residue [cysteine‐rich domain (CRD)] sequence mediates at least in part Wnt ligand binding, whereas it is the third ‐propeller that binds Dickkopf (Dkk) proteins. As is the case for many potent signal transduction pathways, there are several regulatory proteins capable of dampening the eVects of the ligands when needed. Besides intracellular Wnt antagonists (e.g., APC, Axin), transmembrane and secreted proteins are also made by cells to control Wnt activity. Secreted frizzled‐related proteins (Sfrps) are Frizzled decoy receptors containing the CRD of frizzled, but no transmembrane segments. They are therefore able to bind in solution and sequestrate Wnts, inhibiting their interaction with both Fzs and LRP5/6 receptors (Finch et al., 1997; Leyns et al., 1997). Unlike Dkk proteins (see later) Sfrps bind to Wnts, preventing interaction of Wnts with both LRP5/6 and Fz receptors, inhibiting both canonical and noncanonical signaling (Hsieh et al., 1999). In contrast, Dkk proteins and Sost, secreted proteins which are also potent inhibitors of Wnt signaling, do not bind to the Wnt family of ligands but rather to LRP5 or ‐6, and consequently inhibit specifically the Wnt canonical pathway. The fact that deletion mutations of Dkk1 (Morvan et al., 2006) or of Sost (Balemans et al., 2001, 2002; Brunkow et al., 2001; Staehling‐Hampton et al., 2002) lead to HBM therefore confirms that it is the canonical pathway that regulates bone formation. There are three members of the Dkk family of proteins. Dkk1 has been shown to bind the third ‐propeller domain of LRP5 and to act as an absolute Wnt antagonist (Glinka et al., 1998). Dkk2 also binds to LRP5 and ‐6, yet depending on cellular context it might have dual activity, inhibiting or activating the canonical pathway (Li et al., 2002). Dkk3 belongs to the Dkk family but fails to bind to LRP5/6 and to inhibit Wnt activation (Krupnik et al., 1999). In addition to LRP5/6, Dkk proteins require the presence of another receptor, Kremen, to inhibit Wnt signaling. Dkk actually binds to both Kremen and LRP5/6 and the Dkk–Kremen–LRP5/6 protein complex is internalized and degraded thus depleting the cell surface from the LRP5/6 receptor, essential for Wnt canonical signaling (Mao et al., 2002). Although sclerostin also

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mediates its antagonistic activity through association with LRP5/6, not much is known regarding the precise interaction sites, the requirement for additional coreceptors or partner proteins (such as Kremen), or the mechanism by which it inactivates the receptor. Thus, Wnt signaling via the canonical pathway requires the presence of LRP5/6 at the cell surface of target cells and gain or loss of function in LRP5 has been linked to dramatic changes in bone mass, eliciting major interest in this receptor.

III. LRP5 and ‐6: Structure and Function Human LRP5 and ‐6 are highly homologous receptors, sharing 71% amino acid identity (Brown et al., 1998; Dong et al., 1998; Hey et al., 1998; Kim et al., 1998), and together with their Drosophila homologue Arrow, form a subfamily of the of the LDLR family (Wehrli et al., 2000). LRP5 and ‐6 are type I single‐span transmembrane proteins with 1615 and 1613 amino acid residues, respectively. They display in the extracellular portion the structural domains defining the LDLR family: the YWTD‐type ‐propeller domain, the epidermal growth factor (EGF)‐like domain, and the LDLR ligand‐binding motif. LRP5/6 has four YWTD ‐propeller domains that are each followed by an EGF‐like domain, which comprises approximately 40 amino acids with conserved cysteine residues; these domains are then followed by three LDLR ligand‐binding motifs. The cytoplasmic domains of LRP5/6 (207 and 218 amino acids respectively) lack recognizable catalytic motifs and share no sequence similarity with other LDLR proteins. They are rich in prolines and serines and contain five reiterated PPP(S/T)P motifs (for a review of LRP5/6 structure see He et al., 2004). The crucial role played by LRP5 and ‐6 in Wnt signaling was originally identified by genetic studies. Drosophila mutants lacking arrow phenotypically resemble the Wg‐dependent mutants (Wehrli et al., 2000), and mouse embryos homozygous for an insertion mutation in the LRP6 gene exhibit developmental defects that are a striking composite of those caused by mutations in individual Wnt genes (Pinson et al., 2000). Furthermore, LRP6 activates Wnt signaling in Xenopus embryos and induces Wnt responsive genes, dorsal axis duplication, and neural crest formation (Tamai et al., 2000). As mentioned earlier, and unlike Fz, which is required for multiple Wnt pathways (Strutt, 2003; Veeman et al., 2003) Arrow/LRP5/6 seem to be specifically required for Wnt/ ‐catenin signaling (Fig. 3). Several reports have addressed the mechanism by which LRP5/6 signals within the ‐catenin pathway. First, Wnt binds to both Fz and to the first ‐propeller of LRP5/6, inducing conformational changes in the receptor cytoplasmic domain (Cong et al., 2004; Liu et al., 2003). Expression of an extracellular truncated LRP5/6 protein behaves as a constitutively active receptor (Mao et al., 2001a,b), indicating that activation is normally


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The LRP5 receptor, HBM and OPPG mutations, and binding partners.

prevented by the extracellular domain in the absence of Wnt ligand. These modifications of the cytoplasmic tail induce the recruitment and interaction of intracellular proteins, including GBP/Frat1 and Axin, one of the components of the ‐catenin inactivation complex together with GSK‐3 and APC (Hay et al., 2005; Mao et al., 2001b). As indicated earlier, Axin acts as a scaVold for APC, ‐catenin, GSK‐3 , and possibly other proteins, allowing the association of these inactivating proteins with—and the phosphorylation of— ‐catenin. Recruitment of Axin and Frat1 to the cytoplasmic tail of the activated LRP5/6 receptor leads to the inhibition of GSK‐3 by Frat1 and destabilization of the Axin/GSK‐3 / ‐catenin complex, thus preventing ‐catenin degradation by proteasomes (Hay et al., 2005). This occurs in parallel with the recruitment of Dissheveled (Dsh) by Wnt‐activated frizzled, contributing to the inhibition of ‐catenin phosphorylation and degradation, leading to its nuclear translocation and transcriptional activity. Although LRP5 and ‐6 were often considered as two redundant proteins it is now evident that there are some important diVerences between these two receptors. First, although both are widely expressed, Lrp5 or Lrp6 deficiency does not result in the exact same phenotype. Deletion of LRP6 leads to perinatal lethality and defects similar to those reported for Wnt1, Wnt3, or Wnt7a knockout animals (Pinson et al., 2000). In contrast, Lrp5/ mice

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develop normally during embryogenesis but display low peak bone mass and later develop osteoporosis and metabolic disorders (Clement‐Lacroix et al., 2005; Fujino et al., 2003; Kato et al., 2002; Magoori et al., 2003). These data clearly suggest that although the two receptors are partially redundant, during embryogenesis LRP6 plays a role that cannot be compensated by LRP5, whereas in the postnatal skeleton LRP5 and ‐6 overlap partially but play additive roles (see later). A key diVerence between these two receptors is their respective aYnity for Dkk proteins with LRP5 and ‐6 displaying distinct binding aYnity for Dkk1 and Dkk2 (Mao and Niehrs, 2003; Mao et al., 2001a, 2002). It remains to be determined whether these receptors also have diVerent aYnities for the various members of the Wnt family of proteins, which could also explain the diVerences observed in genetic studies. Finally, traYcking and membrane localization also diVer between LRP5 and ‐6. Forced expression of LRP6 results in massive localization of this receptor at the cell membrane (Mao and Niehrs, 2003; unpublished data) whereas overexpression of LRP5 most often does not lead to the accumulation of this receptor at the cell membrane but rather in cytoplasmic vacuoles (Zhang et al., 2004; unpublished data). The chaperonin Mesd has been shown to interact with the third ‐propeller of LRP5 or ‐6 and is required for proper targeting of the receptor to the membrane (Zhang et al., 2004). Further analysis will be required to understand whether these diVerences play any role in the respective roles of LRP5 and ‐6 in the regulation of bone development and bone mass.

IV. Human Genetic Evidence for the Role of Wnt Signaling in Bone Metabolism A pivotal role for members of the Wnt gene family in the process of skeletal development was established prior to the discovery of the link between LRP5 and bone mass. Initially, this was attributed to the involvement of Wnt proteins in chondrocyte diVerentiation in early mesenchymal condensations, aVecting skeletal maturation and, later, bone growth. During embryonic development, abnormal expression of Wnt family members and/or their regulatory molecules can disrupt the Wnt‐controlled balance and results in skeleton malformation (Church and Francis‐West, 2002). But the link to peak bone mass and its maintenance has emerged only recently from human genetic analysis and from transgenic animal models. The OPPG syndrome is an autosomal recessive disorder involving both skeletal and eye abnormalities (Gong et al., 1996). AVected patients display very low bone mass and are prone to developing fractures and deformations with no significant defects in calcium homeostasis or bone turnover


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parameters (Gong et al., 1996, 2001). Obligate carriers of OPPG mutations, although not strongly aVected, have a lower bone mass and an increased incidence of fractures. In addition to the skeletal abnormalities, many patients with OPPG mutations are born with severe disruption of an ocular structure (Gong et al., 2001). Early studies by Warman et al. (Gong et al., 1996) had assigned the OPPG locus to the 11q12–13. In a separate study, Johnson et al. (1997) later also mapped the HBM trait, inherited as an autosomal dominant recessive trait with aVected individuals being clinically healthy, to the same chromosomal region. These studies clearly linked this chromosomal region to the establishment and/or maintenance of bone mass, strongly suggesting the possibility that the same gene was aVected in opposite manners in both OPPG and HBM. Positional cloning later allowed both groups to identify mutations in the Lrp5 gene as the cause of OPPG as well as HBM (Boyden et al., 2002; Gong et al., 2001). All mutations identified in OPPG patients result in LRP5 loss of function whereas a gain‐of‐function mutation was identified in HBM, albeit, as discussed later, it is not intrinsic to the receptor but rather aVecting bonding aYnity of secreted negative regulators (see later). In a report, missense mutations in Lrp5 have also been associated with Van Buchem disease, autosomal dominant sclerosteosis, and osteopetrosis type I syndromes (Van Wesenbeeck et al., 2003). Like in HBM, these mutations are located in the first ‐propeller of LRP5, suggesting that it might lead to a similar gain of function of this receptor, possibly linked to altered aYnity for the Wnt antagonists Dkk1 and/or Sost (see later). Sclerosteosis and Van Buchem diseases are characterized by generalized overgrowth of bone tissue, mostly apparent in cranial bones and in the diaphysis of long bones (Hamersma et al., 2003), and maybe caused by mutations in either the LRP5/6 receptors (Van Wesenbeeck et al., 2003) or in the antagonists Sost. Loss of function of the Sost gene, which encodes for sclerostin, a secreted protein which is selectively expressed by osteocytes and some osteoblasts (Poole et al., 2005; van Bezooijen et al., 2004), is associated with sclerosteosis (Balemans et al., 2001; Brunkow et al., 2001), while downregulation of the same gene can cause Van Buchem disease (Balemans et al., 2002; Staehling‐Hampton et al., 2002). As previously discussed, sclerostin is, like Dkk, a ligand of the LRP5/6 receptors and behaves as an antagonist of the canonical Wnt pathway.

V. How Is the HBM Mutation Constitutively Activating LRP5 and Wnt Signaling? Although the HBM mutation Lrp5 G171V has been reported to increase Wnt signaling in vitro and in vivo (Boyden et al., 2002), the precise molecular

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mechanism by which this mutation results in an enhancement of Wnt signaling and subsequently in an increase in bone mass is still a matter of debate. Cells transfected with Lrp5 G171V do not respond better to Wnt than do cells transfected with the wild‐type cDNA, and this mutation does not confer a constitutive activity to Lrp5 (Boyden et al., 2002). It has, however, been shown that Lrp5 G171V mutation impairs the antagonism of Wnt signaling by Dkk1 (Ai et al., 2005; Boyden et al., 2002; Zhang et al., 2004), probably by altering the aYnity of Dkk1 for LRP5. As shown by our observation in Dkk1 happloinsuYcient mice, such a decrease in aYnity could well be suYcient to increase Wnt signaling in target cells and, thereby, increase bone mass (Morvan et al., 2006). The HBM mutation may also result in a reduction in the number of LRP5 molecules at the cell surface via a disruption in the interaction of Lrp5 with its chaperone protein Mesd (Zhang et al., 2004). The identification of other missense mutations in Lrp5 which result in an increased bone density (Van Wesenbeeck et al., 2003) and the systematic study of how the corresponding proteins reach the cell surface and aVect Wnt signaling confirm that these mutations confer a reduced aYnity to and inhibition by Dkk1 (Ai et al., 2005). Thus, although the eYciency of the diVerent mutants to transit the cell and reach the surface varies from a mutant to another, all were capable of transducing Wnt signaling at the same level as wild‐type Lrp5. Taken together, these results firmly established in humans that conditions with an increased bone density are often associated with gain‐of‐function mutations in the LRP5 gene or loss of function of one of the LRP5 antagonists, such as sclerostin, whereas loss of function in LRP5 leads to low bone mass. These observations have elicited a strong eVort to better understand the mechanisms involved through the analysis of the bone phenotype of mice genetically manipulated to reduce or enhance the expression of distinct elements of the Wnt cascade.

VI. Mouse Genetics Confirms the Essential Role of the Wnt/b‐Catenin Pathway in Bone A. Gain and Loss of Function of the LRP5/6 Receptors Deletion of the Lrp5 gene in mice leads to a phenotype essentially identical to what is observed in OPPG patients: low bone mass due to decreased bone formation and a persistent embryonic eye vascularization (Kato et al., 2002). Lrp6‐ deficient mice are not viable, emphasizing LRP6‐specific roles in embryogenesis. Nevertheless, Lrp6 haploinsuficiency also leads to a significant reduction in bone mass and, in double LRP5/6 mutants, further accentuates the low bone mass phenotype observed in Lrp5‐deficient mice (Fig. 4).


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LRP5‐null mice form less bone, lose bone, and mimic OPPG.

These results show that Lrp6 participates in the control of bone mass accrual in more ways than a simple redundancy with Lrp5 (Holmen et al., 2004). In contrast, overexpression of LRP5 in bone only results in a modest increase in bone mass, the eVects being most likely rate‐limited by endogenous levels of Wnts (Babij et al., 2003). Finally, transgenic mice expressing the human LRP5 gain‐of‐function mutation (G171V) specifically in bone display a HBM phenotype very similar to that described in humans with the autosomal dominant HBM trait (Babij et al., 2003). Thus, gain‐ or loss‐of‐function mutations of the LRP5 and/or ‐6 in mice can regulate bone mass and, for the most part, recapitulate the OPPG or HBM phenotypes in humans, confirming the essential and highly conserved nature of this regulatory pathway.

B. Genetic Alterations of Secreted Wnt Antagonists An increase in trabecular bone mineral density and mineral apposition rate has also been reported in mice deficient for the secreted Wnt antagonist

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Sfrp1 in the absence of any other apparent phenotype outside the skeleton (Bodine et al., 2004). This moderate phenotype does not aVect younger animals but is evident only in adult animals. Actually, Sfrp1 deletion prolongs trabecular bone accretion while trabecular bone mineral density declines with age in the control littermates. In contrast, in mice overexpressing other secreted Wnt antagonists, WIF‐1 or Dkk1, display a decreased bone mineral density and an increased susceptibility to bone fractures (Cao et al., 2004; Li et al., 2004). As for LRP6 but in an opposite manner, Dkk1 haploinsuficiency induces a dramatic increase in trabecular bone and bone formation rate (Morvan et al., 2006), demonstrating that simply shifting the endogenous balance between Wnts, their receptors, and/or their antagonists is suYcient to induce an anabolic response in the skeleton. Thus, genetic alterations of Wnt antagonists in mice not only reproduce the human phenotypes of gain and loss of function of LRP5 but also demonstrate that activation of Wnt signaling by decreasing the levels of expression of endogenous Wnt antagonists is suYcient to induce a HBM phenotype. The only exception to this general rule is a report showing that Dkk2 deficiency in mice results not in an increased bone mass as would be expected but rather in osteopenia (Li et al., 2005). This unexpected observation suggests that either Dkk2 plays a diVerent role than Dkk1 in skeletal development and remodeling or that the promoters that were used to conditionally delete this Dkk family member aVected a very specific time point during the diVerentiation of osteoblasts, leading to decreased bone formation. Further work will be required to clarify this important point.

C. Genetic Alterations of Intracellular Components of Wnt Signaling Since genetic manipulation of secreted or extracellular domains of membrane elements of the Wnt cascade aVect bone formation, as discussed earlier, modulation of intracellular players in mice was expected to have similar eVects. Surprisingly, if osteoblast‐specific APC deficiency and overexpression of a stabilized ‐catenin, both of which result in increased levels of ‐catenin in the cytoplasm and translocation to the nucleus, increased bone mass; this appears to have been mostly not only due to an increase in bone formation, as expected, but also due to a reduction in the number of osteoclasts (Glass et al., 2005; Holmen et al., 2005b). Further supporting the hypothesis that ‐catenin is also involved in the regulation of osteoclast diVerentiation, an increase in the osteoclast number is observed in osteoblast‐selective ‐catenin–deficient mice and Tcf‐deficient mice (Glass et al., 2005). Although initially surprising, these results simply confirm the well‐ established fact that osteoblasts also control osteoclasts via the regulation of


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osteoprotegrin (OPG) and RANK ligand (RANKL) (Boyle et al., 2003). The relative levels of OPG and RANKL were altered in these mutant mice (Glass et al., 2005; Holmen et al., 2005b). It is, however, important to note here that pharmacological manipulation of the ‐catenin inactivating complex by GSK‐3 inhibitors or lithium chloride, which also result in stabilization of ‐catenin, yet not in an osteoblast‐specific manner, seem to aVect mostly osteoblasts and bone formation rather than osteoclasts (see later). It is therefore possible that stabilization of ‐catenin not only in osteoblasts but also nonspecifically in other cells prevents the osteoblast‐mediated decrease in osteoclast diVerentiation while maintaining the stimulation of bone formation. These observations could be of importance in the context of pharmacological manipulation of Wnt signaling in order to increase bone mass in patients.

VII. How Is Wnt Signaling Affecting Bone Homeostasis? The bone phenotypes of the OPPG and HBM human mutations and of the transgenic models described earlier clearly established that the osteoblast lineage is the main direct cellular target of Wnt signaling in bone. All the elements of the Wnt signaling cascade (including Wnts, frizzleds, LRP5/6, ‐catenin, and TCF/Lef) are expressed in osteoblasts (Hens et al., 2005; Kato et al., 2002), and TOPGAL mice (mice expressing a transgene consisting of a ‐galactosidase gene under the control of a TCF responsive element) (DasGupta and Fuchs, 1999) show that the canonical Wnt cascade is active during bone development and growth in osteoblasts as well as in osteocytes and chondrocytes (Hens et al., 2005). With age, Wnt activation seems to decrease in osteoblasts but remains prominent in chondrocytes and osteocytes, an observation of importance in the context of the restricted expression of Sost to a subset of osteocytes (Poole et al., 2005) and their recognized role in mechanostransduction (Burger and Klein‐Nulend, 1999) and the fact that mechanical deformation has been shown to induce TOPGAL transgene expression in primary calvarial osteoblasts (Hens et al., 2005). A number of studies have also shown a marked increase in the expression of extracellular Wnt antagonists, such as Dkk1, Dkk2, Wif‐1, Sfrp2, and sclerostin, during the late phase of osteoblast diVerentiation (Li et al., 2005; Poole et al., 2005; Vaes et al., 2005; van Bezooijen et al., 2004; Zhang et al., 2004), suggesting that inhibition of Wnt signaling in osteoblasts is a necessary step for the terminal maturation of osteoblasts and mineralization of the extracellular matrix (van der Horst et al., 2005). Such a time‐dependent role of Wnt signaling in osteoblast maturation could explain the unexpected changes in bone mass in Dkk2 knockout (Fig. 5).

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Figure 5 Schematic representation of the eVects of Wnt signaling on osteoblast diVerentiation and function and, indirectly, on osteoclast diVerentiation and activation.

VIII. Effects of Wnts on Osteoblasts Osteoblasts are derived from pluripotent mesenchymal precursors, which have the capacity to diVerentiate into other cell lineages including adipocytes, myocytes, chondrocytes, and adipocytes (TriYt, 1996). The Wnt canonical signaling has been shown to play a major role in the fate of mesenchymal precursors both during embryogenesis and adult life. During embryonic development Wnt/ ‐catenin signaling represses the diVerentiation of mesenchymal cells into


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chondrocytes and adipocytyes and promotes osteoblast diVerentiation (Day et al., 2005; Hill et al., 2005; Hu et al., 2005; Rawadi et al., 2003; Ross et al., 2002). Overexpression of Wnt10a in mouse marrow leads to a dramatic inhibition of adipocytes and an increase in bone mass with an increased number of osteoblasts (Bennett et al., 2005). Similarly, treatment of SAMP6 mice with the GSK‐ inhibitor lithium chloride results in a significant reduction of adipocytes in the marrow accompanied by an increase in the number of osteoblasts and bone mass (Clement‐Lacroix et al., 2005), suggesting that the enhancement of osteoblast diVerentiation may occur at the expense of adipocyte progenitors. An enhancement of bone formation can result from an increase in the number of osteoblasts and increase in their rate of diVerentiation and/or an increase in their intrinsic activity. Genetic and pharmacological studies (see later) show that activation of Wnt signaling leads to an increase in the number of osteoblasts. The only in vivo proliferation data reported so far shows that the Lrp5/ osteoblasts display a low mitotic index, suggesting that Wnt signaling regulates osteoblast proliferation positively (Kato et al., 2002), as well as increasing the commitment of precursors and the progression toward fully diVerentiated cells. Wnt signaling is also known to have a strong antiapoptotic function, and apoptosis of osteoblasts is decreased in animal models (Chen et al., 2001) and a significant reduction in the number of apoptotic osteoblast and osteocytes is observed in Lrp5 G171V transgenic mice (Babij et al., 2003). In addition, the Wnt antagonists sclerostin and sFRP1 induce apoptosis in osteoblasts (Bodine et al., 2005; Sutherland and Musgrove, 2002). Thus, the Wnt canonical pathway can control not only the proliferation and diVerentiation of osteoblast precursors but also the survival of mature osteoblasts. In addition to their eVects on the osteoblast lineage determination, proliferation and apoptosis, in vitro models strongly suggest that Wnt signaling modulates the transcription of genes essential for bone matrix synthesis and mineralization (Rawadi et al., 2003). Only Wnts signaling via the canonical pathway is able to increase the expression of alkaline phosphatase in mesenchymal pluripotent cell lines (Gong et al., 2001) and Wnt3a has been shown to increase the expression of collagen type I and osteopontin in the mesenchymal cell line C3H10T1/2 (Hu et al., 2005).

IX. Is the Wnt Pathway the Final Common Pathway of Bone Morphogens? How does the Wnt pathway interact with other pathways known to play important roles in controlling osteoblast function and activity?

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To some extent, the ability of morphogens of the BMP and Hh protein families to control osteoblast diVerentiation/function is dependent on the capacity of these proteins to induce Wnts (Hu et al., 2005; Rawadi et al., 2003). Therefore, the induction of alkaline phosphatase (ALP) activity by BMPs or Hh is significantly decreased in the presence of the Wnt antagonist Dkk1 demonstrating an autocrine loop for the control of the expression of ALP. Although Dkk1 inhibits mineralization of MC3T3‐E1 cells (Rawadi et al., 2003), it has been proposed that downregulation of Wnt signaling by Dkk1 and Dkk2 is necessary for late‐stage osteoblast diVerentiation of KS483 osteoblast‐like cells (van der Horst et al., 2005). Thus, the three families of Wnt, Hh, and BMP morphogens frequently interact and regulate one another’s expression or function during the development of many tissues. BMPs are general targets of Hh signaling. The Drosophila BMP ortholog, decapentaplegic (dpp), is induced by Hh (Dominguez et al., 1996) and its transcription is regulated by the transcriptional mediator of Hh signaling Cubitus interrupts (Ci) (Aza‐Blanc and Kornberg, 1999; Methot and Basler, 1999). There are also examples where Hh is both upstream and downstream of BMPs (Zhang et al., 2000). Wnt5a has been identified as a target of Shh in hair follicle morphogenesis (Reddy et al., 2001), and the finding that Gli1 consistently induces a distinct set of Wnt genes in animal cap explants and in skin tumors suggests that Wnt regulation by Gli proteins is a general phenomenon (Mullor et al., 2001). Finally, Wnt signaling inhibits neural diVerentiation of embryonic stem cells by controlling BMP expression (Haegele et al., 2003). Because bones display continuous regeneration in adult life, it is not so surprising that most of the morphogens are expressed in adult bones (mainly by osteoprogenitors and mature osteoblasts) and that all these families of proteins play a crucial role in postnatal bone accrual, bone remodeling, and fracture healing. As it is the case for a series of tissues during the development, the interconnection between the signaling cascades triggered by Wnt, Hh, and BMP family members seem to be very important to control bone homeostasis in adults. Distinct autocrine/paracrine loops have been proposed to play a role in osteoblast diVerentiation. First of all, Hh signaling controls BMP expression in osteoprogenitor cells. Whereas full‐length Gli3 increases BMP‐2 promoter activity, truncated Gli3 markedly represses BMP‐2 gene transcription (Haegele et al., 2003). The mammalian orthologs of Ci, Gli1, and Gli3, have been demonstrated to stimulate luciferase activity driven by either BMP‐4 or BMP‐7 promoters in cotransfection experiment conducted in COS‐7 cells providing direct evidence that the Gli proteins are capable of inducing the BMP expression (Kawai and Sugiura, 2001). Wnt signaling functions downstream of Indian hedgehog (Ihh) in the development of osteoblast lineage, and Wnt7b has been identified as a potential endogenous ligand regulating


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osteogenesis (Hu et al., 2005). Finally, integrity of Wnt signaling is a requisite for induction of osteogenesis by Hh and BMP, and Wnt1 and Wnt3a gene expression is induced by BMP‐2 in mesenchymal pluripotent cells (Rawadi et al., 2003). It therefore appears that the Wnt signaling pathway is downstream of two of the main signaling cascades regulating skeletal development, Hh and BMPs. Each of these pathways target a specific set of genes in osteoblast precursors, including the ligand for the next pathway: Hh induces BMPs and BMPs induce Wnts. It is only when all three are fully functional, as a result of the paracrine balance between the ligands and their endogenous antagonists that all the genes that define a mature, bone matrix‐producing osteoblast can be expressed. Often, the ligands also increase the expression of their own antagonists. Given that the skeleton continues to remodel after embryogenesis and growth, these local cascades will play a major role in establishing peak bone mass and preventing bone loss in adults, acting on osteoblasts to regulate both bone formation and bone resorption. Bone remodeling being in large part regulated by the biomechanical forces exerted on the skeleton, Wnt signaling may be a key molecular link between mechanical stimulation and bone formation and remodeling.

X. Is Wnt Signaling the Pathway Linking Mechanosensing to Bone Remodeling? A link between Wnt signaling and the response to mechanical loading has been hypothesized from several observations. First, bones from Lrp5‐deficient mice and Lrp5‐G171V transgenic mice display, respectively, a reduction or an increase in the response to loading (Robinson et al., 2003; Sawakami et al., 2004). Second, mechanical loading activates Wnt signaling in osteoblasts in culture (Hens et al., 2005). But importantly the link to mechanosensing may be more the fact of osteocytes than of osteoblasts. Osteocytes play a crucial role in sensing mechanical forces in the skeleton, and not only express all the components of Wnt signaling but also they are the main source of sclerostin in bone, where canonical Wnt signaling in mature bone is almost exclusively active in osteocytes (Hens et al., 2005). There is therefore a strong possibility that the main role of Wnts in the adult skeleton is linked to osteocytic mechanosensing sending Wnt‐dependent signals to osteoblasts, and possibly to osteoclasts, thereby translating mechanical forces into the regulation of the process of bone remodeling, allowing the mechanical adaptation of the skeleton and individual bones not only in terms of bone mass but also bone shape and geometry. In turn, this would provide an exceptional

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opportunity to regulate bone mass and the biomechanical function of the skeleton through pharmacological intervention.

XI. Pharmacological Manipulation of Wnt Signaling in Bone There are numerous targets for pharmacological intervention within the Wnt pathway: the receptors, the secreted antagonists, and intracellular kinases oVer significant opportunities for drug discovery, which could include small chemical compounds, neutralizing antibodies, as well as small mimicking peptides. One of the most ‘‘drugable’’ targets in the Wnt pathway is GSK‐3 , a multifunctional serine/threonine kinase expressed in all eukaryotic cells (Doble and Woodgett, 2003). This enzyme is, however, a key regulator of many signaling pathways, including Wnt, several receptor tyrosine kinases, and GPCRs. This is an atypical kinase because it is constitutively active and is mainly regulated through inhibition. Given the association of abnormal function of GSK‐3–dependent pathways and numerous human diseases, GSK‐3 is a potential therapeutic target besides bone diseases, for example, in neurodegenerative diseases and in diabetes. One of the best‐known GSK‐3 inhibitor used in clinics is lithium chloride (Klein and Melton, 1996). Lithium inhibits GSK‐3 through competition with Mg2þ, but it is also able to inhibit other enzymes. Several more specific and selective inhibitors of GSK‐3 have also been identified which compete for ATP binding (Martinez et al., 2002). In vitro GSK‐3 inhibitors induce ‐catenin stabilization and its translocation to the nucleus as well as activation of Wnt target genes, thus mimicking to a great extent Wnt activity. When tested in vitro, GSK‐3 inhibitors (including lithium), like Wnt, are able to stimulate alkaline phosphatase expression in osteoblast progenitors (Clement‐Lacroix et al., 2005). As would be expected of GSK‐3 in vivo, treatment of wild‐type mice with lithium chloride resulted in a significant increase in bone mass. A marked increase in bone mass was also obtained when treating osteopenic animals, such as Lrp5/ or SAMP6, with lithium (Clement‐Lacroix et al., 2005). Finally, a similar anabolic eVect in mice is observed with a more specific and bioavailable GSK‐3 inhibitor (Clement‐Lacroix et al., 2005). Finally, neutralizing monoclonal antibodies have been successfully used to target RANKL for osteoporosis treatment (Bekker et al., 2004). It is therefore reasonable to consider that a similar approach for Wnt signaling could be developed by targeting Wnt inhibitors such as Dkks, Sfrps, WIF‐1, or sclerostin. Strong evidence of increased bone mass was reported using monoclonal neutralizing antibodies targeting sclerostin in rats (Warmington et al., 2004). Altogether, these data validate the fact that therapeutic intervention by modulating Wnt pathway results in an anabolic eVect. The true challenge


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will be to determine whether it is possible to aVect bone formation without activating the signaling events that have been reported to be associated with tumor development in the colon and other tissues.

XII. Conclusion and Perspectives Progress in the mapping of the human genome has allowed the rapid identification of the genes responsible for several genetic diseases of the skeleton. The finding of both gain‐ and loss‐of‐function mutations in the LRP5 receptor regulating in opposite manner bone formation and bone mass in humans provided an entirely novel insight on how bone cells were regulated, oVering novel opportunities for therapeutic intervention in an area of major medical need. The extraordinary burst of genetic and molecular investigations that followed have now firmly established the Wnt canonical signaling pathway as a master regulator of osteoblast diVerentiation and function in the adult skeleton. Most pharmaceutical companies involved in the prevention and treatment of osteoporosis are now engaged in the identification of compounds that could regulate this pathway in humans, thereby potentially allowing not only to prevent bone loss in aging individuals but also to restore lost bone mass in patients already suVering from an established disease. Both in academia and in industry, the major question to address in the next couple of years is to identify the components of the pathway that regulate osteoblasts versus those that aVect other cells and their potential transformation. To date, data from patients with the HBM mutation or mice in which the pathway has been specifically activated suggest that it may be possible to aVect bone without aVecting other tissues negatively.

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Eukaryotic DNA Replication in a Chromatin Context Angel P. Tabancay, Jr. and Susan L. Forsburg Molecular and Computational Biology Section University of Southern California Los Angeles, California 90089

I. Chromatin Structure Regulates Access to the DNA A. DNA Is Packaged into Chromatin B. Chromatin Remodeling C. Histone Modification D. Heterochromatin Is Transcriptionally Silent II. Replication Origins and Origin‐Binding Proteins A. Origins B. Origin Recognition Complex Binding C. Origins, Chromatin, and Transcription III. Assembly of the Prereplication Complex on Chromatin A. Proteins of the preRC B. MCMs and Chromatin IV. Origin Activation and Replication A. Kinases B. Activating the MCM Helicase and Unwinding the DNA V. Replication Elongation Through Chromatin A. Replicating Through a Chromatin Template B. Nucleosome Assembly During Replication C. Maintaining Epigenetic Marks During Replication VI. Conclusions Acknowledgments References

There has been remarkable progress in the last 20 years in defining the molecular mechanisms that regulate initiation of DNA synthesis in eukaryotic cells. Replication origins in the DNA nucleate the ordered assembly of protein factors to form a prereplication complex (preRC) that is poised for DNA synthesis. Transition of the preRC to an active initiation complex is regulated by cyclin‐dependent kinases and other signaling molecules, which promote further protein assembly and activate the mini chromosome maintenance helicase. We will review these mechanisms and describe the state of knowledge about the proteins involved. However, we will also consider an additional layer of complexity. The DNA in the cell is packaged with histone proteins into Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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chromatin. Chromatin structure provides an additional layer of heritable information with associated epigenetic modifications. Thus, we will begin by describing chromatin structure, and how the cell generally controls access to the DNA. Access to the DNA requires active chromatin remodeling, specific histone modifications, and regulated histone deposition. Studies in transcription have revealed a variety of mechanisms that regulate DNA access, and some of these are likely to be shared with DNA replication. We will briefly describe heterochromatin as a model for an epigenetically inherited chromatin state. Next, we will describe the mechanisms of replication initiation and how these are aVected by constraints of chromatin. Finally, chromatin must be reassembled with appropriate modifications following passage of the replication fork, and our third major topic will be the reassembly of chromatin and its associated epigenetic marks. Thus, in this chapter, we seek to bring together the studies of replication initiation and the studies of chromatin into a single holistic narrative. ß 2006, Elsevier Inc.

I. Chromatin Structure Regulates Access to the DNA A. DNA Is Packaged into Chromatin In order to fit into the nucleus, DNA is packaged with histones and other proteins to form a chromatin fiber (Khorasanizadeh, 2004; Luger, 2006; Luger and Hansen, 2005). The basic repeating unit of chromatin is the nucleosome core particle, which is formed when 146 bp of DNA double helix wraps around a histone octamer to form a characteristic disklike structure. Histones are small, basic proteins and the core histones H2A, H2B, H3, and H4 are highly conserved in all eukaryotes. Each nucleosomal histone octamer contains two dimers of histones H2A‐H2B, and a tetramer of (H3‐H4)2. Variant histones can replace canonical H2A and H3 in some regions; for example, CenpA, a variant histone H3, is found in the core of the centromere and is required for kinetochore formation, while H2AZ, an H2A variant, is found associated with specific regions of transcriptionally active chromatin (HenikoV and Ahmad, 2005; Jin et al., 2005; Kamakaka and Biggins, 2005). The DNA wrapped around the histones forms a 10‐nm chromatin fiber with a distinctive ‘‘beads on a string’’ morphology. This is further compacted into higher order structures and ultimately condensed during mitosis to facilitate chromosome segregation. Importantly, chromatin packaging is not uniform throughout the genome. Regions containing actively transcribed genes are more loosely packaged (euchromatin), whereas other regions are more rigidly packaged and generally transcriptionally silent (heterochromatin). Even within a single region, nucleosomal spacing may vary. The position of the nucleosome relative

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Figure 1 Chromatin structure is modulated by three activities. (A) Histone‐modifying enzymes covalently modify exposed histone tails to aVect DNA access and protein binding. (B) Chromatin remodelers change the relationship of nucleosomes to underlying DNA, for example, sliding histones from one position (outlined nucleosome) to another (filled in nucleosomes). (C) Histone chaperones perform assembly of nucleosomes de novo. DiVerent colors in the nucleosomes refer to H2A‐H2B, and H3‐H4 pairs. Me, methyl group; Ac, acetyl group.

to DNA sequence, and the nature of individual histone‐DNA interactions, all aVect access to the underlying DNA. Proteins bound to the chromatin are not limited to DNA‐binding proteins, but include proteins that recognize the histones themselves. Thus, ‘‘chromatin binding’’ is not synonymous with ‘‘DNA binding.’’ The cell must constantly access the DNA for transcription, replication, and repair functions. Thus nucleosomes are not inert and must be mobile. Their association with the DNA is subject to considerable regulation that can change function of the underlying DNA sequence. We will discuss two broad mechanisms important for modulating nucleosomes, and regulating access to the underlying DNA: chromatin remodeling and histone modifications (Fig. 1). Although we will introduce them separately, these mechanisms are often closely linked and work together to aVect chromatin structure.

B. Chromatin Remodeling Remodeling typically refers to the movement and relocalization of nucleosomes relative to DNA sequence. We will consider nucleosome assembly associated with newly synthesized DNA in Section V; here, we will discuss changes in access to existing chromatin. Remodeling is carried out by members of a large family of ATPases, each of which is typically the catalytic subunit of a large protein complex (Becker and Horz, 2002; Cairns, 2005; Haushalter and Kadonaga, 2003; Lusser and Kadonaga, 2003; Tsukiyama, 2002; Vignali et al., 2000). These ATP‐dependent remodeling complexes fall into several broad families, defined by sequences outside their ATPase domains (Becker and Horz, 2002; Cairns, 2005; Dirscherl and Krebs, 2004;

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Lusser and Kadonaga, 2003; Mohrmann and Verrijzer, 2005; Tsukiyama, 2002). These enzymes utilize the energy derived from ATP hydrolysis to change the relationship between nucleosomes and the underlying DNA (Becker and Horz, 2002; Haushalter and Kadonaga, 2003; Lusser and Kadonaga, 2003; Vignali et al., 2000). This can be done in several diVerent ways. One way is nucleosomal sliding in which the enzyme changes the nucleosome position relative to the associated DNA sequence. Another strategy is by nucleosome displacement in which the histone octamer is actually removed from the DNA. A third strategy is to tighten or loosen the association of the DNA with the nucleosome. Movement of nucleosomes can also change superhelical torsion. Lastly, remodeling enzymes can function by replacing the core histones of a nucleosome with other histones or histone variants. The eVects of these activities have been best studied in transcription, where they change the accessibility of a promoter site for sequence‐specific transcription factors and influence gene expression. However, their role in controlling DNA accessibility is likely to be important for replication as well. Four major families of remodelers are particularly important. The best‐ studied group is the SWI/SNF family (Mohrmann and Verrijzer, 2005). The catalytic subunits of this family are associated in a large protein complex with many other proteins, including an actin related protein. Biochemically, these proteins cause relatively large changes in histone orientation, including sliding over large distances and even ejection of histones. SWI/SNF proteins typically contain a bromodomain motif at the C‐terminus; this motif typically bind acetylated histones (Section I.C) (de la Cruz et al., 2005; Khorasanizadeh, 2004). This suggests that acetylation, a common histone modification that we will discuss later, is linked to the activity of this group of remodelers. The second class of remodelers, ISWI (Imitation SWItch) has been implicated in nucleosome assembly and spacing (Corona and Tamkun, 2004; Dirscherl and Krebs, 2004). It is thought to make relatively small‐scale changes of the DNA around the core nucleosome, with short‐distance sliding. This class includes diVerent related complexes with overlapping subunit membership, including ISWI and chromatin accessibility complex (CHRAC); there are several related catalytic subunits in most species. ISWI proteins interact physically with cohesion proteins and are required for replication of heterochromatin (Collins et al., 2002) (Sections IV and V). The CHD family is named for its chromodomain (de la Cruz et al., 2005; Lusser and Kadonaga, 2003). Chromodomains are typically methyl‐histone binding motifs that are found in a range of chromatin‐associated proteins with diVerent functions (Section I.C). CHD family members are often associated with histone modifiers, particularly histone acetylases and deacetylases (Pray‐Grant et al., 2005; reviewed in Lusser and Kadonaga, 2003).


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This provides another strong link between chromatin remodeling and covalent histone modification. CHD proteins are associated with transcriptional elongation and chromatin assembly in a variety of species. The final class, called the ‘‘split ATPase’’ or INO80 family, is unusual because the ATPase domain splits in half by a long interruption (Cairns, 2005; van Attikum and Gasser, 2005a). Ino80, the founding member of this family, also associates with actin related proteins, which are thought to target the protein to phosphorylated histones in regions surrounding double‐strand breaks (Tsukuda et al., 2005; van Attikum and Gasser, 2005a). SWR1, the other member of this family, has a histone exchange function, and works to ‘‘swap’’ variant histones into the nucleosomes of actively transcribed genes, in particular a special form of H2A called H2AZ (Korber and Horz, 2004; Mizuguchi et al., 2004; van Attikum and Gasser, 2005a). Any chromatin transactions that require DNA access are likely to involve some of these chromatin remodelers. Significantly, most of them were identified because of their role in transcription. Yet the same issues of access also aVect replication, recombination, and repair. Thus chromatin remodelers are likely to be involved in all reactions that require a DNA template. While elucidating their role in DNA replication apart from a role in transcription is challenging, some of them have been linked to DNA replication, and we will discuss them again later in this chapter.

C. Histone Modification The second major mechanism controlling chromatin access is histone modification (Cosgrove and Wolberger, 2005; Jenuwein and Allis, 2001; Peterson and Laniel, 2004; Santos‐Rosa and Caldas, 2005). The core histones have N‐terminal tails that protrude from the octamer. These tails, as well as accessible parts of the core octamer, are subject to covalent modification with a wide range of chemical groups including acetylation, methylation, phosphorylation, and ubiquitylation. Distinct enzymes are responsible for adding or removing these modifications on diVerent residues, and on diVerent histones. These modifications have a wide variety of eVects on heterochromatin structure, centromere assembly, transcriptional activation, and silencing, DNA repair, and chromosome segregation. The best‐studied modification is acetylation on lysine residues (Kurdistani and Grunstein, 2003; Sterner and Berger, 2000; Verdone et al., 2005). Histone acetyltransferases (HATs) fall into several distinct families that share sequence similarity but recognize diVerent histone residues. Many of them are associated with transcriptional activation, and were identified because they stimulated transcription activity or act as coactivators. However, like

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the chromatin remodelers, it is now recognized that their eVects extend well beyond transcription. Acetylation is a reversible modification, with relatively fast kinetics. It is reversed by histone deacetylases (HDACs) which can be categorized into several diVerent families diVering by structure and specificity. Like the HATs, the HDACs typically target specific histone residues so that some of these enzymes may directly antagonize each other. Methylation of histones is also a significant modification, although it does not undergo the same rapid global turnover as acetylation (Lachner et al., 2003; Martin and Zhang, 2005; Peters and Schubeler, 2005; Shilatifard, 2006). There are a number of histone methyltransferases (HMTs), again varying with the residue modified and with the number of methyl groups added. Demethylating activities have been harder to identify, but several distinct activities have now been found (Holbert and Marmorstein, 2005; Kubicek and Jenuwein, 2004; Tsukada et al., 2006). Importantly, the same lysine residue in a histone may be subject to acetylation or methylation, but not at the same time. Thus, the modifying enzymes may antagonize one another. Several additional covalent modifications are also observed in histones. Phosphorylation occurs on diVerent histones in response to various signals. For example, residue S10 of H3 (H3S10) is phosphorylated during mitosis throughout the genome and provides a robust marker of mitotic progression (Fry et al., 2004; Prigent and Dimitrov, 2003). On the other hand, phosphorylation of the H2A variant H2AX by checkpoint kinases ATM and ATR occurs only in the region of double‐stranded breaks in the DNA, and helps localize the repair and checkpoint response (Fernandez‐Capetillo et al., 2004; Foster and Downs, 2005; Thiriet and Hayes, 2005). Monoubiquitylation on the C‐termini of H2A or H2B has been associated with gene activation and inactivation (Gill, 2004; Horn and Peterson, 2006; Zhang, 2003), and the related SUMO molecule may also modify histones to facilitate repression (Shiio and Eisenman, 2003; Xhemalce et al., 2004). Perhaps the pattern of histone modifications is not endlessly variable but diVerent modifications may be coupled to one another. For example, there is some evidence that some HAT enzymes may be associated with a deubiquitinylating enzyme, linking these modifications and allowing one to act as a target for another (Czermin and Imhof, 2003; Fischle et al., 2003). At first, acetylation was associated exclusively with transcriptional activation, and the hypothesis was that the acetylated histones ‘‘loosened’’ the DNA on the nucleosome making the DNA more accessible or the nucleosome more mobile (Cosgrove and Wolberger, 2005). A new study shows that acetylation on H4K16 prevents compaction of the chromatin fiber (Shogren‐Knaak et al., 2006), which would be consistent with a more open state required for RNA polymerase access; however, with the discovery of other acetylated residues associated with transcriptionally inactive regions, obviously the story is more complicated. Now, it appears that histone tail


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modifications not only have intrinsic eVects on chromatin structure but also provide binding sites for a variety of chromatin proteins (Cosgrove and Wolberger, 2005; de la Cruz et al., 2005). Distinct protein motifs recognize diVerent modifications, and provide a mechanism to recruit a variety of binding factors. Not surprisingly, some of these binding domains are not only found on chromatin remodelers, which provides one way to coordinate remodeling and modification, but they are also found on other proteins related to a wide range of DNA and chromatin activities. For example, chromodomains bind to methylated histones, and are found in the CHD class of chromatin remodelers as well as chromatin‐binding proteins such as heterochromatin protein 1 (HP1, or Swi6 in fission yeast; Brehm et al., 2004). Over the last decade, it has been suggested that the pattern of histone modifications and their associated proteins presents the cell with an additional level of heritable information: an epigenetic histone code that is independent of the underlying DNA sequence (Cosgrove and Wolberger, 2005; Jenuwein and Allis, 2001; Peterson and Laniel, 2004). Particular patterns of modification are associated with diVerent genome characteristics. For example, actively transcribed genes are likely to be associated with chromatin that is acetylated on certain residues of histones H3 and H4, ubiquitylated on H2B, and methylated on K4 of H3. Such a code may certainly impact the replication process (Sections II and III) and, because the code can be inherited, there must exist a mechanism to recreate it on newly synthesized DNA as it is assembled into chromatin (Section V).

D. Heterochromatin Is Transcriptionally Silent The code established by heterochromatin provides a particularly good model to explain the basics of epigenetic inheritence (Allshire, 2004; Grewal and Elgin, 2002; Maison and Almouzni, 2004; Martienssen et al., 2005; Wallace and Orr‐Weaver, 2005). Heterochromatin is typically rigid and compact with regularly spaced nucleosomes and is less accessible than most euchromatin. Constitutive heterochromatin includes permanent structural domains such as centromeres and telomeres, while facultative heterochromatin occurs in regions that are only transiently silenced. Heterochromatic silencing provides a capacity to silence large regions of the chromosome, rather than repress expression on a gene‐by‐gene basis. Here, we will briefly describe heterochromatin (Fig. 2). Its replication will be discussed in Section V. In most eukaryotes, transcriptionally inactive heterochromatin is typically underacetylated, methylated on K9 of H3, and bound by the chromatin‐ binding protein HP1/Swi6 (Allshire, 2004; Eissenberg and Elgin, 2000; Martienssen et al., 2005; Wallace and Orr‐Weaver, 2005). We will consider

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Figure 2 Heterochromatin structure relies on binding of HP1/Swi6 to methylated histones. This can recruit the histone methyltransferase (HMT) to methylate neighboring histones and spread the signal.

a ‘‘generic’’ HP1, although there are several isoforms in metazoa with distinct functions (e.g., Hediger and Gasser, 2006; Lomberk et al., 2006). HP1/Swi6 is mediated by association between its chromodomain and methylated H3K9. Adjacent HP1 molecules are postulated to form a more compact and structurally rigid form of chromatin that is particularly important for centromere and telomere function. There is positive feedback associated with HP1, because binding of HP1 can recruit the HMT, Su(var)3–9/Clr4, to methylate adjacent histones. This in turn recruits additional HP1/Swi6. This spreads the HP1‐associated structure into adjacent regions along the chromatin fiber (Fig. 2); how the cell limits this spreading to prevent heterochromatization of the entire genome remains an important question. Establishment of this mark de novo requires a number of conserved proteins that apparently function independently, including the enzymes of the RNAi machinery (Almeida and Allshire, 2005; Bailis and Forsburg, 2002; Martienssen et al., 2005) as well as histone deacetylation (Freeman‐ Cook et al., 2005; Horn and Peterson, 2006; Jia et al., 2005; Shankaranarayana et al., 2003; Yamada et al., 2005), ubiquitylation and methylation (Bannister et al., 2001; Horn and Peterson, 2006; Horn et al., 2005; Nakayama et al., 2001b). Phosphorylation of H3S10 during mitosis is incompatible with HP1 binding to the adjacent H3K9Me residue (Fischle et al., 2005; Hirota et al., 2005), and may provide the cell with means to periodically disrupt HP1 and remodel the heterochromatin, perhaps during S phase. Although HP1 clearly has additional, more subtle eVects not limited to silencing, to a first approximation, its binding to methylated histone H3K9 can be used to define heterochromatin (reviewed in Hediger and Gasser, 2006; Maison and Almouzni, 2004; but see Gilbert et al., 2003). This mechanism is conserved in most eukaryotes, with the striking exception of Saccharomyces cerevisiae, which lacks HP1 and assembles silent chromatin by a diVerent mechanism involving Sir1–4 (Pillus and Rine, 2004; Rusche et al., 2003). However, the Sir1 protein in S. cerevisiae, while its sequence is unique to that species, plays a role not unlike that of HP1/Swi6 (Section V.B).


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Heterochromatin proteins and associated modifications have many tantalizing links to DNA replication proteins. We will discuss silencing and heterochromatin again in the following section. But first, we must consider the chromatin context in which DNA synthesis initiates at the origin of replication.

II. Replication Origins and Origin‐Binding Proteins A. Origins DNA replication in eukaryotic cells begins at replication origins (Bell and Dutta, 2002; DiZey, 2004; Forsburg, 2004; Gerbi et al., 2002). Origins of replication are distributed widely throughout a typical eukaryotic genome to ensure timely replication of DNA. However, origin structure varies considerably from species to species (Bell, 2002; Bielinsky and Gerbi, 2001; Cvetic and Walter, 2005; Gilbert, 2001a; Robinson and Bell, 2005). In the well‐ studied budding yeast S. cerevisiae, origins are determinate, and associated with specific DNA elements of about 100 base pairs, which includes a defined consensus sequence that is essential for full activity (Bell, 2002; Cvetic and Walter, 2005; Robinson and Bell, 2005; Weinreich et al., 2004). These origins fire in an orderly and reproducible pattern that ensures the timely duplication of the genome (Lengronne et al., 2001; Raghuraman et al., 2001; Yabuki et al., 2002). In contrast, in the fission yeast Schizosaccharomyces pombe, origin elements are A/T‐rich regions of about a kilobase, with asymmetric strand composition (Antequera, 2004; Dai et al., 2005). They lack a defined sequence consensus and are degenerate in that multiple mutations are required to eliminate their function (Clyne and Kelly, 1995; Dubey et al., 1996; Kim et al., 2001; Okuno et al., 1997). Fission yeast origins often cluster in groups in which only one origin is likely to be used per cell cycle (Dubey et al., 1994; Patel et al., 2005; Sivakumar et al., 2004). They are also relatively ineYcient compared to origins in S. cerevisiae, meaning that a given origin fires in only a fraction of the cell cycles (Dubey et al., 1994; Gomez and Antequera, 1999; Patel et al., 2005; Segurado et al., 2002). In metazoans, a number of specific origins have been identified that result in specific replication initiation zones (Bell, 2002; Bielinsky and Gerbi, 2001; Cvetic and Walter, 2005; DePamphilis, 1999; Gilbert, 2001a; Robinson and Bell, 2005; Todorovic et al., 1999). However, in contrast to budding yeast, binding of replication proteins to these origins is not sequence specific (see in a later section; Kong et al., 2003; Remus et al., 2004; Vashee et al., 2003) and mutational analysis suggests that, as in fission yeast, crucial elements are dispersed and redundant (Aladjem and Fanning, 2004; Cvetic and Walter,

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2005; Gilbert, 2004; Kohzaki and Murakami, 2005). The extreme example is in embryonic systems such as Xenopus oocytes in which origins appear to be stochastically defined, perhaps by a spacing mechanism (Blow et al., 2001; Hyrien et al., 1995; Lucas et al., 2000; reviewed in Cvetic and Walter, 2005; Gilbert, 2001a; Robinson and Bell, 2005). Remarkably, these cells can eYciently replicate exogenous DNA in a cell cycle dependent fashion (Harland and Laskey, 1980; Mechali and Kearsey, 1984). Thus, there must be mechanisms independent of DNA sequence that can define origins and determine their activity. While embryonic origins are randomly distributed, zygotic origins are more defined (Bielinsky, 2003; Cvetic and Walter, 2005; Gilbert, 2004) so that the process of development appears linked to a more defined use of origin regions. Studies primarily in budding yeast but also in fission yeast show that there is substantial variation between origins. One source of variation is eYciency. Weak or ineYcient origins are unlikely to fire unless replication is delayed, while strong origins will fire in most S phases. The determinants that make an origin weaker are not fully understood. A better understood variable is timing. Elegant studies in budding yeast show that some origins reliably fire early whereas others routinely fire late during S phase (Ferguson et al., 1991; Friedman et al., 1995; Raghuraman et al., 1997, 2001). Such a distribution may be important to distribute replication factors, which may be limiting. In other species as well, replication timing is neither uniform throughout S phase (Gilbert, 2002a,b; Goren and Cedar, 2003; Kim and Huberman, 2001) nor throughout the nucleus (Gilbert, 2001b). There is evidence that in some cases, but not all, this late firing may be determined by additional cis‐acting sequences (Friedman et al., 1996; Yompakdee and Huberman, 2004). In particular, regions of heterochromatin are generally late replicating (reviewed in Gilbert, 2002b; Goren and Cedar, 2003; but see Kim et al., 2003; Weidtkamp‐Peters et al., 2006), while transcriptionally active regions often replicate early (Holmquist, 1987; Selig et al., 1992). Consistent with this, an active silencer in budding yeast can delay origin firing (Zappulla et al., 2002). This has led to considerable speculation about links between replication timing and transcription (Gilbert, 2002b; Kohzaki and Murakami, 2005; Schwaiger and Schubeler, 2006), some of which we will consider in the following section.

B. Origin Recognition Complex Binding Remarkably, despite the significant diVerences in origin sequence and structure, all eukaryotes use the same broad palette of proteins to initiate DNA replication (Bell and Dutta, 2002; DiZey, 2004; Forsburg, 2004; Gerbi et al., 2002). The crucial complex that defines the origin is called origin recognition complex, ORC, which is bound to the origin throughout the cell cycle


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(Fig. 3A) (Bell, 2002; Bell and Stillman, 1992; DiZey and Cocker, 1992; Spradling, 1999). ORC is conserved in eukaryotes and in archaea (Bell, 2002; Kelman and Hurwitz, 2003; Tye, 2000) and comprises six subunits. In S. cerevisiae, ORC is a sequence‐specific binding protein that recognizes the core sequence of the origin (Bell, 2002). In contrast, S. pombe Orc4 directs the complex to a type of sequence, rather than a conserved sequence element. It contains a distinct ‘‘A/T hook’’ motif that perhaps anchors it on the large A/T‐rich origin (Chuang and Kelly, 1999; Kong and DePamphilis, 2001). ORC from Drosophila, Xenopus, or humans shows no sequence specificity (Kong et al., 2003; Remus et al., 2004; Vashee et al., 2003), indicating that other methods are required to recruit it to particular initiation zones. If ORC is tethered to an ectopic region via a nonrelated DNA‐binding domain, it confers origin activity (Takeda et al., 2005): thus, its binding is suYcient to define an origin. It is useful to think of ORC as a marker of potential origins of replication. However, ORC localization is not restricted to active replication origins but also found in silencers and unused origins, so not all its sites will be used as origins; it may serve a broader function as a landing platform for a range of chromosomal proteins (Dillin and Rine, 1997; Fox et al., 1995; Rusche et al., 2002; Wyrick et al., 2001). Intriguingly, ORC is required to assemble silenced chromatin in S. cerevisiae, and there exist separation of function alleles of ORC that distinguish silencing from replication (Bose et al., 2004; Dillin and Rine, 1997; Foss et al., 1993, 1995; Suter et al., 2004). Consistent with this, mutants of Drosophila Orc2 have a Su(var) phenotype, meaning they suppress the variegation of gene expression associated with heterochromatin boundaries. In Drosophila, Xenopus, and humans, ORC interacts with HP1 (Huang et al., 1998; Lidonnici et al., 2004; Pak et al., 1997; Prasanth et al., 2004); we have observed similar results in fission yeast (J. M. Bailis and S. L. Forsburg, unpublished data). Considerable evidence now links HP1 and ORC in heterochromatin function, although the mechanism remains elusive (Section V.B). This provides our first link between replication and chromatin structure.

C. Origins, Chromatin, and Transcription Studies in transcription suggest a variety of ways that sequence‐specific transcription factors, histone modifiers, and chromatin remodelers work together to facilitate access to the DNA (Khorasanizadeh, 2004; Luger, 2006; Mellor, 2005; Shilatifard, 2006). This provides a paradigm for the interplay of chromatin structure and nucleic acid transactions. Generally, accessible and transcriptionally active chromatin is hyperacetylated on histones H3 and H4, and methylated on the K4 residue of histone H3.

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Figure 3 A model showing how ordered assembly of proteins leads to replication origin firing. (A) Replication origins are marked by ORC binding. This may be modulated by histone acetylation of chromatin (HAT? and Hbo1?) or by HAT association with ORC (Hbo1?).


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The nucleosomes are actively relocalized or remodeled. The order of these events varies from promoter to promoter, depending on its spatial and temporal context. By linking replication to transcription, the cell may co‐ opt the transcriptional program to open up the chromatin for DNA synthesis, possibly linking timing of these events (Gilbert, 2002b; Kohzaki and Murakami, 2005; Schwaiger and Schubeler, 2006). This would be consistent with changes in origin timing and usage that accompany developmental transitions in Drosophila and mammals (Antequera, 2004; Cvetic and Walter, 2005; Gilbert, 2002b). A genomic study in budding yeast suggests that promoters typically have well‐positioned nucleosomes flanking a short (200 bp) region that is nucleosome free (Bernstein et al., 2004; Yuan et al., 2005). This could suggest an entry point for transcription factors that can then modulate positioning of the remaining nucleosomes. In budding yeast, links between transcription and replication are long established. Origins are typically intergenic in S. cerevisiae (Raghuraman et al., 2001; Wyrick et al., 2001) and contain transcription factor binding sites. For example, the transcription factor Abf1 binds at sites at some replication origins and is required for activity (DiZey and Stillman, 1989; Marahrens and Stillman, 1992), as does the Mcm1 transcription factor (Chang et al., 2004). Origins are also located in promoters in S. pombe (Antequera, 2004; Antequera and Bird, 1999; Dai et al., 2005) although their activity is independent of transcription (Gomez and Antequera, 1999; Segurado et al., 2003). A positive link to replication has also been linked to a variety of transcription factors in diVerent systems (Cheng and Kelly, 1989; Cheng et al., 1992; Danis et al., 2004; Kohzaki et al., 1999; Lemaitre et al., 2005; Li et al., 1998; Maiorano et al., 2005). This is particularly apparent in viral systems, where binding of transcription factors is thought to facilitate nucleosome mobilization around the origin (Cheng and Kelly, 1989; Cheng et al., 1992; Kohzaki and Murakami, 2005; Murakami and Ito, 1999; Stedman et al., 2004). Importantly, the data suggest that this positive eVect is independent of RNA synthesis, instead reflecting chromatin access or nucleosome organization (Cheng and Kelly, 1989; Cheng et al., 1992). In fact, active transcription inhibits origin activity in S. cerevisiae (Nieduszynski et al., 2005) and (B) ORC is activated by Cdc6 and Cdt1 association. (C) MCM complexes load via activated Cdc6 and spread along the DNA. This may be regulated by binding of Hbo1 (Hbo1?). The complex of ORC, Cdc6, Cdt1, and MCMs is the prereplication complex (preRC). (D) Mcm10 binds. DDK phosphorylates MCM. CDK phosphorylates proteins in the TopBP1/Dpb11/Rad4 complex resulting in its activation. CDK phosphorylation of Cdc6 results in Cdc6 proteolysis. Cdt1 is inactivated by geminin binding. (E) Cdc45 and GINS bind, forming the initiation complex (IC). (F) Unwinding exposes single‐stranded DNA, allowing binding of RPA and recruitment of polymerases. The leading strand polymerase is locked in place by the PCNA sliding clamp. Excess MCMs are removed from the chromatin as the forks pass. Adapted from Forsburg, 2004.

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hamster cells (Saha et al., 2004) and causes stalling of the replication fork (Prado and Aguilera, 2005). The logical inference is that it is not transcription per se but chromatin access that may provide the positive link between replication and transcription. Consistent with this potential eVect on chromatin structure, a number of studies link origin activation to nucleosomal organization in the origin region. In yeast, sequences encompassing the core origin sequence elements are void of nucleosomes but regions neighboring the core sequence have positioned nucleosomes (Ravindra et al., 1999; Thoma et al., 1984; Venditti et al., 1994), similar to what is seen in many promoters (Bernstein et al., 2004; Yuan et al., 2005). Introducing nucleosomes inappropriately into the ARS1 origin sequences inhibits origin activity (Simpson, 1990) and a transcriptional silencer delays origin firing (Zappulla et al., 2002). Thus, increasing chromatin structure in the origin region occludes origin activity. There is good evidence that S. cerevisiae ORC together with the Abf1 transcription factor act to position nucleosomes at the origin in a suitable arrangement for origin activation (Lipford and Bell, 2001). If this activity is blocked, ORC is still able to bind to the origin, but lacks the ability to recruit downstream factors (Lipford and Bell, 2001). This is consistent with studies in both Drosophila and Xenopus showing that chromatin restructuring occurs in parallel to origin of replication utilization (Blumenthal et al., 1974; Hyrien et al., 1995). In higher eukaryotes, a number of chromatin remodeling events coincide with the initiation of DNA replication (Alexiadis et al., 1997; Iizuka and Stillman, 1999; Stedman et al., 2004; Zhou et al., 2005). Importantly, specialized chromatin structure during origin assembly is distinct from that at initiation (Bell et al., 1995; Dillin and Rine, 1997; Ehrenhofer‐Murray et al., 1995; Fox et al., 1995; Palacios DeBeer et al., 2003). This suggests that in replication, as in transcription, entry of one binding factor may be required first to position nucleosomes for additional protein loading. Nucleosomal positioning at origins is also aVected by the related chromatin remodeling complexes, ISWI and CHRAC (Alexiadis et al., 1998; Langst and Becker, 2001). Intriguingly, ISWI is enriched at regions of pericentromeric heterochromatin that are actively replicated (Collins et al., 2002). Its depletion results in decreased replication of heterochromatin regions and delayed S phase progression. This suggests that ISWI is able to remodel the chromatin structure to allow either origin assembly or replication fork progression through particular compact regions. Likewise, localization of SNF2h in humans (an ISWI related protein) is enhanced at origin‐associated sequences during G1/S arrest, and depletion of SNF2h reduces origin firing (Zhou et al., 2005). In budding yeast, an origin‐specific role for the SWI/SNF remodeling factor has also been identified (Flanagan and Peterson, 1999). In a minichromosome, some origins fire eYciently in the absence of SWI/SNF but others do not, suggesting an origin‐specific sensitivity


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to chromatin context mediated by this remodeler. In the genome, swi/snf mutant cells may accommodate this eVect by relying on adjacent origins to activate replication through a refractory region. An important question is whether this putative replication‐associated remodeling acts upstream or downstream of ORC binding. Regulation by chromatin remodeling suggests that origins may be associated with ‘‘active chromatin’’ states, which immediately suggests acetylation (Weinreich et al., 2004). In budding yeast, timing of origin firing is aVected by histone acetylation because loss of the HDAC Rpd3 leads to accelerated origin firing, at least of late origins (Aparicio et al., 2004; Vogelauer et al., 2002). Loss of a diVerent HDAC, Sir2, genetically suppresses temperature sensitive mutations in a variety of origin‐binding proteins, which suggests that the origins are more eYcient or more accessible to attenuated proteins when acetylation is increased (Pappas et al., 2004). Mutants in sir2 also increase origin firing in rDNA (Pasero et al., 2002). If loss of an HDAC increases origin firing, presumably by leading to increased levels of histone acetylation, then increasing HDAC activity should reduce origin firing by reducing acetylation. Consistent with this, targeting the Drosophila Rpd3 HDAC close to an origin reduces origin firing (Aggarwal and Calvi, 2004). The eVect is not limited to manipulating HDAC activity. An increase in origin activity is observed if the Gcn5 HAT is tethered next to an origin in yeast (Vogelauer et al., 2002), again consistent with acetylation being more permissive for replication. Tethering of the PCAF or Gcn5 HATs adjacent to the polyoma virus replication origin likewise stimulates replication in animal cells (Xie et al., 2002), and acetylation corresponds with assembly of ORC and associated factors at origins of Kaposi’s sarcoma‐associated herpesvirus (Stedman et al., 2004). Increased HAT activity correlates with the onset of DNA replication in rat liver cells (Weiss and Puschendorf, 1988) and histone acetylation facilitates replication elongation in vitro (Alexiadis et al., 1997). Histone acetylation is also associated with origin activation in Drosophila (Aggarwal and Calvi, 2004) and Xenopus (Danis et al., 2004). Together, these results suggest that histone acetylation is more permissive for DNA replication. This is consistent with the study suggesting that histone acetylation promotes transcription by preventing chromatin compaction (Shogren‐Knaak et al., 2006), and suggests that access to replication origins may be aVected by many of the same constraints that regulate promoter access during transcription. However, while histone acetylation may be helpful, is it essential? The evidence here is less clear. While the experiments above all suggest that the acetylation state of the origins may influence their accessibility, most of the HATs and HDACs in yeast are not essential for cell viability or DNA replication (Kurdistani and Grunstein, 2003; Sterner and Berger, 2000). Their eVects, described earlier, might be thought of as fine‐tuning, rather than an essential activity. Alternatively, there might be considerable overlap

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in the eVects of the histone modifications that prevent the cell from being completely dependent on any single HAT or HDAC. Recent data identify a candidate HAT involved in replication, although its mechanism is not clear. Human Orc1 interacts with an MOZ, Ybf2/Sas3, Sas2, and Tip60 (MYST) family HAT, called Hbo1 (Fig. 3A) (Burke et al., 2001; Iizuka and Stillman, 1999). An obvious model suggests that the HAT may directly aVect replication origin access, perhaps being localized by ORC assembly to the chromatin to facilitate assembly of other proteins. But, other data disagree: temperature sensitive orc mutants in S. cerevisiae are suppressed by loss of a MYST family HAT, which provides genetic evidence that an HAT inhibits ORC activity (Dillin and Rine, 1997; Ehrenhofer‐ Murray et al., 1997; Foss et al., 1993, 1995). It is also important to remember that HATs are not limited in activity to histone substrates (Chen et al., 2001; Hasan and Hottiger, 2002). One study suggests that Hbo1 can actively acetylate ORC and associated proteins (Iizuka et al., 2006). Thus, involvement of a HAT in replication does not prove that histone acetylation is the relevant target: the replication proteins themselves may also be targets for modification. We will discuss additional evidence in support of this theory in the following section, where we describe additional replication factors.

III. Assembly of the Prereplication Complex on Chromatin The prereplication complex (preRC) which assembles at the replication origin contains several multiprotein complexes that render the origin competent for activation (Bell and Dutta, 2002; DiZey, 2004; Forsburg, 2004; Gerbi et al., 2002; Johnson and O’Donnell, 2005). The preRC is poised to fire, but is not able to initiate DNA synthesis without additional protein factors. Assembly of the preRC is tightly coupled to the cell cycle clock established by cyclin‐dependent kinases (CDKs); it occurs at the end of mitosis, when CDK levels start to fall. Increased CDK activity prevents further preRC assembly, restricting it to that part of the cycle where B‐type cyclins are inactive (Baum et al., 1998; Coue et al., 1996; Kearsey et al., 2000; Mendez and Stillman, 2000; Su and O’Farrell, 1997; Tanaka et al., 1997).

A. Proteins of the preRC 1. Cdc6 and Cdt1 At the replication origin, ORC is bound by Cdc6 (an ATPase) and Cdt1 during G1 phase firing (Fig. 3B; see Table I for nomenclature in diVerent species) (Bell and Dutta, 2002; DiZey, 2004; Forsburg, 2004; Gerbi et al., 2002; Johnson and

Table I Synonyms for Conserved Replication Proteins Protein

S. cerevisiae

Cdc6 Cdc7 Cdc45 Ctf4 Cdt1 Dbf4 Dpb11 Geminin GINS: PSF2 PSF3 SLD5 PSF1 HP1 Mcm2 Mcm3 Mcm4

Cdc6 Cdc7 Cdc45 Ctf4, Chl15, Pob1 Tah11, Sid2 Dbf4, Dna52 Dpb11 No homologue

Ddc18 Hsk1 Sna41, Cdc45 Mcl1 Cdt1 Dfp1, Him1 Rad4, Cut5 No homologue

Psf2, Cdc102 Psf3 Sld5, Cdc105 Psf1, Cdc101 No homologue Mcm2 Mcm3 Mcm4, Cdc54

Psf2, Bsh3 Psf3 Sld5 Psf1 Swi6 Mcm2, Cdc19, Nda1 Mcm3 Mcm4, Cdc21

Mcm5 Mcm6 Mcm7 Mcm8 Mcm10 ORC1–6

Mcm5, Cdc46 Mcm6 Mcm7, Cdc47 No homologue Mcm10, Dna43 Orc1–6

Mcm5, Nda4 Mcm6, Mis5 Mcm7 No homologue Cdc23, Mcm10 Orp1–6, Orc1–6

?, not yet identified.

S. pombe

Drosophila

Human

Xenopus

Cdc6 Cdc7 Cdc45 ? Dup (double‐parked) ChiVon Mus101 Geminin

Cdc6 Cdc7 Cdc45 AND‐1 Cdt1 Dbf4, ASK TopBP1, Cut5 Geminin

? ? ? ? Su(var)2–5 DmMcm2 DmMcm3 DmMcm4 Dpa (disk proliferation abnormal) DmMcm5, DmCdc46 DmMcm6 DmMcm7 ? Mcm10 DmORC1–6

Psf2, Bsh3 ? ? ? HP1, , BM28, Mcm2 Mcm3 Mcm4

Cdc6 Cdc7 Cdc45 ? Cdt1 Dbf4 Xmus101 Geminin GINS: Psf2 Psf3 Sld5 Psf1 HP1 Mcm2 Mcm3 Mcm4

Mcm5 Mcm6 Mcm7 Mcm8 Mcm10 ORC1–6

Mcm5 Mcm6 Mcm7 ? Mcm10 XOrc1–6

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O’Donnell, 2005). Cycles of ATP hydrolysis by ORC and Cdc6 promote the loading of multiple mini chromosome maintenance (MCM) complexes, suggesting that Cdc6 may act as a clamp loader to assemble the ring‐shaped MCM complex on the DNA (see the following section) (Davey et al., 2002; Fujita et al., 1999; Johnson and O’Donnell, 2005; Lee and Bell, 2000; Mendez and Stillman, 2003; Perkins and DiZey, 1998; Randell et al., 2006; Speck et al., 2005; Weinreich et al., 1999). The molecular contribution of Cdt1 to this activity is unclear. Cdc6 is regulated by multiple mechanisms so that its activity is restricted to the G1/S phase of the cell cycle. Activation of the replication origin by CDK (see later) leads to phosphorylation of Cdc6 and its protein degradation or nuclear export (Delmolino et al., 2001; Jallepalli et al., 1997; Jiang et al., 1999; Mailand and DiZey, 2005; Mimura et al., 2004; Nguyen et al., 2001). Cdt1 is also regulated. In metazoans, a small protein called geminin binds Cdt1 and prevents its activity in G2 or mitosis (Arias and Walter, 2005; Tada et al., 2001; Wohlschlegel et al., 2000; Yanagi et al., 2002). 2. The MCM Helicase The MCM complex is a heterohexameric ring of six related proteins called Mcm2–7 (Forsburg, 2004; Maiorano et al., 2006). The completed assembly of ORC, Cdc6, Cdt1, and MCM forms the preRC, marking the origin as competent for initiation (Fig. 3C). Although biochemical experiments suggest that ORC and Cdc6 are dispensable once MCMs are bound (Donovan et al., 1997; Hua and Newport, 1998), in vivo experiments in the yeasts suggest that continued expression of Cdc6 throughout G1 is required to maintain the preRCs (Baum et al., 1998; Cocker et al., 1996; Piatti et al., 1995; Santocanale and DiZey, 1996). MCM proteins share a core sequence with homology to the AAA class of ATPases, which includes specific sequence motifs that define the MCM family (Cvetic and Walter, 2006; Forsburg, 2004; Maiorano et al., 2006). Outside of the core motif, MCMs are distinct from one another so that (for example) the Mcm2 from yeast and humans is more closely related than the Mcm2 and Mcm3 from yeast. MCMs are very ancient, and are also found in archaeal species, where there is typically only one family member that forms a homohexamer (Kearsey and Labib, 1998; Kelman and White, 2005). The hexameric ring structure is highly suggestive of a replicative DNA helicase (Forsburg, 2004; Patel and Picha, 2000; Sclafani et al., 2004), and as will be discussed later, the MCM complex is a compelling candidate for this role. However, at the preRC, the MCM complex fulfills the role of replication assembly factor. MCMs in most organisms are constitutively located in the nucleus throughout the cell cycle, but binding to the chromatin is cell cycle regulated.


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MCMs only bind chromatin during G1/S phase and are dislodged from the chromatin as S phase proceeds (Forsburg, 2004). Although there is always a soluble fraction of MCM protein in the nucleus throughout the cell cycle, access of the MCMs to chromatin via Cdc6 and Cdt1 is a key regulatory event necessary to control origin usage. The exception is S. cereivsiae in which MCMs cycle in and out of the nucleus during the cell cycle (Hennessy et al., 1990; Nguyen et al., 2000; Yan et al., 1993); their nuclear localization reflects chromatin association in other species. Cytological studies in metazoa indicate that MCMs are liberally distributed throughout the nucleus, associated with unreplicated chromatin, and are not concentrated at the origin (Dimitrova et al., 1999; Krude et al., 1996; Madine et al., 1995). This excess of MCMs distant from the origin (termed the ‘‘MCM paradox’’) (Hyrien et al., 2003; Laskey and Madine, 2003) may suggest that MCMs load at the preRC and spread along the DNA (Hyrien et al., 2003; Laskey and Madine, 2003), perhaps similarly to a translocating DnaB helicase in bacteria (Kaplan and O’Donnell, 2002; Kaplan et al., 2003b). Several experiments suggest that there are multiple MCM complexes associated with a single origin (Bowers et al., 2004; Edwards et al., 2002; Ogawa et al., 1999; Ritzi et al., 1998). Genome‐wide analysis using ‘‘ChIP on a chip’’ (chromatin immunoprecipitation and PCR, followed by microarray analysis of the products) suggests that there are 600–700 MCM sites per S. cerevisiae genome, of which approximately 10% do not colocalize with ORC (Wyrick et al., 2001). This is slightly more than the anticipated number of origins (Raghuraman et al., 2001; Wyrick et al., 2001), but substantially less than the 30–50,000 estimated MCM molecules per yeast cell (Donovan et al., 1997; Lei et al., 1996). Even given the possibility that a significant fraction of the MCMs remains soluble in the nucleus, it would appear that this method underestimates the MCMs on the chromatin. This suggests that the more distal MCMs are distributed randomly in the regions adjacent to the origin, although it is a formal possibility that they form larger order structures or have weaker associations. The function of these distal MCMs is unclear, and has been subject of considerable speculation. Among the possibilities: they could contribute to genome stability, facilitate replication through structurally complex regions, or ensure proper spacing of origins. The ratio of MCMs bound to ORC at the origin was examined using a Xenopus in vitro system in which origin sequences are not defined by sequence but determined stochastically (Edwards et al., 2002). On an 80 bp fragment of DNA, ORC and MCM assemble at a 1:1 ratio. However, as the fragment length was increased, the amount of MCM (but not of ORC) also increased, approaching the estimate of 20–40 MCMs per ORC observed for sperm chromatin in Xenopus extracts (Edwards et al., 2002; Walter and Newport, 1997). A number of investigators have suggested that the spreading MCMs

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contribute to the choice of initiation sites which may facilitate optimal spacing of potential origins (Blow, 2001; Edwards et al., 2002; Hyrien et al., 2003). This suggests that only a subset of the MCMs is an active participant in DNA synthesis. Curiously, cells are sensitive even to modest reductions of MCMs, suggesting that these abundant proteins may have functions beyond DNA replication (Lei et al., 1996; Liang et al., 1999). On the basis of extensive physical and genetic interactions with other factors, MCM proteins have been implicated in diverse chromosome transactions including genome stability, transcription, and chromatin modification (Bailis and Forsburg, 2004; Forsburg, 2004). Intriguingly, mouse Mcm2 binds histone H3‐H4, and although it is not known whether this depends on particular histone modifications, the binding is suYciently robust that it can be used to aYnity purify the MCM complex (Holland et al., 2002; Ishimi et al., 1996). However, MCM association with chromatin is disrupted as S phase proceeds, so any additional functions are likely to be linked to S phase progression or maintenance of genome stability specifically during S phase (Bailis and Forsburg, 2004; Forsburg, 2004).

B. MCMs and Chromatin As discussed earlier, studies in S. cerevisiae show that ORC and the associated factor Abf1 are required to position nucleosomes at the origin (Lipford and Bell, 2001). Failure to carry out this remodeling does not disrupt ORC binding, but prevents Cdc6 and MCM association with the origin, suggesting that ORC needs to adjust nucleosome spacing to allow MCMs access to the DNA. Several additional intriguing links couple MCM activity to chromatin modification. First, a study in budding yeast suggests that mutations in the HDAC Sir2 can suppress hypomorphic mutations in Cdc6 or MCM (Pappas et al., 2004), which suggests that chromatin acetylation (which would increase in the absence of an HDAC) may facilitate access of preRC proteins to DNA. This model is consistent with a second observation: overexpression of a member of the Gcn5‐containing HAT complex partially suppresses the temperature sensitive phenotypes of an mcm5 mutant (Dziak et al., 2003). This could reflect increased acetylation of an origin or associated proteins. It could also suggest direct interactions between HATs and MCMs. A third observation from humans, Drosophila, and Xenopus shows that the Hbo1 HAT protein, which we discussed earlier in Section II.C, binds MCM as well as ORC (Burke et al., 2001; Iizuka and Stillman, 1999; Stedman et al., 2004). Mutational analysis of the interaction between the Drosophila proteins, using a two‐hybrid assay, localized the interaction to the zinc‐finger domain of Hbo1 and the N‐terminus of Mcm2 (Burke et al., 2001). Disruption of


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Hbo1 by siRNA or antisense methods inhibits viral replication (Stedman et al., 2004) and MCM chromatin association in humans (Iizuka et al., 2006). Depletion of this HAT from Xenopus extracts disrupts MCM binding (Iizuka et al., 2006), although it does not seriously deplete the amount of MCMs, and nor does it aVect ORC or Cdc6 binding. This defect in MCM loading can be rescued by addition of Cdt1, which would suggest that the depletion of Hbo1 removes or inhibits active Cdt1, which is required for MCM loading. Does Hbo1 in some way couple Cdt1 activity to MCMs by binding both proteins? The mechanism remains unclear. It is also not clear whether this eVect is mediated through histones or through direct interaction with the preRC proteins. Curiously, Hbo1 does not have a clear orthologue in the yeasts. It is a member of the MYST family of HATs (Utley and Cote, 2003). There are several diVerent family members in most species, defined by sequence characteristics outside of the MYST HAT domain, and they are typically found in large protein complexes. Metazoan Hbo1 appears to define a distinct subfamily that is diVerent from the MYST subfamilies in yeasts: S. cerevisiae Esa1, Sas2, and Sas3, or S. pombe Mst1 and Mst2 (Burke et al., 2001; Clarke et al., 1999; Go´mez et al., 2005; Iizuka and Stillman, 1999; Reifsnyder et al., 1996; Smith et al., 1998). Thus, the replication function associated with Hbo1 is either absent from yeast or fulfilled by another HAT, whether a MYST family HAT or another family. Intriguingly, one candidate may be Gcn5, which is in the GNAT family of HAT proteins (Berger, 1999; Kurdistani and Grunstein, 2003). Overexpression of one of the subunits of the Gcn5 complex suppresses an mcm5 mutant in yeast although the mechanism is not clear (Dziak et al., 2003). This suggests it may play a positive role in MCM function. Intriguingly, data suggest that budding yeast Gcn5 is functionally redundant with the MYST family member Sas3 (Howe et al., 2001). However, the double mutant gcn5 sas3 arrests with a replicated DNA content, suggesting that replication initiation is not specifically aVected in this strain. Is the interaction between Hbo1 and preRC proteins related to Hbo1’s HAT function? The ING proteins are often subunits of HAT complexes (Gong et al., 2005; Russell et al., 2006). Five diVerent ING subunits in humans were identified and association with diVerent MYST family HAT proteins was examined (Doyon et al., 2006). Ing3 associates with the MYST family protein Tip60/Esa1 (Doyon et al., 2004, 2006), which is implicated in repair (Morrison and Shen, 2005; van Attikum and Gasser, 2005a). Ing4 associates specifically with Hbo1. Ing5, a new member of the family, independently associated with several MYST family HATS: Hbo1, Moz, and Morf. AYnity purification reveals Ing5 also associates with MCMs (Doyon et al., 2006). This may occur through its association with Hbo1, and the phenotypes are compatible: depletion of Ing5 using siRNA specifically blocks human cells in S phase.

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These results suggest that Ing5 is an essential subunit in a replication‐specific Hbo1 complex. Again, the yeasts lack the full complement of ING proteins observed in human cells, but data suggest that the related proteins also function in HAT complexes (Loewith et al., 2000). These data are consistent with the model that an Hbo1‐related HAT complex plays a positive role in preRC assembly. The ING proteins may be helpful in determining whether there is an MCM‐Hbo1‐like connection in yeast. Of course, as we discussed, HATs are not limited to histone substrates (Chen et al., 2001; Hasan and Hottiger, 2002). The HATs Gcn5 and PCAF stimulate replication from a viral replication origin and acetylate polyoma T antigen (a viral equivalent of MCMs) (Xie et al., 2002). Consistent with this, Iizuka et al. (2006) observe acetylation of ORC, Cdc6, Mcm2, and geminin by Hbo1 in vitro. An acetyltransferase activity that is not associated with a HAT enzyme has been identified in association with mammalian Mcm3, and is required for replication initiation (Takei et al., 2001, 2002). These data suggest that acetylation of preRC proteins may provide positive regulation independent of chromatin structure. Protein acetylation has not been examined during the standard characterization of replication proteins, but these results suggest that the potential for regulated acetylation of replication factors should be tested systematically.

IV. Origin Activation and Replication Once the MCMs are loaded, the origin is ready for activation and conversion to an initiation complex (IC) that begins unwinding the DNA and actual DNA synthesis (Bell and Dutta, 2002; Forsburg, 2004; Sclafani et al., 2004). This activation requires several steps, mediated by two distinct kinases, Dbf4‐dependent kinase (DDK) and cyclin‐dependent kinase (CDK) (Fig. 3D and E). The net eVect of these signals is to facilitate loading of additional protein factors that activate the unwinding ability of the MCM proteins and allow DNA synthesis to begin. The pathways controlled by these kinases essentially converge to allow a fail‐safe activation of the origin, and simultaneous inactivation to ensure that the origin cannot refire.

A. Kinases 1. DDK The conserved DDK consists of a catalytic subunit Cdc7 and a regulatory subunit that confers substrate specificity (S. cerevisiae Dbf4; see Table I for nomenclature in other species) (Forsburg, 2004; Jares et al., 2000; Johnston


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et al., 1999; Sclafani, 2000). Dbf4 is transcriptionally and posttranslationally regulated to restrict activity of its kinase partner to S phase of the cell cycle. Current data suggest that the DDK acts at individual replication origins to phosphorylate a subset of MCM proteins and other S phase factors (Forsburg, 2004; Jares et al., 2000; Johnston et al., 1999; Sclafani, 2000). While DDK activity is limited temporally to S phase, its function is not limited to replication: it has also been implicated in additional functions in heterochromatin assembly, gene silencing, repair, and recombination (Axelrod and Rine, 1991; Bailis et al., 2003; Costanzo et al., 2003; Fung et al., 2002; Hollingsworth and Sclafani, 1993; Hollingsworth et al., 1992; Njagi and Kilbey, 1982; Snaith et al., 2000; Takeda et al., 2001). These roles likely involve substrates distinct from the replication proteins; for example, the kinase also phosphorylates the heterochromatin protein HP1/Swi6 in fission yeast, without aVecting its localization (Bailis et al., 2003). Mutants of S. cerevisiae cdc7 increase repression at a damaged silencer, although the mechanism is unclear (Axelrod and Rine, 1991). In vivo data linking DDK and MCMs come from S. cerevisiae genetics. An allele of mcm5 called bob1 bypasses Cdc7 function (Hardy et al., 1997; Jackson et al., 1993). Curiously, Mcm5 is the only MCM subunit that is not phosphorylated by DDK in vitro, so this allele does not simply mimic a phosphorylation event but has a more unusual eVect. An attractive model is that the mcm5‐bob1 mutation causes a change in the shape of the MCM complex that mimics the eVect of DDK phosphorylation on other subunits, and therefore allows subsequent loading of Cdc45 (Hardy et al., 1997; Sclafani et al., 2002). The structural eVect of a bob1 mutation was tested in the archaeal MCM (Fletcher et al., 2003). It results in a modest but discernable shift in the position of one domain of the protein. 2. CDK The cyclin‐dependent kinases (CDKs) are also essential for replication initiation (Blow and Nurse, 1990; Booher and Beach, 1987; Dirick et al., 1995; Duncker et al., 1999; Dutta and Stillman, 1992; Fang and Newport, 1991; Hua and Newport, 1998; Piatti et al., 1996; Sclafani et al., 2002; Tanaka et al., 1997). Importantly, mcm5‐bob1 is still dependent on CDK activity, indicating that this kinase either functions in parallel or downstream of DDK (Jackson et al., 1993; Sclafani et al., 2002). If DDK provides the signal at individual origins, CDK appears to provide the link to the cell cycle. CDK phosphorylates ORC, MCMs, replication protein A (RPA), and DNA polymerase alpha (Coue et al., 1996; Dutta and Stillman, 1992; Foiani et al., 1995; Hendrickson et al., 1996; Hua and Newport, 1998; Masumoto et al., 2002; Montagnoli et al., 2002; Noguchi et al., 2002), as well as the Drc1/Sld2 protein, phosphorylation of which is essential for S phase

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progression (Kamimura et al., 1998; Masumoto et al., 2002; Noguchi et al., 2002; Wang and Elledge, 1999). Phosphorylation by CDKs also inactivates the preRC to prevent its reassembly at the origins (Coverley et al., 1998; Hendrickson et al., 1996; Ishimi and Komamura‐Kohno, 2001; Ishimi et al., 2000; Tanaka et al., 1997). A simplified view is that the DDK pathway activates individual replication origins, while the CDK pathway provides a connection to overall cell cycle progression. These pathways (which certainly have additional substrates) converge to initiate DNA synthesis.

B. Activating the MCM Helicase and Unwinding the DNA 1. Cdc45: A Rate‐Limiting Factor The most significant event in origin firing appears to be binding of a conserved protein called Cdc45 which, like the MCMs, is required both for initiation and elongation steps of DNA replication (Fig. 3D and E) (Hopwood and Dalton, 1996; Labib et al., 2000; Masuda et al., 2003; Reid et al., 1999; Tercero et al., 2000). Cdc45 is less abundant than the MCMs, and it is rate limiting for replication (Aparicio et al., 1999; Edwards et al., 2002; Mimura et al., 2000; Wohlschlegel et al., 2002). If the ratio of MCM to ORC is near 40:1, the ratio of Cdc45 to ORC is about 2:1 (Edwards et al., 2002). Consistent with this, Cdc45 does not assemble at all origins all at once, but apparently as they are fired (Aparicio et al., 1999; Mimura et al., 2000; Zou and Stillman, 1998). A plausible model is that DDK phosphorylation of MCMs results in an allosteric change in the MCM complex that allows binding of a limiting amount of Cdc45 to the MCMs adjacent to the origin. This leaves the bulk of the MCMs inactive. But CDK activity is also required for Cdc45 loading (Mimura and Takisawa, 1998; Zou and Stillman, 1998). This may be mediated at least in part by a protein called TopBP1/Dpb11 (Table I), which is part of a CDK‐regulated complex and is also required for Cdc45 loading (Hashimoto and Takisawa, 2003; Kamimura et al., 1998; Masumoto et al., 2002; Noguchi et al., 2002; Van Hatten et al., 2002). Genetic and molecular analyses indicate that TopBP1/Dpb11 is required both for initiation of DNA replication and, independently, for checkpoint responses to unreplicated or damaged DNA (Garcia et al., 2005). These data suggest that there may be two independent inputs into the replication initiation pathway to create a fail‐safe signaling mechanism. One signal reads the origin identification and preRC assembly of the MCMs, which is activated by DDK phosphorylation. The second signal monitors the cell cycle response provided by the CDK. The consequence of Cdc45 loading is the unwinding of DNA to open a single‐stranded region and allow binding of the single‐strand DNA‐binding


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protein RPA (Mimura et al., 2000; Schepers and DiZey, 2001; Walter and Newport, 2000). There is considerable evidence that MCMs have associated helicase activity, and in vivo data strongly support that all six MCMs are required for unwinding (Forsburg, 2004). Data indicate that Cdc45 may be the crucial cofactor that converts the MCM complex to an active DNA helicase in vivo (Masuda et al., 2003; Pacek and Walter, 2004; Pacek et al., 2006), consistent with Cdc45 association with MCMs at the moving replication fork (Aparicio et al., 1997; Claycomb et al., 2002; Gambus et al., 2006; Pacek et al., 2006; Tercero et al., 2000). But Cdc45 may not be alone in this activity. A ring‐shaped four‐protein complex called GINS is conserved in archaea and eukaryotes. GINS is mutually interdependent for origin binding with Cdc45 and TopBP1/Dpb11 complexes (Kanemaki et al., 2003; Kubota et al., 2003; Marinsek et al., 2006; Takayama et al., 2003). Importantly, GINS is also found at the replication fork and appears to be essential for unwinding activity (Gambus et al., 2006; Marinsek et al., 2006; Pacek et al., 2006). Although its precise role in replication remains unclear, GINS also aVects chromosome segregation and interacts with the chromosome passenger proteins that are involved in kinetochore function (Huang et al., 2005). Whether this is linked to its replication activity remains to be seen. How is the unwinding activity of the replicative helicase (which we now must consider a complex of MCMs, Cdc45, and GINS) linked to the enzymes of DNA synthesis? The best candidate is a rather elusive protein called Mcm10, which appears to participate in Cdc45 loading and also regulates DNA polymerase alpha stability (Fig. 3D and E) (Christensen and Tye, 2003; Fien et al., 2004; Gregan et al., 2003; Ricke and Bielinsky, 2004; Sawyer et al., 2004; Wohlschlegel et al., 2002; Yang et al., 2005). Mcm10 not only appears to be distinct from the MCM/GINS/Cdc45 unwinding activity (Pacek et al., 2006) but is also associated with the replication fork (Ricke and Bielinsky, 2004), which by now is becoming rather crowded. The precise arrangement of the MCM unwinding complex in relation to the other proteins of the fork is not clear, but is the subject of considerable speculation (Laskey and Madine, 2003; Sclafani et al., 2004; Takahashi et al., 2005). 2. Assembling the Active Fork Once unwinding is initiated, the cell assembles the active DNA synthesis machinery (replisome) and begins DNA replication (Fig. 3F) (Bell and Dutta, 2002; Garg and Burgers, 2005; Hubscher et al., 2002; Johnson and O’Donnell, 2005; Waga and Stillman, 1998). First, DNA polymerase alpha, associated with primase, initiates a short RNA primer and begins its extension. Then, a polymerase switch occurs in which a more processive DNA polymerase (delta or epsilon) is loaded, along with PCNA, the ‘‘sliding clamp’’ processivity

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factor (Maga and Hubscher, 2003). PCNA loading depends on a molecular wrench, the clamp‐loading complex replication factor C (RFC) (Jeruzalmi et al., 2001). This promotes the polymerase switch (Tsurimoto et al., 1990; Yuzhakov et al., 1996, 1999). Leading strand synthesis is highly processive and eYcient, once the polymerase switch has occurred, and follows closely behind the helicase. Synthesis on the lagging strand, however, requires multiple cycles of priming and extension as the underlying DNA is exposed; this synthesis occurs in the direction away from the helicase. Finally, the discontinuous fragments of lagging strand DNA, called Okazaki fragments, are processed by nucleases and DNA ligase to assemble a single DNA chain (MacNeill, 2001). Leading and lagging strand synthesis remains coupled to ensure completion of DNA synthesis and to protect the underlying DNA.

V. Replication Elongation Through Chromatin Our concept of the replication fork works very well, until we consider one significant problem: the DNA is not available to the replisome as long as it is wrapped around nucleosomes as chromatin. How does the replication fork gain access to the DNA and unwind the double helix? What protein(s) function as the cowcatcher that moves the nucleosomes out of the way? And, how are the nucleosomes reassembled on the newly synthesized DNA? Studies have identified a number of factors and mechanisms that are likely to play a role in these activities (Falbo and Shen, 2006; Franco and Kaufman, 2004; Krude and Keller, 2001; Polo and Almouzni, 2006). Our current model for how these are coordinated is in Fig. 4.

A. Replicating Through a Chromatin Template The first question to ask is how are the existing nucleosomes moved out of the way of the MCM helicase complex so that it can unwind DNA? During replication, studies suggest old nucleosomes are broken apart and redistributed randomly to the DNA behind the replication fork (Gruss et al., 1993; Lucchini et al., 2001; Randall and Kelly, 1992; Sogo et al., 1986). Thus, some activity must work ahead of the fork to disrupt the nucleosomes and transfer them behind. The most compelling candidate for this activity is called FACT (facilitates chromatin transactions) also referred to as SPN or DUF (Belotserkovskaya and Reinberg, 2004; Formosa, 2003; Singer and Johnston, 2004). FACT is best studied for its role in transcription. Studies show that it is an elongation factor, binding histones, disassembling nucleosomes, and redistributing the old histones behind the RNA polymerase (Belotserkovskaya et al., 2003; Kaplan et al., 2003a; Kireeva et al., 2002; Mason and Struhl, 2003;


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Figure 4 A model‐coupling chromatin assembly to fork passage. See Fig. 3 for identity of replication proteins. Old histones are indicated in solid colors; new histones are indicated by cross‐hatching. (A) The FACT complex dislodges nucleosomes ahead of the fork and passes existing histones behind the fork for reassembly. This may require multiple FACT complexes, or be accomplished by a single linked event at the fork. (B) The CAF‐1 complex binds PCNA and facilitates loading of new histones. ASF activates CAF‐1. (C) In budding yeast, Asf1 and HIR contribute to assembly of nucleosomes independent of the replication fork.

Saunders et al., 2003). This is precisely the role it is likely to play in DNA replication. There is now considerable evidence that FACT plays a role in replication. First, FACT interacts genetically and physically with DNA polymerase alpha (Schlesinger and Formosa, 2000; Wittmeyer and Formosa, 1997; Wittmeyer et al., 1999; Zhou and Wang, 2004), and FACT mutants are sensitive to the DNA synthesis inhibitor, hydroxyurea (Formosa et al., 2001). Second, FACT is required for DNA polymerase to elongate through chromatinized templates: depletion of FACT from Xenopus extracts halts DNA synthesis, independent of transcription (Okuhara et al., 1999), and mutants in budding yeast show replication defects (Schlesinger and Formosa, 2000). Genetic studies in yeast link FACT to the HIR proteins involved in nucleosome assembly (Formosa et al., 2002), which is consistent with evidence that yeast HIR proteins function during S phase (see later). Most intriguingly, a study using a series of tandem aYnity purifications of replication fork proteins show a link between FACT proteins and MCMs

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(Gambus et al., 2006), which is exactly what would be predicted if this complex is opening up the chromatin template ahead of a replication fork. FACT is a heterodimer composed of two subunits, Spt16/Cdc68 and SSRP (yeast Pob3), which are conserved in many systems (Belotserkovskaya and Reinberg, 2004; Formosa, 2003; Singer and Johnston, 2004). The genes encoding Spt16 and Pob3 are essential in S. cerevisiae (Wittmeyer and Formosa, 1997). In yeast, these two subunits interact with the high‐mobility group (HMG) proteins Nhp6a and Nhp6b (Brewster et al., 2001; Formosa et al., 2001); an HMG domain is present in the mammalian SSRP protein but missing in the yeast Pob3 orthologue (Orphanides et al., 1998; Wittmeyer and Formosa, 1997). HMG proteins are abundant small proteins that show dynamic association with chromatin (Bianchi and Agresti, 2005). The Nhp6 proteins appear to be required to recruit the yeast FACT complex to chromatin (Brewster et al., 2001; Formosa et al., 2001), and it seems feasible that the HMG domain of mammalian SSRP similarly directs the complex to chromatin. FACT also interacts with other classes of chromatin proteins as well as histones. It interacts with the chromatin remodeling complex CHD (Kelley et al., 1999; Krogan et al., 2002; Simic et al., 2003) which is required for replication in a cell‐free system (Robinson and Schultz, 2003). Yeast FACT also associates with the MYST family HATs from budding yeast, ScEsa1 and ScSas3 (Formosa et al., 2002; John et al., 2000); the MYST family relative, Hbo1, has already been discussed in the preceding sections. A likely model is that FACT loads onto the chromatin at the time of MCM activation to open access to the DNA, remaining there through its physical association with MCMs and the replication fork complex (Fig. 4A) (Gambus et al., 2006). FACT may also link events at the fork with sister chromatid cohesion, which is established during S phase and essential for normal chromosome segregation (Nasmyth, 2001; Uhlmann, 2003). Ctf4 is a DNA‐polymerase‐associated protein that is required for cohesion and also shows physical and genetic interactions with FACT (Gambus et al., 2006; Hanna et al., 2001; Kouprina et al., 1992; Williams and McIntosh, 2002; Wittmeyer and Formosa, 1997; Zhou and Wang, 2004). With so many protein interactions involving this complex (Fig. 4), a significant current challenge will be to determine how FACT is localized to the fork and dissect the details of its mechanism during S phase.

B. Nucleosome Assembly During Replication Since the process of DNA replication eVectively doubles the amount of DNA in the cell, simply relocalizing existing histones is not suYcient to rechromatinize the template. Histones must also be assembled de novo on the newly synthesized DNA. This process is carried out by conserved assembly


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complexes that contain histone‐binding proteins (histone chaperones). These proteins may act in concert with the chromatin remodeling ATPases (Section I) to load the nucleosomes and correctly position them relative to the DNA (Lusser and Kadonaga, 2003; Polo and Almouzni, 2006). This reaction occurs very quickly and is essential for viability (Han et al., 1987; Jackson, 1990; Kim et al., 1988; Nabatiyan and Krude, 2004; Nelson et al., 2002; Worcel et al., 1978). Nucleosomes are assembled in two stages, beginning with H3‐H4 (Smith and Stillman, 1991; Worcel et al., 1978; reviewed in Franco and Kaufman, 2004; Gunjan et al., 2005; Polo and Almouzni, 2006). The newly assembled nucleosomes following the replication fork may be a mix of old H2A‐H2B and new H3‐H4, or vice versa (Jackson, 1987). This suggests that nucleosome assembly occurs in two stages, which might involve diVerent histone chaperones. Considerable data suggest that H3‐H4 may be deposited as a tetramer (Gruss et al., 1993; Jackson, 1987, 1990), although some suggest it may assemble as a dimer like H2A‐H2B (English et al., 2005; Tagami et al., 2004). Deposition of two H3‐H4 dimers may therefore be coupled. Importantly, assembly of chromatin during S phase is essential. DNA synthesis in the absence of histones leads to damage and activates the checkpoint (Hoek and Stillman, 2003; Ye et al., 2003). Three protein complexes have been identified that are important for de novo nucleosome assembly during S phase (Falbo and Shen, 2006; Franco and Kaufman, 2004; Gunjan et al., 2005; Loyola and Almouzni, 2004; Polo and Almouzni, 2006), each of which is described in the following sections. 1. Chromatin Assembly Factor‐1 Chromatin assembly factor‐1 (CAF‐1) only promotes nucleosome assembly on newly synthesized DNA during replication or repair (Fig. 4B) (Falbo and Shen, 2006; Franco and Kaufman, 2004; Gunjan et al., 2005; Loyola and Almouzni, 2004; Polo and Almouzni, 2006). This activity was first observed in vitro when histones were deposited on plasmid DNA during DNA replication (Smith and Stillman, 1989; Stillman, 1986). Subsequently, it was shown to assemble nucleosomes de novo on newly synthesized DNA (Kaufman et al., 1995; Krude, 1995; Shibahara and Stillman, 1999; Verreault et al., 1996). Consistent with this, CAF‐1 can be localized to replication foci in S phase cells and in repair foci (Green and Almouzni, 2003; Krude, 1995; Taddei et al., 1999). It consists of three subunits and is highly conserved in eukaryotes (Kamakaka et al., 1996; Kaufman et al., 1995, 1997; Kaya et al., 2001; Smith and Stillman, 1989; Verreault et al., 1996). CAF‐1 binds histones H3 and H4 acetylated on K5 and K12. This acetylation pattern is typical of newly synthesized histones although the marks are rapidly lost after assembly (Jackson et al., 1976; Ruiz‐Carillo

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et al., 1975; Taddei et al., 1999). This links CAF‐1 to newly synthesized histones, rather than displaced ‘‘old’’ histones; however, this acetylation is not essential for CAF‐1 to function (Sharp et al., 2001; Sherr, 1994). Some species have two diVerent H3 isoforms; in these, CAF‐1 associates with H3.1, which is only synthesized during S phase (Franklin and Zweidler, 1977; Tagami et al., 2004). This provides one way to temporally limit its activity to S phase. The other H3 isoform, H3.3, is associated with actively transcribed chromatin and is the only form of H3 in the yeasts (Ahmad and HenikoV, 2002; Baxevanis and Landsman, 1998). Physical association with PCNA, the sliding clamp processivity factor, localizes CAF‐1 to regions of new DNA synthesis (Fig. 4B; see Section IV.C) (Krawitz et al., 2002; Moggs et al., 2000; Shibahara and Stillman, 1999; Zhang et al., 2000). This localization, and the preference for the S‐phase synthesized histone H3.1, limits CAF‐1 activity to replication and repair foci. CAF‐1 also associates with the chromatin remodeling complex ISWI (ACF/CHRAC) (Corona and Tamkun, 2004), which, as discussed earlier, has been linked to replication initiation and replication through heterochromatin (Alexiadis et al., 1998; Collins et al., 2002). Evidence in Drosophila suggests that ISWI, when part of a complex called ACF, works with CAF‐1 to assemble properly spaced nucleosome arrays (Ito et al., 1997, 1999). CAF‐1 is essential for S phase progression in human cells, and inhibiting its activity leads to S phase arrest and DNA damage (Hoek and Stillman, 2003; Ye et al., 2003). This suggests that proper assembly of the chromatin is essential for S phase progression and cell viability. Curiously, however, CAF‐1 is not essential for viability in yeast, although mutations do result in disruptions of silencing and genome stability (Enomoto and Berman, 1998; Kaufman et al., 1997; Monson et al., 1997; Myung et al., 2003). This indicates that other histone assembly factors can function redundantly with CAF‐1 during S phase, at least in S. cerevisiae. It has been suggested that this redundancy reflects the single H3 isoform in yeast, which eliminates one form of temporal regulation; in other species, chromatin assembly factors may be restricted temporally by availability of diVerent isoforms (see discussions in Franco and Kaufman, 2004; Loyola and Almouzni, 2004). Since gene expression often reflects chromatin structure and the histone code (see Section I), it is perhaps not surprising that CAF‐1‐mediated assembly of histones can aVect transmission of these signals and aVect silencing (Franco and Kaufman, 2004; Wallace and Orr‐Weaver, 2005). Consistent with this function of CAF‐1, mutations in its binding partner PCNA can also aVect silencing (Sharp et al., 2001; Zhang et al., 2000). This is not a yeast‐specific eVect, because silencing defects are also associated with CAF‐1 mutations in multicellular eukaryotes (Kaya et al., 2001; Quivy et al., 2001; Tchenio et al., 2001). Strengthening this link, CAF‐1 interacts with traditional mediators of silencing, including HP1 (Murzina et al., 1999;


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Quivy et al., 2004) and the MYST family HAT Sas2, which is required to establish silencing boundaries (Katan‐Khaykovich and Struhl, 2005; Kimura et al., 2002; Meijsing and Ehrenhofer‐Murray, 2001; Suka et al., 2002). Again, we see a link between silencing and heterochromatin, and events in DNA replication. The eVect of CAF‐1 on inheritence of epigenetic marks is discussed in Section V.C. 2. Anti‐Silencing Function‐1 (ASF) An additional conserved protein factor acts with CAF‐1 to promote eYcient replication‐coupled histone deposition (Fig. 4B) (Falbo and Shen, 2006; Franco and Kaufman, 2004; Loyola and Almouzni, 2004). A complex containing Asf1 stimulates CAF‐1 activity (Mello et al., 2002; Sharp et al., 2001; Tyler et al., 1999) but cannot assemble nucleosomes on its own (Tyler et al., 1999). It binds to histones H3‐H4 (Tyler et al., 1999), but unlike CAF‐1 it does not distinguish between H3 isoforms in higher cells (Tagami et al., 2004). Intriguingly, ASF has been linked directly to replication fork progression by regulating stability of proteins at the replication fork (Franco et al., 2005). Whether this is a mechanism to couple ASF‐mediated nucleosome deposition to the replication fork, or an independent activity, remains to be determined. Asf1 has also been shown to interact with Rad53, the DNA damage and S phase checkpoint kinase (Daganzo et al., 2003; Emili et al., 2001; Hu et al., 2001; Schwartz et al., 2003). Significantly, yeast ASF is essential in the absence of CAF‐1, or when the interaction between CAF‐1 and PCNA is disrupted (Krawitz et al., 2002; Sharp et al., 2001; Sutton et al., 2001; Tyler et al., 1999). Thus, it is not simply a CAF‐1 cofactor, but also functions independently. Consistent with this, its phenotypes are distinct from those of CAF‐1 mutants (Ramey et al., 2004; Tong et al., 2004; Tyler et al., 1999). Mutants in asf1 show elevated levels of gross chromosomal rearrangements (Myung et al., 2003) and increased plasmid loss rates that might reflect defects in DNA replication or chromosome segregation (Le et al., 1997). Because yeast Asf1 acts to stimulate CAF‐1 (perhaps as a histone donor) but becomes essential in the absence of CAF‐1, we can conclude that an alternative histone chaperone‐ deposition complex can work along with Asf1 in S phase in budding yeast. The likely candidate is the HIR complex. 3. HIR Proteins The HIR complex is another H3‐H4 chaperone: HIRA in mammals, and Hir1, 2, and 3 in yeast (Franco and Kaufman, 2004; Gunjan et al., 2005; Loyola and Almouzni, 2004). Genetic interactions with FACT complex genes (Formosa et al., 2002) and ASF1 (Sharp et al., 2001; Tyler et al., 1999) suggest

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that HIR proteins also contribute to chromatin assembly. In the absence of CAF‐1, Asf1 and HIR proteins compensate to allow histone deposition (Fig. 4C) (Kaufman et al., 1998; Krawitz et al., 2002; Ray‐Gallet et al., 2002). In contrast to CAF‐1, vertebrate HIRA binds the H3.3 isoform that is associated with active chromatin, so it is not temporally linked to S phase. Thus, HIR provides a replication‐independent histone assembly activity (Ahmad and HenikoV, 2002; Franco and Kaufman, 2004; Loyola and Almouzni, 2004; Tagami et al., 2004). This provides a clear functional distinction between CAF‐1 and HIR, but they are linked through their use of Asf1 as a common partner to prime either deposition complex (Franco and Kaufman, 2004; Gunjan et al., 2005; Loyola and Almouzni, 2004).

C. Maintaining Epigenetic Marks During Replication As described earlier, heterochromatin in most eukaryotes is characterized by distinct patterns of histone modification and binding of HP1 and serves as a useful model for epigenetic inheritence. We have described how nucleosomes are disrupted during S phase and new histones are recruited. But how are epigenetic marks such as those in heterochromatin preserved during DNA replication? Most studies have focused on the establishment of silencing de novo rather than maintenance of existing signals. There is a long‐standing link between DNA replication and establishment of silencing (Miller and Nasmyth, 1984). Early models suggested a direct link between replication fork passage and silencing, but this pleasing theory has been disproven by showing that silencing, while S phase dependent, does not require a passing replication fork (Kirchmaier and Rine, 2001; Li et al., 2001). However, compelling links to replication proteins show that there is a connection, at least temporally, between silencing and replication. Links between replication proteins and heterochromatin are uncovering the mechanism (Maison and Almouzni, 2004; Wallace and Orr‐Weaver, 2005). As discussed in Section II, replication origins are closely tied to transcriptional silencing. First, silencer elements in both budding and fission yeast include potential origins of replication (Ekwall et al., 1991; Fox et al., 1993). While the mechanism involving replication proteins in this process is elusive, the data suggest replication activity at silencers is not essential. Rather, they are likely to be a platform for ORC loading (Foss et al., 1993; Leatherwood and Vas, 2003; Loo et al., 1995). This is supported because silencing phenotypes in ORC are genetically separable from those aVecting replication (Bell et al., 1995; Dillin and Rine, 1997). Importantly, S. cerevisiae ORC also interacts with the silencing protein Sir1 (Bose et al., 2004; Fox et al., 1997; Triolo and Sternglanz, 1996), suggesting that ORC contributes to recruitment of silencing proteins.


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Intriguingly, Sir1 is also found at centromeres and interacts with CAF‐1 (Sharp et al., 2003). Although Sir1 is not found in other eukaryotes, it functions analogously to the HP1/Swi6 protein in many respects (Pillus and Rine, 2004). Significantly, HP1 interacts with ORC in Drosophila (Huang et al., 1998; Pak et al., 1997; Shareef et al., 2001), Xenopus (Pak et al., 1997), humans (Prasanth et al., 2004), and fission yeast (J. M. Bailis, P.‐C. Li, and S. L. Forsburg, unpublished data). Data from a number of cell types suggest ORC binds specifically in heterochromatic regions (Lidonnici et al., 2004; Palacios DeBeer et al., 2003; Prasanth et al., 2004). Many of these data support an interpretation that ORC helps to recruit HP1 to specific regions to reestablish heterochromatin marks, although it has also been suggested that HP1 recruits ORC to facilitate replication (Leatherwood and Vas, 2003). However, as always, things are more complicated. Although ORC’s silencing function is genetically separable from its essential replication role, other preRC proteins nevertheless have eVects on silencing that suggest complicated links between replication and heterochromatin. Mutations in budding yeast CDC7, CDC45, and other replication genes can restore silencing to defective silencers (Axelrod and Rine, 1991; Ehrenhofer‐Murray et al., 1999). Does this suggest that reduced eYciency of firing of those origins in replication mutants increases their silencing potential? Some data suggest origin firing reduces silencing, but this may not be uniform (Dubey et al., 1991; McConnell et al., 2006; Zappulla et al., 2002). On the other hand, mutations in MCM5 and MCM10 genes may disrupt silencing, suggesting these proteins actively promote silencing (Dziak et al., 2003; Liachko and Tye, 2005). Again, the mechanism is unclear, but it is interesting to speculate that this might involve the distal MCMs (MCM paradox, see Section III) or known interactions with RNA polymerase (Forsburg, 2004). The HP1/Swi6 heterochromatin protein is an additional substrate for the S. pombe Hsk1 (Cdc7) kinase (Bailis et al., 2003). This is genetically separable from the function of the kinase in promoting replication (it phosphorylates MCMs, see Section III), and appears to be important in establishing centromeric cohesion. However, because this occurs in S phase, it provides a way of temporally linking chromatin events to the cell cycle, independent of a replication fork (Bailis and Forsburg, 2003). How is existing HP1 assembled onto newly replicated heterochromatin? A significant link between replication and heterochromatin comes from S. pombe in which DNA polymerase alpha (see Section IV.B) binds to the HP1/Swi6 protein and is required for HP1/Swi6 recruitment to heterochromatin and subsequent silencing (Ahmed et al., 2001; Nakayama et al., 2001a; Singh and Klar, 1993). This implicates a passing replication fork with maintenance of HP1 binding and heterochromatin structure. However, the polymerase domain responsible for the interaction is not conserved, suggesting this means of recruiting Swi6 may be specific to fission yeast. What else can do the job?

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The key may be chromatin assembly. Importantly, CAF‐1 also binds HP1 (Murzina et al., 1999; Quivy et al., 2004). This provides a possible mechanism for maintenance of heterochromatin through DNA replication, which is the time when CAF‐1 is active. Such a model is consistent with mutations in CAF‐1 aVecting maintenance of silencing (Enomoto and Berman, 1998; Kaufman et al., 1997; Kaya et al., 2001; Quivy et al., 2001; Tchenio et al., 2001). PCNA recruits CAF‐1 to the fork and CAF‐1 recruits HP1 as it is dislodged by replication through the heterochromatin. Association with CAF‐1 leads to a local increase in HP1 concentration. Since HP1 can recruit the H3K9 methyltransferase (see Section I.C; Fig. 2), this localization provides a mechanism to restore the epigenetic marks on the newly deposited histones, to reestablish heterochromatin on newly synthesized DNA (Maison and Almouzni, 2004; Wallace and Orr‐Weaver, 2005). This pleasing mechanism may provide a paradigm for how epigenetic marks are maintained during replication. The challenge will be to determine how other histone modifiers are recruited to the replication apparatus to facilitate inheritance of other marks on newly synthesized DNA.

VI. Conclusions As we have described, we now have a good grasp of the events required to initiated DNA replication at eukaryotic replication origins. For reasons of space, we have not been able to consider the linked issues of DNA repair and checkpoint responses, but these also affect the picture we have painted and have recently been reviewed (Morrison and Shen, 2005; Thiriet and Hayes, 2005; van Attikum and Gasser, 2005a,b; Vidanes et al., 2005; Wurtele and Verreault, 2006). The challenge now is to understand how the events of replication initiation and genome duplication are mechanistically linked to chromatin modification, remodeling, and assembly. In this chapter, we have introduced some of the connections between molecules aVecting replication and chromatin structure. These two areas of study, so long distinct, are beginning to merge. We can expect a powerful synergy of approaches to investigate the complete question of chromosome replication in a chromatin context.

Acknowledgments We thank Jennifer Aparicio, Judd Rice, and members of our laboratory for critical reading of the chapter and helpful discussions. We are indebted to colleagues who shared manuscripts, reprints, and many conversations, and we apologize to those whom we were unable to cite due to space limitations. SLF acknowledges support from NIH GM059321 and American Cancer Society RPG‐00–132–04‐CCG. APT is supported by NIH T32 CA009320.


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The Regulatory Network Controlling the Proliferation–Meiotic Entry Decision in the Caenorhabditis elegans Germ Line Dave Hansen* and Tim Schedl { *Department of Biological Sciences, University of Calgary Calgary, Alberta, Canada T2N‐1N4 { Department of Genetics Washington University School of Medicine Saint Louis, Missouri 63110

I. Introduction II. Temporal and Spatial Pattern of Proliferation and Initiation of Meiotic Development III. Summary of the Core Regulatory Pathway Controlling the Proliferation–Meiotic Development Decision A. GLP‐1 Notch Signaling B. GLD‐1 and GLD‐2 Meiotic Entry Pathways IV. Control of the Adult Pattern of Proliferation and Meiotic Entry A. Stem Cells Versus Transit Amplifying Cells B. Spatial Control of Notch Signaling V. Regulation of the Activity of the GLD‐1 and GLD‐2 Pathways A. GLP‐1 Notch Signaling Control of GLD‐1 and GLD‐2 Pathway Activity B. FBF Functions Between GLP‐1 Notch Signaling and the GLD‐1 and GLD‐2 Pathways C. FOG‐1, FOG‐3, and FEM‐3 Function Redundantly with FBF D. FBF‐1 and FBF‐2 Functions Are not Completely Redundant VI. Additional Components and Pathways Regulating the Proliferation–Meiotic Entry Decision A. FBF May also Function in the GLD‐2 Pathway B. Cross Talk Between the GLD‐1 and GLD‐2 Pathways C. A Third Pathway Downstream of Notch Signaling D. MOG Gene Products Act to Promote Meiotic Entry E. ATX‐2 May Promote Proliferation Independent of GLP‐1 Notch Signaling F. Connection Between the Proliferation–Meiotic Development Decision and Germ Line Sex Determination VII. Conclusions Acknowledgments References

The germ line of sexually reproducing animals, at some point in development, consists of both proliferating and diVerentiating cells. Proliferation is needed to increase cell number, ensuring that a suYcient Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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quantity of gametes is produced. Meiotic development is needed to produce gametes that can support embryogenesis, each with half the ploidy of the somatic cells. For the reproductive strategy of a given species, regulating the timing and number of gametes, and thus controlling the timing of diVerentiation and the extent of proliferation, is very important for reproductive fitness. Therefore, animals have evolved regulatory mechanisms that tightly control and balance the proliferation–initiation of meiotic development (meiotic entry) decision. Genetic analysis has identified signaling mechanisms involved in controlling this balance in some animals, including mice, Drosophila, and Caenorhabditis elegans. In this chapter, we present our understanding of the genetic hierarchy controlling the proliferation–meiotic entry decision in C. elegans. A core regulatory network controls the decision under all known conditions (developmental stage, sex, and growth temperature). It consists of a canonical Notch signaling pathway promoting proliferation by inhibiting two redundant mRNA regulatory pathways, the GLD‐1 and GLD‐2 pathways, which promote meiotic entry. Superimposed on the core network is a complex set of factors, some yet to be identified, and many with regulatory relationships still poorly understood, which control the activities of the GLD‐1 and GLD‐2 pathways and possibly parallel pathways. Some of the complexity arises from these regulators acting only under certain conditions. We also highlight major areas where we lack knowledge. For example, it is unknown if the entire population of proliferating cells are stem cells capable of self‐renewal or if only a small portion are stem cells and the rest are transit amplifying cells. ß 2006, Elsevier Inc.

I. Introduction The germ line is a specialized tissue in sexually reproducing animals that is necessary to pass genetic material to future generations. Since the gametes of two parents contribute genetic information to each oVspring, it is necessary for diploid germ cells to undergo meiosis, a complex developmental program with two specialized cell divisions, to produce haploid gametes that can unite at fertilization to generate the zygote. Germ cells in the parents, in particular the germ line stem cells, must first proliferate in order to increase cell number and thus provide a pool of cells from which gametes can be formed. The reproductive fitness of the animal depends upon a balance being maintained between proliferation and initiation of meiotic development. How this balance is maintained varies between species and often between sexes of the same species. In female mammals, all germ cells exit proliferation and initiate meiotic development in the embryo (Morelli and Cohen, 2005). The germ cells then arrest in prophase I until sexual maturity. In mammalian males, germ cells do not enter meiosis until after birth, but prior to puberty.

6. Proliferation–Meiotic Entry in Caenorhabditis elegans

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A population of proliferative cells is maintained throughout adult life, allowing the male to produce large numbers of gametes and remain reproductive throughout adulthood (Morelli and Cohen, 2005). Human males produce 1012–1013 gametes over their lifetime (Reijo et al., 1995). Evidence suggests that retinoic acid (RA) promotes mammalian germ cells to enter meiosis and that RA is degraded in embryonic testes, thereby preventing embryonic meiosis in males (Koubova et al., 2006). In Drosophila, the transition from proliferation to meiotic development is regulated by signals emanating from somatic niche cells (Wong et al., 2005). Germ cells that remain in direct contact with the cap cells (female) or the hub (males) remain proliferative. The asymmetric sisters of these cells will go through additional rounds of proliferation, transit amplification, but are normally destined to enter meiosis. The distinction between the stem cells and the transit amplifying cells has become less clear with the observation that some transit amplifying cells can revert into stem cells in both the male and female Drosophila germ lines (Brawley and Matunis, 2004; Kai and Spradling, 2004). In this chapter, we present our understanding of how the decision between germ line proliferation (self‐renewal) and initiation of meiotic development (diVerentiation) in Caenorhabditis elegans is controlled. First is a brief background on C. elegans and the pattern of germ line proliferation and meiotic development. We then present the core proliferation–meiotic entry regulatory network. The connection between the regulatory network and the pattern of proliferation and meiotic entry follows, with subsequent elaboration on the network model by incorporating additional genes and control processes. Throughout we highlight major open questions. For other reviews on the proliferation–meiotic entry decision in C. elegans please see Crittenden et al., 2003; Kimble and Crittenden, 2005; Seydoux and Schedl, 2001; and Wong et al., 2005.

II. Temporal and Spatial Pattern of Proliferation and Initiation of Meiotic Development C. elegans exists as two sexes, hermaphrodite and male. Males generate as many as 3000 sperm and reproduce only by mating with a hermaphrodite. Hermaphrodites are somatically female, with the germ line producing first sperm and then oocytes. Hermaphrodites can reproduce either using their own sperm, by self‐fertilization, or by using male sperm when mating has occurred. The hermaphrodite generates 320 sperm and thus only 320 self‐ progeny. However, the hermaphrodite can generate many more oocytes throughout her reproductive lifespan of 2 weeks, and when mated can produce as many as 1400 progeny (Kimble and Ward, 1988). Since about half of all germ cells undergoing oogenesis die through the process of

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physiological apoptosis (Gumienny et al., 1999), more than 2800 germ cells undergoing gametogenesis must have been generated. It appears that the general processes regulating the decision between proliferation and meiotic entry are similar between the male and the hermaphrodite, and most research has focused on studying the hermaphrodite germ line. Therefore, we will mostly discuss the hermaphrodite germ line. The adult hermaphrodite has two symmetric U‐shaped gonad arms, each containing 1000 germ cells (Fig. 1A). In each gonad arm the germ line has a distal to proximal polarity, with a population of proliferative cells at the distal end followed by entry into meiotic prophase, progression through meiotic prophase, and gametogenesis as germ cells move proximally (Fig. 1B). The adult pattern is formed postembryonically through cell division, and diVerentiation in the germ line and somatic gonad. In the hatched L1 larva, there are two germ cells (Z2, Z3) and two somatic cells (Z1, Z4) surrounded by a basement membrane. Z1 and Z4 divide by an essentially invariant cell lineage pattern to generate the somatic gonad while Z2, Z3, and all germ cell daughters divide by a random pattern (Kimble and Hirsh, 1979). If the animal has suYcient food, Z2 and Z3 begin to proliferate in the L1 stage. Z1 and Z4 also divide, generating 12 somatic gonad cells by the end of the L1 stage, including two distal tip cells (DTCs) that promote germ cell proliferation (see later). By the end of the L2 stage in the hermaphrodite, the remaining ten somatic gonad cells rearrange to form the centrally located somatic gonad primordium, bisecting the proliferating germ cells into two populations, one anterior, capped by the anterior DTC, and other posterior, capped by the posterior DTC. In the L3 and L4 stages, the respective DTCs lead the two growing gonad arms to form the anterior and posterior U‐shaped gonad arms, centered around the proximal somatic gonadal uterine and spermathecal structures generated from the somatic gonad primordium. During this time period the somatic gonadal sheath lineage forms a single cell layer covering the proximal 3/4 of each gonad arm, positioned between the germ cells and the surrounding basement membrane (Hall et al., 1999). Polarization of the germ line into a proliferative population near the DTC and initiation of meiotic development more proximally begins in early L3 and continues throughout the reproductive life (Hansen et al., 2004a; Kimble and White, 1981). The size of the region containing the proliferative cells, the proliferative or mitotic zone, increases as the hermaphrodite ages and as the gonad arm lengthens. In early L3, the proliferative zone is 12 cell diameters long, but increases in size to 20 cell diameters long in the adult (Fig. 2A) (Crittenden et al., 1994; Hansen et al., 2004a; Lamont et al., 2004). Proliferation is nevertheless very rapid as the germ cell number increases 20‐fold from early L3 to adulthood. Robust proliferation continues in adulthood, during which the proliferative zone contains 200–250 cells, and then declines as self‐sperm is exhausted (Killian and Hubbard, 2005;


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Figure 1 Basic anatomy of the C. elegans hermaphrodite germ line. (A) Diagram of an adult C. elegans hermaphrodite emphasizing the position and polarity of the germ line. The gonad consists of anterior and posterior U‐shaped arms, centered around the proximal uterus (containing light‐green shaded embryos). At the distal end of each gonad arm, capped by the distal tip cell (DTC; yellow), is a population of proliferating germ cells (green proliferative nuclei). As cells move proximally, toward the uterus, they enter meiotic prophase in the transition zone (red meiotic prophase nuclei). Cells continue proximally as they progress through meiotic prophase, eventually forming gametes. The first 40 germ cells in each gonad arm that enter meiosis in larvae form sperm (blue, which are stored in the spermatheca) while all remaining germ cells that enter meiosis form oocytes (orange). Ovulated oocytes are fertilized as they pass through the spermatheca and embryos begin their cell divisions while in the uterus, and then are passed to the environment through the vulva. (B) Fluorescent image of one young adult hermaphrodite gonad arm that has been dissected away from the body of the animal (distal to the left). Surface view distally, interior view proximally. The top and bottom image are of the same arm. Top image (blue) shows DNA morphology from DAPI staining. Bottom image shows the proliferative cell nuclei (green) detected with antibodies specific to REC‐ 8 while meiotic prophase nuclei (red) are detected with HIM‐3 antibodies (see text for details). Most of the germ line is syncytial, with each nucleus partially enclosed by plasma membranes (Hall et al., 1999; Hirsh et al., 1976). By convention, the term ‘‘germ cell’’ refers to a nucleus, its associated cytoplasm, and surrounding membrane. For more comprehensive reviews of C. elegans germ line development please see Hubbard and Greenstein (2000, 2005) and Schedl (1997).

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Figure 2 The core regulatory network controlling the proliferation–meiotic entry decision. (A) Magnified view of the distal end of the dissected gonad arm fluorescent micrograph in Fig. 1. Surface view. The top image shows DNA morphology using DAPI staining (blue). The bottom image is the same gonad arm with proliferative cell nuclei identified as staining with antibodies against REC‐8 (green) and meiotic prophase nuclei staining with antibodies against HIM‐3 (red). The proliferative zone is the region from the very distal end up to, but not including, the most distal cell with crescent‐shaped morphology (transition zone nucleus). The transition zone is the region from the most distal crescent‐shaped cell to the most proximal crescent‐shaped cell. The meiotic entry region is the portion of the germ line that contains both proliferative (REC‐8–positive, HIM‐3–negative) and meiotic (REC‐8–negative, HIM‐3–positive) cells in the same plane (Hansen et al., 2004a). The transition from proliferation to meiotic development is not synchronous, as some proliferative cells (green nuclei) are more proximal than some cells that have initiated meiotic development (red nuclei) (Hansen et al., 2004a). (B) Panels showing the two alternate states of the core regulatory network controlling the proliferation–meiotic entry decision. The top panels shows the relative activity of the network near the DTC, where cells are proliferative, while the bottom panel shows the relative activity of the network near the transition zone, where cells enter meiotic


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Lamont et al., 2004). The first 40 germ cells that initiate meiotic development in each gonad arm develop as sperm and all subsequent germ cells that enter meiosis develop as oocytes. Following meiotic entry, germ cells orderly progress through the stages of meiotic prophase and gametogenesis in an assembly‐line‐like fashion (Fig. 1B). The switch from proliferation to initiation of meiotic development is assessed primarily by cytological examination of nuclear morphology. A convenient approach is 40 ,6‐diamidino‐2‐phenylindole (DAPI) staining to visualize chromosome morphology and organization. In the proliferative region, cells in mitosis (metaphase, anaphase, and telophase) display the expected DAPI morphology (Fig. 2A). Just proximal to the proliferative region is the transition zone where cells are in the earliest stages of meiotic prophase (leptotene/zygotene). These cells display an asymmetric organization of the nucleolus and chromatin, associated with pairing and synapsis, leading to a crescent‐shaped DAPI‐stained nuclear morphology (Fig. 2A) (Dernburg et al., 1998; Francis et al., 1995a; MacQueen and Villeneuve, 2001). Antibody staining has been used to confirm and refine these regions. Particularly useful has been antibody staining to REC‐8, a meiotic cohesion that under mild fixation conditions is detected in the nucleoplasm and chromatin of all proliferating germ cells (Hansen et al., 2004a; Pasierbek et al., 2001), and antibody staining to HIM‐3, a meiotic chromosome axes protein that is found in all meiotic prophase nuclei (Fig. 2A) (Zetka et al., 1999). REC‐8 staining and HIM‐3 staining are almost completely mutually exclusive. Cell nuclei that are REC‐8–positive are HIM‐3–negative and vice versa. There are, however, a small number of weakly REC‐8–positive cell nuclei per gonad arm that are also weakly positive for HIM‐3. These cells are perhaps in the very early stages of meiotic prophase. It still is not completely clear how accurately these markers show commitment to entering meiotic prophase. It is likely that this commitment is made around the time of premeiotic S‐phase, which perhaps is prior to HIM‐3 loading on to the chromosomal axes. However, the REC‐8 and HIM‐3 markers have emphasized the asynchronous nature of the transition from proliferation to entry prophase. Factors with high activity are shown in large font, while factors with little or no activity are shown in small font. Arrows depict activating or positive interaction between factors while barred lines depict an inhibitory relationship. Dashed lines indicate interactions that are no longer active. For cells near the DTC (top panel), GLP‐1 Notch signaling is active due to interaction of the LAG‐2 ligand with the GLP‐1 receptor (see text and Fig. 3). Active GLP‐1 Notch signaling promotes proliferation by inhibiting the GLD‐1 and GLD‐2 pathways that promote meiotic entry. For cells near the transition zone (bottom panel), GLP‐1 Notch signaling is reduced or eliminated. In the absence of repression mediated by GLP‐1 Notch signaling, the GLD‐1 and GLD‐2 pathways promote the activities of factors involved in entry into meiotic prophase and inhibit the activities of factors necessary for proliferation. See text for details.

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into meiotic prophase in the adult hermaphrodite. All cells do not enter meiosis at the same position in the gonad arm. Rather, in the adult a region of approximately seven cell diameters, referred to as the meiotic entry region, consists of a mixture of cells, some of which are proliferative (REC‐8–positive and HIM‐3–negative) and others that are in early meiotic prophase (REC‐8–negative and HIM‐3–positive) (Hansen et al., 2004a) (Figs. 1 and 2). Therefore, there does not appear to be a strict boundary separating proliferative and meiotic cells. Rather, the boundary is relatively ragged. This diVers from the situation in larval animals where the transition is very abrupt (Hansen et al., 2004a). This diVerence is possibly due to the relatively low number of cells in the larval germ line as compared to the adult.

III. Summary of the Core Regulatory Pathway Controlling the Proliferation–Meiotic Development Decision A. GLP‐1 Notch Signaling The first insight into the control of the proliferation–meiotic development decision came from a pivotal experiment by Kimble and White (1981), where they sought to determine the consequences to germ line development of ablating the DTC with a laser microbeam. They found that the proliferative population was lost with all germ cells prematurely initiating (and completing) meiotic development. Thus the DTC must signal the germ line to promote proliferation and/or inhibit meiotic development. They also demonstrated that mispositioning the DTC created a new polarized germ line organization with proliferation adjacent to the ectopic DTC followed at a distance by meiotic entry. Together these experiments demonstrated that the DTC is necessary and suYcient to create a niche for the proliferative cells and polarizing germ line development. These findings lead to the idea that genes and cells that function in the proliferation–meiotic development decision could be identified experimentally from mutations or cell ablations that caused either a premature meiotic entry phenotype, like with the DTC ablation, or an overproliferation (tumorous) phenotype due to a failure of germ cells to enter meiotic prophase. Genetic and molecular analysis has revealed that the conserved Notch signaling pathway is the principal molecular mechanism by which the DTC niche controls proliferation (Fig. 2B). The four members of the canonical Notch signaling pathway are the DSL ligand (acronym for Delta‐Serrate‐ LAG‐2), Notch receptor, CSL transcription factor (CBF‐1/RBPJ‐Su(H)‐ LAG‐1), and the MAML transcriptional coactivator (Mastermind‐LAG‐3) (Lubman et al., 2004). In the C. elegans gonad the conserved transmembrane


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LAG‐2 DSL ligand is expressed in the DTC (Fig. 3) (Henderson et al., 1994; Tax et al., 1994). As the DTC comes in contact with germ cells, LAG‐2 interacts with the germ line‐expressed GLP‐1 Notch transmembrane receptor (Austin and Kimble, 1987, 1989; Yochem and Greenwald, 1989). This interaction is thought to result in cleavage of the intracellular portion of GLP‐1 from the transmembrane portion of the protein (Mumm and Kopan, 2000). The intracellular portion of GLP‐1, referred to as Notch intracellular domain (NICD) or GLP‐1(INTRA), then is thought to translocate to the nucleus and bind the LAG‐1 CSL transcription factor (Fig. 3) (Christensen et al., 1996). CSL transcription factors have been shown in other systems to be transcriptional repressors, but on binding with INTRA, they become transcriptional activators (Mumm and Kopan, 2000). The same is likely to occur with LAG‐1. GLP‐1(INTRA) and LAG‐1 form a ternary complex

Figure 3 GLP‐1 Notch signaling in the distal gonad. On the right is a diagram of the proliferative zone and part of the transition zone of an adult hermaphrodite; surface view. The somatic DTC (yellow) caps the germ cells in the very distal end of the gonad and has projections that extend more proximally. Some of the proliferative germ cells (nuclei shown in dark green) are in direct contact with the DTC while others are not. As cells move proximally, they enter meiotic prophase and their DNA takes on a crescent‐shaped morphology (red). On the left is a blow‐up of contact between the DTC and adjacent germ cells. Also shown are depictions of the relative positions of the canonical GLP‐1 Notch pathway components during active signaling. LAG‐2, the DSL ligand, is expressed on the surface of the DTC. It comes in direct contact with the GLP‐1 Notch receptor, which is expressed on the surface of the germ cells. This causes cleavage of the receptor and subsequent translocation of the intracellular portion, GLP‐1 (INTRA), to the nucleus. GLP‐1(INTRA) forms a ternary complex with the LAG‐1 CSL transcription factor and the SEL‐8/LAG‐3 MAML transcriptional coactivator. This complex then activates genes necessary for proliferation. For cells approaching the transition zone, GLP‐1(INTRA) will be reduced or absent, the ternary complex will not be assembled, GLP‐1 Notch target genes will not be transcribed, and cells will enter meiotic prophase. See text for details.

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with SEL‐8/LAG‐3 MAML, then as a complex activates transcription of downstream target genes (Doyle et al., 2000; Nam et al., 2006; Petcherski and Kimble, 2000; Wilson and Kovall, 2006). If the activities of any of these canonical Notch pathway components are reduced or eliminated in the C. elegans germ line, germ cells cease proliferating and prematurely enter meiosis; similar to the phenotype observed when the DTC is ablated (Austin and Kimble, 1987; Christensen et al., 1996; Doyle et al., 2000; Henderson et al., 1994; Lambie and Kimble, 1991; Petcherski and Kimble, 2000; Tax et al., 1994). Constitutively active glp‐1 gain‐of‐function (gf) alleles have been identified that, based on worm genetic and mammalian cell culture experiments, result in ligand independent generation of GLP‐1(INTRA) (Berry et al., 1997; Pepper et al., 2003; Vooijs et al., 2004). In glp‐1(gf) mutants, germ cells fail to enter meiosis, but instead continue to proliferate and form a germ line tumor, which is opposite to the premature meiotic entry phenotype caused by a loss of GLP‐1 Notch signaling. Thus, the GLP‐1 Notch signaling pathway acts as a binary switch; when it is active proliferation results, when it is inactive cells enter meiosis (Fig. 2). Cells close to the DTC have active GLP‐1 Notch signaling, thus are proliferative. As the cells move proximally away from the DTC, the LAG‐2 ligand and GLP‐1 receptor are no longer in contact, resulting in a lack of GLP‐1(INTRA) and the resulting loss of the GLP‐1(INTRA), LAG‐1, and SEL‐8/LAG‐3 ternary transcriptional activation complex. Therefore, genes involved in promoting proliferation and/or inhibiting meiotic entry are no longer active, allowing cells to enter meiotic prophase as they move into the transition zone.

B. GLD‐1 and GLD‐2 Meiotic Entry Pathways Two redundant mRNA regulatory pathways, GLD‐1 and GLD‐2, named after their founding members, function downstream of GLP‐1 Notch signaling to promote meiotic development (Kadyk and Kimble, 1998) (Fig. 2B). GLD‐1 is a maxi‐KH/STAR domain containing RNA‐binding protein similar to mouse Quaking and Drosophila held out wings (How) (Jones and Schedl, 1995). Also within the GLD‐1 pathway is NOS‐3, which is a protein similar to Drosophila Nanos (Hansen et al., 2004b; Kraemer et al., 1999). GLD‐2 is a poly(A) polymerase catalytic subunit that complexes with GLD‐ 3, a BicC‐related KH domain containing RNA‐binding protein, which perhaps provides the mRNA specificity for the catalytic domain (Eckmann et al., 2004; Wang et al., 2002). The GLD‐1 and GLD‐2 pathways function redundantly in promoting meiotic entry; each is suYcient for meiotic entry to occur. If the activity of a gene (or genes) in one pathway is eliminated, proliferation and meiotic entry occur essentially normally (Eckmann et al., 2004; Francis et al., 1995b; Hansen et al., 2004b; Kadyk and Kimble, 1998).


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However, if the activity of at least one gene in each pathway is eliminated, a synthetic tumorous germ line phenotype is observed (Eckmann et al., 2004; Hansen et al., 2004b; Kadyk and Kimble, 1998). This tumor is the result of a failure of germ cells to initiate meiotic development at the appropriate time/ place and is similar to the germ line tumor formed in animals carrying a strong glp‐1 gain‐of‐function allele (Eckmann et al., 2004; Francis et al., 1995b; Hansen et al., 2004a,b; Kadyk and Kimble, 1998). The regulatory relationship between GLP‐1 signaling and the GLD‐1 and GLD‐2 pathways was determined from genetic epistasis experiments; the gld‐1 gld‐2 double null meiotic entry failure is epistatic to the premature meiotic entry defect of glp‐1 null as the triple mutant has a tumorous germ line (Kadyk and Kimble, 1998). Thus GLP‐1 Notch signaling promotes germ line proliferation by inhibiting the meiosis‐promoting activities of the GLD‐1 and GLD‐2 pathways (Fig. 2). GLD‐1 and GLD‐2 pathway genes have additional, nonoverlapping, functions in germ line development. GLD‐1 promotes the male sexual fate in the hermaphrodite germ line and is required for meiotic prophase progression (Francis et al., 1995a,b). gld‐1 null single mutants have a tumorous germ line; however, this overproliferation is not due to a meiotic entry failure but rather is the result of pachytene germ cells returning to a mitotic cell cycle. NOS‐3 functions to promote the female germ line sexual fate, opposite to the sex‐determining function of GLD‐1, but does not have an essential function in meiotic prophase progression (Kraemer et al., 1999). GLD‐2 is necessary for meiotic progression/gametogenesis in both hermaphrodites and males (a diplotene‐like arrest occurs in a gld‐2 null mutant) but has no essential function in germ line sex determination (Kadyk and Kimble, 1998). GLD‐3 promotes the male sexual fate in the germ line and also plays a role in oogenesis (Eckmann et al., 2002). In meiotic prophase progression, GLD‐2 and GLD‐3 function redundantly rather than interdependently as they do in regulating meiotic entry (Eckmann et al., 2002). Thus the meiotic entry mRNA regulators must have additional targets on which they uniquely act. How might the GLD‐1 and GLD‐2 mRNA regulatory pathways function redundantly to promote initiation of meiotic development? For all GLD‐1 target RNAs studied thus far, which are primarily involved in GLD‐1’s roles in sex determination and meiotic prophase progression during oogenesis, GLD‐1 functions as a translational repressor (Jan et al., 1999; Lakiza et al., 2005; Lee and Schedl, 2001, 2004; Marin and Evans, 2003; Xu et al., 2001). Thus GLD‐1 likely promotes meiotic entry by translational repression of proliferation‐promoting genes. The GLD‐2/GLD‐3 complex likely promotes gene activity; lengthening poly(A) tails both stabilizes and promotes the translation of mRNAs (Wang et al., 2002). Thus, the GLD‐2/GLD‐3 multisubunit poly(A) polymerase likely promotes meiotic entry by increasing the

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amount of meiotic entry gene products through increased translation— either directly or indirectly. However, the model of GLD‐1 repressing proliferation‐promoting genes and GLD‐2/GLD‐3 activating meiotic entry gene products cannot be as simple as stated. If so, then one would not expect the essentially normal pattern of proliferation–meiotic entry observed in the gld‐1, gld‐2, or gld‐3 null single mutants. One possibility is that GLD‐1 and GLD‐2/GLD‐3 do not have unitary molecular functions. Alternatively, there may be additional pathways that act in parallel to the GLD‐1 and GLD‐2 pathways (see later). An important unanswered question is what are the mRNA targets of both GLD‐1 and GLD‐2/GLD‐3 related to their activities in regulating the proliferation–meiotic entry decision?

IV. Control of the Adult Pattern of Proliferation and Meiotic Entry A. Stem Cells Versus Transit Amplifying Cells The adult proliferative zone contains 200–250 germ cells (Killian and Hubbard, 2005; Lamont et al., 2004). It is unknown if the entire population of proliferative cells is homogeneous or if there are diVerent cell types within the population. One possibility (Fig. 4, model 1) is that all of the proliferative cells are stem cells, in that they all are capable of dividing to give both self‐ renewing and diVerentiating daughter cells. Another possibility (Fig. 4, model 2) is that only the distal‐most cells in the proliferative zone are stem cells and more proximal cells have ‘‘committed’’ to diVerentiate but undergo one or more proliferative cycles before they enter meiotic prophase (transit amplifying cells). The second model of stem cells followed by transit amplifying cells would be similar to what is observed in other systems, such as in the testes of male mammals and in both the male and the female Drosophila germ lines (McLean, 2005; Yamashita et al., 2005). In mammalian males, there are relatively few spermatogonial stem cells in the population of proliferating germ cells. DiVerentiating proliferative cells divide through spermatocytogenesis to amplify the number of cells that go on to enter meiotic prophase and diVerentiate as sperm (McLean, 2005). This amplification step is needed in order to produce the large number of sperm that are necessary for reproductive fitness. In Drosophila males, only approximately nine stem cells exist at the apical tip of the testis (Wong et al., 2005). As these cells divide asymmetrically, one daughter cell maintains its stem cell character (self‐renews), and the other daughter cell, the gonialblast, initiates diVerentiation. As a stereotypic part of the diVerentiation program, the gonialblast undergoes four rounds of mitotic divisions before overt diVerentiation.


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Therefore, the majority of proliferating cells in the Drosophila testis are not stem cells but rather transit amplifying cells. The stem cells can be identified in this population based on their proximity to certain somatic cells and the availability of markers (Wong et al., 2005). In C. elegans, unlike Drosophila, there are currently no obvious cytological diVerences between cells in the proliferative region for both the hermaphrodite and male germ lines. Therefore, it is unknown if these cells are equivalent, or if only a small number are actual stem cells. A key experiment to distinguish between these two models would be to isolate cells from diVerent regions of the proliferative zone and test their ability to repopulate a gonad arm that has been depleted of germ cells. To date, no repopulation assays have been performed with the C. elegans germ line. Repopulation assays are technically challenging because the proliferative cells of the germ line are not completely cellularized (Hall et al., 1999; Hirsh et al., 1976) (Fig. 1 legend). Therefore, individual cells, or even small groups of cells, cannot be isolated and transplanted to a diVerent gonad. Experiments, however, suggest that the cycling behavior of cells in the proliferative zone is nonhomogeneous. Cells within the first two to three cell diameters from the DTC have a lower rate of cell division than cells further away from the distal end. Cells approximately six to seven cell diameters from the DTC have the highest frequency of cell division (Maciejowski et al., 2006). In some stem cell systems, the niche‐associated stem cells cycle more slowly than transit amplifying cells (Cotsarelis et al., 1990). Thus, the adult C. elegans germ line may be analogous to other systems in which the stem cells cycle more slowly; the DTC niche‐associated cells within the first two to three cell diameters from the distal end being the stem cells and the other proliferative cells being transit amplifying (also see later).

B. Spatial Control of Notch Signaling An understanding of the spatial control of the proliferative zone is hindered by our lack of knowledge regarding precisely what cells have active GLP‐1 Notch signaling. On binding of LAG‐2 to the GLP‐1 receptor, GLP‐1 (INTRA) is generated; it translocates to the nucleus and forms a ternary complex with the LAG‐1 transcription factor and the SEL‐8/LAG‐3 coactivator. This leads to transcription of downstream genes. However, in which nuclei along the distal–proximal axis of the proliferative zone the ternary complex is present and active is unknown. The Notch intracellular domain has been diYcult to detect cytologically in any system, in part due to its low abundance and due to masking of the antigen (Cheng et al., 2003; Mumm and Kopan, 2000). GLP‐1 in its receptor form has been detected on the surface of the germ cells in C. elegans, and is expressed at its highest level in

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Figure 4 Models relating two possible spatial patterns of GLP‐1 Notch signaling with two possible behaviors of cells within the proliferative zone. Included in each model is a diagram of the distal end of an adult hermaphrodite gonad arm; surface view. The diagram illustrates the relative positions of the DTC (yellow), stem cells (green), transit amplifying cells (blue), and cells that have just entered meiotic prophase (red). Below each diagram is a graph showing the known abundance of the GLP‐1 receptor on the surface of the germ cells (black) (Crittenden


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the first 20 cell diameters of the distal end and at a lower level more proximally (Crittenden et al., 1994) (Fig. 4). This does not, however, mean that GLP‐1 Notch signaling is active over this entire region. In gld‐1 null mutant animals meiotic entry is relatively normal even though the GLP‐1 receptor is expressed at the surface of cells throughout the germ line (Crittenden et al., 1994; Marin and Evans, 2003). Normally, glp‐1 translation is inhibited outside of the proliferative zone by GLD‐1 through a negative feedback loop (Marin and Evans, 2003). Since meiotic entry is essentially normal in gld‐1 null mutants with high GLP‐1 receptor levels throughout the gonad, this demonstrates that surface expression of GLP‐1 does not necessarily correlate with the presence of active GLP‐1(INTRA)– LAG‐1–SEL‐8/LAG‐3 ternary complexes within the cell. The spatial distribution of GLP‐1(INTRA)–LAG‐1–SEL‐8/LAG‐3 ternary complexes can be incorporated into the two models for control of the proliferative zone (Fig. 4). In model 1, the active GLP‐1 Notch signaling ternary complex is found throughout the proliferative zone, with higher levels at the distal end and lower levels near the transition zone. The continuing presence of the ternary complex would lead to GLP‐1 Notch signaling‐dependent proliferation throughout the zone. At some point a threshold would be crossed where low GLP‐1 Notch signaling and high GLD‐1 and GLD‐2 pathway activity would lead to meiotic entry. In this model, the proliferative zone would primarily contain self‐renewing stem cells. In model 2, the active GLP‐1 Notch signaling ternary complex is found only in the distal‐most cells. In this case, stem cells only exist close to the DTC where GLP‐1 Notch signaling is active. The remaining proliferative cells undergo transit amplification via a mechanism that is not directly promoted by GLP‐1 Notch signaling. The switch from transit amplification to meiotic entry would depend on when a threshold activity of the GLD‐1 and GLD‐2 pathways, which were initially repressed by GLP‐1 Notch signaling, is finally achieved. The size and shape of the DTC presumably influences what cells have active GLP‐1 Notch signaling since the LAG‐2 ligand is bound to the DTC membrane. The DTC caps the very distal end of the gonad and has tentacle‐like et al., 1994) and GLD‐1 protein (red) (Hansen et al., 2004b; Jones et al., 1996) and two possible spatial patterns of the GLP‐1(INTRA)–LAG‐1–SEL‐8/LAG‐3 ternary complex (green), which depicts active GLP‐1 Notch signaling. In the first model, all of the proliferative cells are a ‘‘stem cell population’’ capable of self‐renewal. GLP‐1 Notch signaling is active throughout the proliferative zone, progressing from high to low as cells move proximally. This pattern of activity may be the inverse of GLD‐1 accumulation, since GLP‐1 Notch signaling inhibits GLD‐1 accumulation. In the second model, GLP‐1 Notch signaling is only active in the cells at the very distal end of the gonad, where LAG‐2 ligand is present. Only the distal‐most cells with active GLP‐1 Notch signaling are stem cells while the more proximal cycling cells are transit amplifying cells that are not capable of indefinite self‐renewal. These models are extreme cases; the true situation may be somewhere in between. See text for details.

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projections that extend, on average, approximately eight cell diameters from the distal end (Hall et al., 1999). Each cell only has a few of these projections and they are quite thin; thus, only a minority of cells in the proliferative zone, even relatively close to the DTC, actually come in contact with LAG‐2 ligand. Therefore, only cells near the distal end are likely to have GLP‐1 (INTRA) entering the nucleus, which would appear to be consistent with model 2. However, it is unknown how rapidly the GLP‐1(INTRA)–LAG‐1– SEL‐8/LAG‐3 ternary complexes decay; it may be over one or a few additional cell cycles, consistent with model 1, or very rapidly, consistent with model 2. One regulator that promotes ternary complex decay is SEL‐10, an F‐box member of the SCF E3 ubiquitin ligase complex involved in proteasome‐mediated degradation of target proteins (Hubbard et al., 1997; Spruck and Strohmaier, 2002). Both C. elegans and mammalian SEL‐10 bind to the intracellular portion of the Notch receptor, leading to the model that SEL‐10 negatively regulates Notch signaling by targeting GLP‐1(INTRA) Notch for degradation by the proteasome (Fig. 4) (Gupta‐Rossi et al., 2001; Hubbard et al., 1997; Oberg et al., 2001). However, sel‐10 mutants have not aided in determining the importance of GLP‐1(INTRA) degradation in regulating GLP‐1 Notch signaling in the proliferative zone because sel‐10 null mutants are wild type. However, genetic data indicate that SEL‐10 must function redundantly with another factor to promote GLP‐1(INTRA) degradation. Redundancy is inferred from sel‐10 null mutants being wild type while a novel glp‐1 gain‐of‐function allele (q35), which should stabilize GLP‐ 1(INTRA) due to a C‐terminal deletion of the PEST sequences implicated in degradation of Notch homologs, has a multivulval phenotype (Hubbard et al., 1997; Jager et al., 2004; Mango et al., 1991). Thus, much remains to be learned about the spatial pattern of the GLP‐1 Notch signaling ternary complex and genes that mediate its decay.

V. Regulation of the Activity of the GLD‐1 and GLD‐2 Pathways A. GLP‐1 Notch Signaling Control of GLD‐1 and GLD‐2 Pathway Activity The GLP‐1 Notch signaling pathway represses the GLD‐1 and GLD‐2 pathways in the distal‐most germ line. As cells move proximally, GLP‐1 Notch signaling is reduced allowing the GLD‐1 and GLD‐2 pathways to promote meiotic entry and/or inhibit proliferation (Fig. 2). Since the activity of either the GLD‐1 or GLD‐2 pathway is suYcient to induce meiotic entry, both pathways must be simultaneously repressed to achieve proliferation. GLD‐1 appears to be regulated primarily through expression. GLD‐1 levels are very


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low at the distal‐most end and then progressively rise to a high level 20 cell diameters from the distal end—the position where germ cells enter meiotic prophase (Hansen et al., 2004b; Jones et al., 1996) (Fig. 5). The rise in GLD‐1 level is roughly 20‐fold (Hansen et al., 2004b). Low GLD‐1 levels at the distal‐most end appear to be necessary for these cells to remain proliferative while high GLD‐1 levels appear to be suYcient for meiotic entry. This is suggested by a gld‐1 gain‐of‐function allele with higher distal GLD‐1 levels, which causes distal germ cells to prematurely initiate meiotic

Figure 5 Factors controlling GLD‐1 accumulation in the distal germ line. Top panel shows two fluorescent photomicrographs of the same distal gonad of an adult hermaphrodite, with DNA morphology from DAPI staining (blue) and GLD‐1 protein levels from staining with anti‐GLD‐1 antibodies (red); surface view. The bottom panel is a graph showing the relative accumulation pattern of GLD‐1 protein (red line) in adult hermaphrodites (Hansen et al., 2004b). The factors controlling GLD‐1 accumulation are shown with inhibitory influences depicted by a bar and promoting influences depicted with an arrow. GLP‐1 Notch signaling inhibits GLD‐1 accumulation, partly through FBF and partly through an unknown factor(s). The GLD‐2/GLD‐3 complex promotes GLD‐1 accumulation redundantly with NOS‐3. GLD‐1 levels are low in the distal end where GLP‐1 Notch signaling is highest. GLD‐1 levels are higher more proximally where GLP‐1 Notch signaling is reduced, where FBF is no longer inhibiting and where GLD‐2/GLD‐3 and NOS‐3 are free to promote GLD‐1 accumulation. See text for details.

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development (Hansen et al., 2004b; Jones et al., 1996). GLP‐1 Notch signaling inhibits the accumulation of GLD‐1 in the distal end; GLD‐1 levels are elevated in the distal end of animals that lack glp‐1 activity (Hansen et al., 2004b). This inhibition of GLD‐1 accumulation is posttranscriptional as gld‐1 mRNA levels are equivalent in the glp‐1(þ) and glp‐1() backgrounds employed (Hansen et al., 2004b). Regulation of NOS‐3, GLD‐2, and GLD‐3 in the adult appears to be, in large part, posttranslational. NOS‐3 levels are constant throughout the entire distal arm of the germ line (Kraemer et al., 1999). GLD‐2 protein is found throughout the germ line, but with moderately lower levels in the proliferative zone (Wang et al., 2002). GLD‐3 levels are highest in the transition zone, flanked by slightly lower levels in the proliferative zone and pachytene region (Eckmann et al., 2002). The increase in GLD‐2 and GLD‐3 levels as germ cells progress from the proliferative to the transition zone is only a few fold, in contrast to the 20‐fold increase in GLD‐1 levels, and thus would seem unlikely to be suYcient to cause a switch from proliferation to initiation of meiotic development. The premature meiotic entry phenotype from loss of GLP‐1 Notch signaling is most likely due to the ectopic activity of the GLD‐1 and GLD‐2 pathways in the distal end. Since GLP‐1 Notch signaling culminates in GLP‐1 (INTRA)–LAG‐1–SEL‐8/LAG‐3 complexes that are thought to transcriptionally activate genes, and since gld‐1, gld‐2, nos‐3, and gld‐3 appear not to be transcriptionally regulated in the distal end, inhibition of the GLD‐1 and GLD‐2 pathways by GLP‐1 Notch signaling must be indirect. Instead, GLP‐1 Notch signaling likely activates the transcription of genes that, directly or indirectly, inhibit the function of GLD‐1 and GLD‐2 pathway components.

B. FBF Functions Between GLP‐1 Notch Signaling and the GLD‐1 and GLD‐2 Pathways One pair of genes that function between GLP‐1 Notch signaling and the redundant GLD‐1 and GLD‐2 pathways is fbf‐1 and fbf‐2 (fem‐3 binding factor) (Crittenden et al., 2002; Lamont et al., 2004) (Figs. 5 and 6). These two genes encode nearly identical proteins that are homologous to the Drosophila translational regulator Pumilio (Zhang et al., 1997). FBF‐1 and FBF‐2, collectively referred to as FBF, are largely redundant in their control of germ line proliferation. In mutants lacking FBF activity, proliferative germ cells enter meiosis prematurely. By the end of the L4 stage, at 20 C, all proliferative cells have entered meiotic prophase (Crittenden et al., 2002). The fbf‐2 promoter region contains four LAG‐1‐binding sites and its expression is altered in glp‐1 mutants; therefore, the activity of fbf‐2 is perhaps


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Figure 6 Additional genetic complexity regulating the proliferation–meiotic entry decision. Superimposed on the core regulatory network (Fig. 2) are additional regulatory interactions, primarily inferred from genetic analysis (see text for details). Positive interactions are shown with arrows while negative or inhibitory interactions are shown with bars. GLP‐1 Notch signaling promotes the activities of FBF, FOG‐1, and unknown factor(s) X. Currently, fbf‐2 is the only known transcriptional target of GLP‐1 Notch signaling that promotes proliferation, others almost certainly exist. FOG‐3 and FEM‐3 may also function with FOG‐1; however, since their analysis is less complete they have not been included. FBF, FOG‐1, and X then inhibit the activities of the GLD‐1 and GLD‐2 downstream pathways. In the case of FBF, gld‐1 and gld‐3 mRNAs are known targets of this repression. A third pathway is inferred to function in parallel with the GLD‐1 and GLD‐2 pathways to inhibit proliferation and/or promote entry into meiotic prophase. Genes that uniquely define the third meiotic entry pathway are not known. It is unclear whether the third pathway is inhibited by FBF, FOG‐1, and/or X or if inhibition is independent of these factors. The GLD‐1 pathway contains NOS‐3 and GLD‐1. NOS‐3 functions upstream of GLD‐1, promoting its accumulation. GLD‐1 likely represses the translation of downstream target mRNAs that are necessary for proliferation; these targets remain to be identified. The GLD‐2 pathway consists of GLD‐2 and GLD‐3 and, inferred from genetic data, it is proposed that FBF is also part of the GLD‐2 pathway. GLD‐2 and GLD‐3 perhaps increase the stability or translatability of the mRNA targets that promote meiotic entry; these targets remain to be identified. The GLD‐2/GLD‐3 complex either directly or indirectly promotes the accumulation of GLD‐1 protein, functioning redundantly with NOS‐3. How FBF acts both to promote proliferation by repressing the meiotic entry pathways and to promote meiotic development by acting in the GLD‐2 pathway is unclear. ATX‐2 promotes proliferation independent of GLP‐1 Notch signaling and is shown acting in parallel, although not redundantly, with GLP‐1 Notch signaling. However, it is possible that ATX‐2 acts downstream in opposition of the targets of one or more of the three meiotic entry pathways. A number of the regulatory interactions shown are context (developmental stage, sex, or growth temperature) specific. For example, the third meiotic entry pathway appears not to function during larval development, FOG‐1 appears not to be required to promote proliferation in hermaphrodites undergoing oogenesis and FBF appears not to be essential in animals grown at 25 C.

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directly regulated by GLP‐1 Notch signaling (Lamont et al., 2004). The FBF protein binds to the GLD‐1 30 untranslated region (UTR) in vitro, and GLD‐1 protein levels increase in the distal end of fbf‐1 single mutants (Crittenden et al., 2002). Furthermore, the premature meiotic entry phenotype in fbf mutants is suppressed in animals heterozygous for a gld‐1 null allele. Together these results demonstrate that FBF inhibits the accumulation of the GLD‐1 protein. They further suggest that the premature meiotic entry defect in fbf mutant animals is caused, at least in part, by increased GLD‐1 activity in the distal end. Therefore, by reducing GLD‐1 levels by half in a gld‐1 null heterozygote, GLD‐1 activity is lowered in the distal end enough to allow for proliferation. All of this supports the model that FBF is a direct link between GLP‐1 Notch signaling and the activity of the GLD‐1 pathway (Fig. 6). Genetic data indicate that nos‐3 and fbf act in opposition, with nos‐3 possibly functioning in between fbf and gld‐1 (Hansen et al., 2004b). nos‐3 mutations suppress the premature meiotic entry defect in fbf mutants, suggesting that NOS‐3 functions downstream of FBF (however, see later). Expression analysis places NOS‐3 upstream of GLD‐1, promoting the expression of GLD‐1 (Hansen et al., 2004b) (Fig. 5; see later). Therefore, FBF and NOS‐3 appear to work in opposition in controlling GLD‐1 accumulation; FBF inhibiting GLD‐1 accumulation, and NOS‐3 promoting accumulation (Fig. 5). FBF and NOS‐3 physically interact in vitro (Kraemer et al., 1999). Both fbf and nos‐3 also regulate the switch from spermatogenesis to oogenesis in the hermaphrodite, but unlike their relationship in regulating proliferation in which they work in opposite directions, in controlling sex determination they work in the same direction, promoting the female fate (Kraemer et al., 1999; Zhang et al., 1997). FBF appears to regulate the activity of the gld‐2 pathway through control of GLD‐3 levels. FBF binds to the 30 UTR of gld‐3 and GLD‐3 levels increase in larval fbf‐1 fbf‐2 double‐mutant males (Eckmann et al., 2004). However, it is unclear how important this regulation is in the adult, where the increase in GLD‐3 levels as germ cells progress from the proliferative to the transition zone is only a few fold (Eckmann, 2004). FBF thus appears to inhibit the GLD‐2 pathway by repressing GLD‐3 accumulation, but this regulation may be restricted to larvae (Fig. 6). FBF is the first identified direct link between GLP‐1 Notch signaling and the downstream GLD‐1 and GLD‐2 pathways. However, genetic data indicate that FBF cannot be the only link inhibiting these two pathways. With respect to the GLD‐1 pathway, FBF cannot be the only transcriptional target of GLP‐1 Notch signaling that inhibits GLD‐1 accumulation. First, it must be remembered that the GLD‐1 pathway is suYcient for cells to enter meiosis (meiotic entry is normal in a gld‐2 mutant); therefore, ectopic GLD‐1 activity in the distal end should cause a strong premature entry into


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meiosis phenotype. Ectopic GLD‐1 activity is presumed to be the cause of premature meiotic entry in a gld‐2; glp‐1 double mutant (Kadyk and Kimble, 1998). Premature meiotic entry in fbf mutants occurs much later than in glp‐1 or gld‐2; glp‐1 mutants; only 16 sperm are formed in each gonad arm of a glp‐1 mutant and 64 in a gld‐2; glp‐1 mutant, while 200 sperm are formed in each gonad arm of an fbf mutant (Austin and Kimble, 1987; Crittenden et al., 2002; Kadyk and Kimble, 1998). Second, in fbf mutants grown at 25 C, the proliferative population is never fully depleted in adult animals, demonstrating that fbf activity is not required to maintain proliferation in the adult at 25 C. glp‐1 activity is required to maintain proliferation in adults at 25 C (Austin and Kimble, 1987). Since high GLD‐1 promotes meiotic entry and GLP‐1 Notch signaling is inducing proliferation by repressing GLD‐1 in larvae and adults at all temperatures, GLP‐1 Notch signaling is perhaps promoting the transcription of another factor (X, in Fig. 5) that acts directly or indirectly to repress GLD‐1 accumulation, at least in larvae at 20 C and adults at 25 C when FBF is not essential.

C. FOG‐1, FOG‐3, and FEM‐3 Function Redundantly with FBF As mentioned earlier, other factors must work with FBF in inhibiting the downstream pathways. FOG‐1, FOG‐3, and/or FEM‐3 may act redundantly with FBF in promoting proliferation (Fig. 6) (Thompson et al., 2005). FOG‐1, FOG‐3, and FEM‐3 also are involved in regulating sex determination, with FOG‐1 and FOG‐3 promoting the male fate in the germ line, and FEM‐3 promoting the male fate in the germ line and soma (Ellis and Schedl, 2006). FOG‐1 is a cytoplasmic polyadenylation element‐binding (CPEB) protein, FOG‐3 belongs to the Tob family of proteins, and FEM‐3 is novel (Ahringer et al., 1992; Chen et al., 2000; Jin et al., 2001; Luitjens et al., 2000). Mutations in each of fog‐1, fog‐3, and fem‐3 enhance the larval fbf premature meiotic entry phenotype suggesting that they function redundantly with fbf in promoting proliferation, although most analysis has been performed with fog‐1 (Thompson et al., 2005). Surprisingly, in contrast to the enhancement of the fbf premature meiotic entry phenotype in fog‐1 null homozygotes, fog‐1 null heterozygotes suppress the premature meiotic entry phenotype, suggesting that FOG‐1 function may switch in a concentration‐dependent manner (Thompson et al., 2005). In germ lines undergoing spermatogenesis (early larval hermaphrodites and males) FOG‐1 protein is detected at low levels in the proliferative region that rises as cells enter meiotic prophase; FOG‐1 protein is not detected in adult hermaphrodite germ lines undergoing oogenesis (Thompson et al., 2005). These results suggest that FOG‐1’s function to promote proliferation may be limited to early larvae and males. FBF was found to inhibit FOG‐1 accumulation in early larvae (Thompson et al., 2005).

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The studies of FBF and FOG‐1 thus indicate that they function redundantly downstream of GLP‐1 to inhibit proliferation in larvae. However, in adult hermaphrodites at 25 C, there must be another factor that functions redundantly with FBF to promote proliferation.

D. FBF‐1 and FBF‐2 Functions Are not Completely Redundant Although fbf‐1 and fbf‐2 are largely redundant, they do have some unique functions. For example, only fbf‐2 has LAG‐1–binding sites; fbf‐1 is perhaps not directly regulated by GLP‐1 Notch signaling (Lamont et al., 2004). FBF‐1 and FBF‐2 protein expression patterns support the model that fbf‐2 is directly regulated by Notch signaling, while fbf‐1 is not. FBF‐2 is expressed at its highest level in the distal end of the germ line where GLP‐1 Notch signaling presumably is most active (Lamont et al., 2004). FBF‐1 expression is lower in the first six cell diameters of the distal end and highest from 6 to 20 cell diameters (Crittenden et al., 2002; Lamont et al., 2004). If GLP‐1 Notch signaling transcriptionally activated fbf‐1, its protein levels should not be lower in the first six cell diameters where GLP‐1 Notch signaling is thought to be at its highest. fbf‐1 and fbf‐2 appear to regulate each other through their 30 UTRs and are thought to fine‐tune the size of the proliferative zone (Lamont et al., 2004).

VI. Additional Components and Pathways Regulating the Proliferation–Meiotic Entry Decision A. FBF May also Function in the GLD‐2 Pathway A major surprise comes from genetic data suggesting that FBF may act in the GLD‐2 pathway to promote meiotic entry (Fig. 6). While both GLP‐1 and FBF promote proliferation, they display the opposite interaction with gld‐1 null. In gld‐1; glp‐1 null double mutants, germ cells prematurely enter meiosis (Francis et al., 1995b). By contrast, gld‐1; fbf‐1 fbf‐2 null triple mutants are tumorous due to a defect in meiotic entry (Crittenden et al., 2002). This surprising result can be rationalized based on what we know about the core proliferation–meiotic entry network (Fig. 2). gld‐1; glp‐1 null double mutants prematurely enter meiosis because of high GLD‐2 pathway activity in the absence of glp‐1 activity. Similarly, in the gld‐1; fbf‐1 fbf‐2 null triple mutant, one would predict high GLD‐2 pathway activity in the absence of FBF, the mediator of GLP‐1 Notch signaling, which should then result in premature meiotic entry. Even if FBF is not the sole inhibitor of the


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GLD‐2 pathway, there should at least be normal GLD‐2 pathway activity near the transition zone resulting in normal meiotic entry, as is observed in the gld‐1 null mutant. Instead, the gld‐1; fbf‐1 fbf‐2 null triple mutant has a tumorous phenotype like the gld‐1 gld‐2 or gld‐1; gld‐3 null double mutants, suggesting that the absence of fbf‐1 fbf‐2 is similar/equivalent to the absence of gld‐2 or gld‐3. Thus, the simplest interpretation is that FBF functions in the GLD‐2 pathway to promote meiotic entry (Fig. 6). Based on previous studies (Hansen et al., 2004b; Kadyk and Kimble, 1998), if FBF functions in the GLD‐2 pathway, then there are two predicted genetic interactions for fbf mutations: 1. fbf‐1 fbf‐2 null should be synthetic tumorous with nos‐3 null. While the fbf‐1 fbf‐2; nos‐3 null mutant is not tumorous, the premature meiotic entry phenotype is suppressed such that adults have a proliferative zone followed by meiotic entry at 20 C (Hansen et al., 2004b). [The fbf‐1 fbf‐2; nos‐3 null triple mutant may not be completely tumorous because either nos‐3 or fbf mutants, or both, may not completely eliminate the activity of their respective pathways. This possibility has previously been suggested for nos‐3 (Hansen et al., 2004b).] While this suppression was previously interpreted as NOS‐3 functioning downstream of FBF in the regulation of GLD‐1 accumulation, it can also be interpreted as a synthetic interaction from aVecting both the GLD‐1 and GLD‐2 pathways. 2. fbf‐1 fbf‐2 should not be synthetic tumorous with gld‐2 or gld‐3. The gld‐2; fbf‐1 fbf‐2 and fbf‐1 fbf‐2; gld‐3 null mutants display the premature meiotic entry phenotype, consistent with the three genes acting in the same pathway. If the above‐mentioned interpretation of the genetic results is correct, then how can FBF act to both promote proliferation and promote meiotic entry? One possibility is based on the finding that FBF‐1 and FBF‐2 are not fully equivalent and that the combination of both transcriptional control and cross‐regulation of FBF‐1 by FBF‐2, and vice versa, could lead to a switch in the aggregate behavior of FBF‐1 FBF‐2 (Lamont et al., 2004). Another possibility is that the activity of FBF or its mRNA targets might change as germ cells progress proximally because of alterations in binding partners or alterations in protein modifications.

B. Cross Talk Between the GLD‐1 and GLD‐2 Pathways The GLD‐1 and GLD‐2 pathways do not function completely independent of one another. GLD‐2 and GLD‐3 were found to promote the expression of GLD‐1 protein, functioning redundantly with NOS‐3 (Figs. 5 and 6) (Hansen et al., 2004b). The gld‐2; nos‐3 and the gld‐3; nos‐3 null double

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mutants have very low levels of GLD‐1 throughout the germ line compared to wild type, while the single mutants show a similar pattern to wild type. It should be noted that promoting GLD‐1 accumulation cannot be the only function of GLD‐2/GLD‐3 in promoting meiotic entry because the phenotype of the gld‐1, gld‐2, and gld‐3 null single mutants are diVerent from each other and from the gld‐1 gld‐2 and gld‐1; gld‐3 null doubles. Thus GLD‐2/ GLD‐3 promote GLD‐1 protein accumulation and separately promote meiotic entry (Fig. 6).

C. A Third Pathway Downstream of Notch Signaling If the GLD‐1 and the GLD‐2 pathways are the only meiotic entry promoting activities downstream of GLP‐1 Notch signaling, then the expectation is that the phenotype of full activation of GLP‐1 Notch signaling should be equivalent to elimination of the GLD‐1 and GLD‐2 pathways that it represses. Animals carrying two copies of the strong glp‐1 gain‐of‐function allele oz112, and also carrying a wild‐type copy of glp‐1 on a free duplication, have completely tumorous germ lines with no evidence of cells entering meiotic prophase (Berry et al., 1997; Hansen et al., 2004a). However, in gld‐1 gld‐2 null mutant adults, even though their germ lines are tumorous, some cells have entered meiotic prophase (Hansen et al., 2004a). The nuclei in these cells stain for the meiotic chromosome axis protein HIM‐3, whereas in the strong glp‐1 gain‐of‐function mutants no HIM‐3 positive nuclei are detected (Figs. 2 and 3) (Hansen et al., 2004a; Zetka et al., 1999). These results indicate that there must be an additional activity(s) promoting meiotic entry in the absence of the GLD‐1 and GLD‐2 pathways. This third pathway must be downstream of GLP‐1 Notch signaling because it must be inhibited in glp‐1(oz112gf) mutant animals. The third pathway is envisioned to act in parallel to the GLD‐1 and GLD‐2 pathways (Fig. 6) (Hansen et al., 2004a). At this point no genes are known that uniquely define the third pathway. Additionally, it is possible that the third pathway is only important in the adult, as all germ cells are proliferative in gld‐1 gld‐2 null mutant larvae (Hansen et al., 2004a).

D. MOG Gene Products Act to Promote Meiotic Entry The MOG‐1, MOG‐4, MOG‐5, and MOG‐6 proteins, originally identified based on their roles in promoting the female fate in the hermaphrodite germ line, are also involved in regulating the proliferation–meiotic entry decision (Belfiore et al., 2004; Graham and Kimble, 1993; Graham et al., 1993). A mutation in any one of the mog genes in animals that have reduced gld‐3


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activity causes a synthetic tumorous phenotype (Belfiore et al., 2004). This tumorous phenotype is due to a meiotic entry defect and is epistatic to complete loss of glp‐1 activity, suggesting that the MOGs function downstream of GLP‐1 Notch signaling (M. Hanazawa and T. Schedl, unpublished observations). The MOG proteins perhaps function in the GLD‐1 pathway based on the synthetic tumorous phenotype of the mog; gld‐2 and mog; gld‐3 double mutants. However, they must also function elsewhere, possibly in the third pathway, because gld‐1; mog double mutants also show synthetic overproliferation phenotypes (M. Hanazawa and T. Schedl, unpublished observations). MOG‐6 is a cyclophilin homolog (Belfiore et al., 2004). MOG‐1, MOG‐4, and MOG‐5 are DEAH box RNA helicases that are orthologous to yeast splicing factors PRP16, PRP2, and PRP22, respectively (Puoti and Kimble, 1999, 2000). Therefore, the MOG proteins perhaps function in mRNA splicing or some other aspects of mRNA metabolism. How the MOG proteins aVect mRNA metabolism to promote meiotic entry remains to be determined.

E. ATX‐2 May Promote Proliferation Independent of GLP‐1 Notch Signaling All factors discussed thus far are thought to regulate the proliferation– meiotic entry decision by either controlling, or being controlled by, GLP‐1 Notch signaling. However, atx‐2 appears not to fall into either of these categories; it may promote proliferation, at least in part, independent of GLP‐1 Notch signaling (Fig. 6) (Maine et al., 2004). ATX‐2 encodes a protein homologous to the mammalian ataxin‐2 protein, which is thought to be involved in RNA metabolism (Kiehl et al., 2000; Shibata et al., 2000). Animals with reduced ATX‐2 activity have a reduced number of cycling cells in the proliferative zone, suggesting that ATX‐2’s normal activity is to promote proliferation (Ciosk et al., 2004; Maine et al., 2004). The gld‐1 gld‐2 null tumorous phenotype is partially suppressed by atx‐2(RNAi) knockdown; a substantial number of germ cells enter meiotic prophase compared to the gld‐1 gld‐2 null alone (Maine et al., 2004). This result suggests that ATX‐2 may act downstream of the GLD‐1 and/or GLD‐2 pathways, may act downstream of GLP‐1 Notch signaling to repress the third meiotic entry pathway, or may act in parallel to GLP‐1 Notch signaling. atx‐2(RNAi) also substantially suppresses the gld‐1 gld‐2 tumorous phenotype in the absence of glp‐1 activity [in gld‐1 gld‐2; glp‐1 atx‐2(RNAi) mutants most germ cells have entered meiosis] indicating that ATX‐2 can not act downstream of GLP‐1 Notch signaling to repress the third meiotic entry pathway (Maine et al., 2004). atx‐2(RNAi) knockdown did not suppress the tumorous phenotype caused by constitutive activation of the strong

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glp‐1(oz112) gain‐of‐function mutant, where the activities of the GLD‐1, GLD‐2, and third pathways should be fully repressed (Maine et al., 2004). Together, these results suggest that ATX‐2 promotes proliferation independent of GLP‐1 Notch signaling, acting in parallel. However, atx‐2(RNAi) knockdown may have been ineYcient in the glp‐1 gain‐of‐function condition leaving open the possibility that ATX‐2 promotes proliferation by opposing the downstream meiotic entry output from the GLD‐1 and/or GLD‐2 pathways. In addition to its role in promoting proliferation, ATX‐2 also functions to promote the female fate in the hermaphrodite germ line (Ciosk et al., 2004; Maine et al., 2004). F. Connection Between the Proliferation–Meiotic Development Decision and Germ Line Sex Determination Many of the genes that function in the proliferation–meiotic entry decision, with the exception of the canonical GLP‐1 Notch cascade, also function in the germ line sex determination pathway (Ellis and Schedl, 2006). However, gene products that promote proliferation can either specify the female fate (e.g., ATX‐2) or the male fate (e.g., FOG‐1), and genes that promote meiotic entry can either specify the female fate (e.g., NOS‐3) or the male fate (e.g., GLD‐1). Thus, it is unclear if there is an intimate connection between the two decisions or if these genes, because they encode mRNA regulators that are likely to have many targets with diverse function, are just pleiotropic.

VII. Conclusions In the reproductive strategy of a given species, control of the proliferation– meiotic entry decision is important for the production of a suYcient number of gametes at the appropriate times. In C. elegans, a large population of proliferative cells is maintained and gametes are produced throughout adulthood. The balance between proliferation and meiotic entry is controlled spatially with cells close to the DTC proliferating and cells further away entering and then progressing through meiotic prophase. GLP‐1 Notch signaling polarizes the germ line, with high levels of signaling at the distal end, near the DTC. GLP‐1 Notch signaling promotes proliferation by inhibiting the redundant GLD‐1 and GLD‐2 mRNA regulatory pathways, which promote meiotic development. As germ cells move proximally, GLP‐1 Notch signaling is reduced, allowing the GLD‐1 and GLD‐2 pathways to promote meiotic development. This core proliferation–meiotic entry network, consisting of canonical GLP‐1 Notch signaling inhibiting the GLD‐1 and GLD‐2 pathways, appears to function under all tested conditions (postembryonic developmental stage, sex, and standard growth temperatures). Additional factors are also involved in


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regulating this decision; however, their regulatory relationships relative to the core network and each other are much more complex because: (1) some appear to be active only under certain conditions; (2) some act in more than one place in the network; (3) some appear to both promote proliferation and promote meiotic development; and (4) some are involved in cross or feedback regulation. Other factors regulating this decision are inferred to exist but remain to be identified. Perhaps the complexity of the noncore genes is an indication that they may function in modulation of the proliferation–meiotic entry decision depending on conditions, such as environment. While further genetic screens will likely uncover new players, additional experimental approaches will be necessary. Many of the gene products acting in the network are mRNA binding proteins and/or function in mRNA metabolism. Therefore, it will be important to identify the mRNA targets of these regulators to further uncover and connect the processes controlled by the proliferation–meiotic entry network. With all of the insights we have gained regarding the proliferation–meiotic entry decision, it is still unclear whether the proliferative zone is composed of a single stem cell population or a small set of stem cells that progress to a larger group of transit amplifying cells and how the spatial control of GLP‐1 Notch signaling might dictate one or the other pattern.

Acknowledgments Research in the Hansen Laboratory is funded by grants from the Natural Sciences and Engineering Research Council of Canada and the Canadian Institutes of Health Research. D. H. is a Scholar of the Alberta Heritage Foundation for Medical Research. Research in the Schedl laboratory is supported by NIH R01 GM63310. We thank our colleagues in the Schedl and Hansen laboratories for stimulating discussions. Note Added in Proof: For additional experimental analysis of the proliferative zone, please consult Crittenden, S., Leonhard, K., Byrd, D., and Kimble, J. (2006). Cellular analysis of the mitotic region in the Caenorhabditis elegans adult germ line. Mol. Biol. Cell 17, 3051–3061.

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Regulation of Angiogenesis by Hypoxia and Hypoxia‐Inducible Factors Michele M. Hickey and M. Celeste Simon Department of Cell and Molecular Biology Abramson Family Cancer Research Institute, and Howard Hughes Medical Institute, University of Pennsylvania Philadelphia, Pennsylvania 19104

I. Introduction II. Mechanisms of Angiogenesis III. The Hypoxia‐Inducible Factor Family A. Structure of HIF Proteins B. Regulation of HIF Activity C. HIF Target Genes IV. V. VI. VII.

The Role of HIF in Developmental Angiogenesis The Role of HIF in Adult Tissues HIF and Ischemic Injury HIF and Cancer A. Tumor Angiogenesis and Hypoxia B. HIF Expression in Tumors C. von Hippel‐Lindau Disease D. The Role of HIF1 and HIF2 in Tumor Angiogenesis E. Therapeutic Implications

VIII. Conclusions Acknowledgments References

Maintenance of oxygen homeostasis is critical for the survival of multicellular organs. As a result, both invertebrates and vertebrates have developed highly specialized mechanisms to sense changes in oxygen levels and to mount adequate cellular and systemic responses to these changes. Hypoxia, or low oxygen tension, occurs in physiological situations such as during embryonic development, as well as in pathological conditions such as ischemia, wound healing, and cancer. A primary eVector of the adaptive response to hypoxia in mammals is the hypoxia‐inducible factor (HIF) family of transcription regulators. These proteins activate the expression of a broad range of genes that mediate many of the responses to decreased oxygen concentration, including enhanced glucose uptake, increased red blood cell production, and the formation of new blood vessels via angiogenesis. This latter process is dynamic and results in the establishment of a mature Current Topics in Developmental Biology, Vol. 76 Copyright 2006, Elsevier Inc. All rights reserved.

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0070-2153/06 $35.00 DOI: 10.1016/S0070-2153(06)76007-0

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vascular system that is indispensable for proper delivery of oxygen and nutrients to all cells in both normal tissue and hypoxic regions. Angiogenesis is essential for normal development and neoplastic disease as tumors must develop mechanisms to stimulate vascularization to meet increasing metabolic demands. The link between hypoxia and the regulation of angiogenesis is an area of intense research and the molecular details of this connection are still being elaborated. This chapter will provide an overview of current knowledge and highlight new insights into the importance of HIF and hypoxia in angiogenesis in both physiological and pathophysiological conditions. ß 2006, Elsevier Inc.

I. Introduction In the absence of a functional blood supply, the growth of both multicellular organisms and tumors is limited by the ability of oxygen to diVuse to cells from blood vessels. Decreased oxygen levels, or hypoxia, can develop within rapidly proliferating tissues or as the result of occlusion of blood vessels. Hypoxia leads to insuYcient cellular energy production as oxygen is essential for oxidative phosphorylation. At the same time, however, excessively high levels of oxygen are detrimental and can result in the production of reactive oxygen species (ROS) that damage cellular organelles and DNA. Therefore, it is imperative that oxygen concentrations be tightly regulated. At the systemic level, oxygen tension is detected by highly sensitive tissues, such as the carotid body, to eVect a rapid physiological response to acute hypoxia that includes increased ventilation and cardiac output. More prolonged hypoxia is also sensed at the cellular level, leading to the activation of molecular pathways to cope with this stress. The key mediators of this response are members of the hypoxia‐inducible factor (HIF) family of proteins. The hypoxic response and the HIF pathway are conserved from Caenorhabitis elegans and Drosophila melanogaster to mice and humans, emphasizing its importance in the maintenance of oxygen homeostasis. These proteins function as transcriptional regulators that stimulate the expression of a multitude of genes important for adaptation to hypoxia, including those encoding glucose transporter‐1 (Glut‐1), which increases cellular glucose uptake, and glycolytic enzymes, which mediate enhanced glycolysis to maintain ATP production in the face of lower oxygen levels. In addition, HIF‐stimulated erythropoietin (Epo) expression improves the oxygen carrying capacity of the blood by enhancing the production of erythrocytes. Another mechanism by which cells can alleviate the increasing metabolic demands presented by hypoxia is via new blood vessel formation through vasculogenesis and angiogenesis. These processes are complex and occur in a stepwise fashion. Angiogenesis is regulated by a balance of positive‐ and

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negative‐acting growth factors and by physiological stresses such as alterations in oxygen levels. Hypoxia stimulates the expansion and remodeling of the existing vasculature to enhance blood flow to oxygen‐deprived tissues. This is accomplished primarily through the activation of HIF target genes involved in various steps of angiogenesis such as vascular endothelial growth factor (VEGF). This regulation of angiogenesis by hypoxia and HIFs is essential for proper embryonic development and recovery after ischemic injury. In addition, numerous lines of evidence suggest that tumors develop regions of hypoxia and that the HIF pathway is an important component of tumor growth and angiogenesis.

II. Mechanisms of Angiogenesis The vascular network mediates the delivery of oxygen and nutrients to all cells of an organism and is essential for normal development and survival. Figure 1 outlines the mechanisms by which the vasculature is formed and highlights several of the key molecules involved in regulating these processes. Mesodermal progenitor cells known as hemangioblasts represent bipotential precursor cells that give rise to both endothelial and hematopoietic cells (Sabin, 1920). A putative hemangioblast, the blast colony‐forming cell (BL‐CFC), has been identified in vitro in embryoid bodies derived from embryonic stem (ES) cells and has been shown to express the VEGF‐receptor‐2 (VEGF‐R2/Flk1) (Choi et al., 1998; Kennedy et al., 1997; Yamaguchi et al., 1993). Hemangioblasts

Figure 1 Overview of vascular development. During embryonic development, mesoderm‐ derived precursor cells known as hemangioblasts are stimulated by VEGF and FGF signaling to give rise to both hematopoietic and endothelial cells (ECs). A primary capillary network is formed by the process of vasculogenesis in which endothelial progenitors, or angioblasts, diVerentiate to form primitive blood vessels in the yolk sac and embryo. VEGF signaling through VEGF‐R2/Flk‐1 is essential for this process. Subsequently, new capillaries are formed by sprouting or splitting of existing vessels through angiogenesis. VEGF is also critical for this process and acts together with angiopoietins to stimulate the proliferation and migration of ECs. These vessels are stabilized through the recruitment of supporting cells to form the mature vasculature.

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have also been detected in vivo in the mouse embryo during gastrulation as mesodermal cells that express brachyury and VEGF‐R2 (Huber et al., 2004). Signaling through VEGF and members of the fibroblast growth factor (FGF) family is required for the diVerentiation of mesoderm into these precursor cells (Faloon et al., 2000; Kennedy et al., 1997). In addition, hypoxia has been shown to enhance the production of BL‐CFCs in a HIF‐dependent manner (Ramirez‐Bergeron et al., 2004). Endothelial cell (EC) precursors, or angioblasts, aggregate within the yolk sac and embryo and subsequently diVerentiate to give rise to a primary capillary plexus in the process of vasculogenesis (Conway et al., 2001). Additional capillaries can be generated by sprouting from and splitting of existing vessels through angiogenesis. Degradation of the extracellular matrix (ECM) by matrix metalloproteinases (MMPs) allows ECs to migrate in response to chemotactic growth factors. VEGF and the angiopoietins, together with FGF proteins and platelet‐derived growth factor (PDGF), mediate the migration and proliferation of ECs to newly formed vessels (Conway et al., 2001; Jain, 2003; Risau, 1997). Primitive blood vessels synthesize ECM proteins and recruit supporting cells, such as pericytes and smooth muscle cells, as they mature. PDGF, angiopoietin‐1 (Ang‐1), and transforming growth factor (TGF)‐ are important in the regulation of these processes (Jain, 2003). Angiogenesis is controlled by a balance between stimulatory and inhibitory factors and the so‐called ‘‘angiogenic switch’’ occurs when this balance shifts in favor of positive stimuli (Carmeliet, 2005; Hanahan and Folkman, 1996). Among these factors, VEGF is essential for most steps in vasculogenesis and angiogenesis, acting in a paracrine manner to stimulate diVerentiation of VEGF‐R2þ angioblasts, proliferation and survival of ECs, and sprouting of new blood vessels (Carmeliet, 2000; Ferrara and Gerber, 2001; Veikkola et al., 2000). The importance of VEGF signaling is supported by the finding that embryos lacking VEGF‐R2 do not form hematopoietic cells or ECs (Shalaby et al., 1995). In contrast, loss of VEGF‐R1 expression results in defective formation of mature blood vessels, demonstrating the role of this receptor later in angiogenesis and vessel maturation (Fong et al., 1995). Regulation of VEGF expression itself is also essential in order to sustain proper angiogenesis during development and to control angiogenesis in pathogenic situations. Deficiency in Vegf expression is early embryonic lethal due to severe defects in vascularization; importantly loss of a single Vegf allele is suYcient to disrupt angiogenesis and cause lethality (Carmeliet et al., 1996; Damert et al., 2002; Duan et al., 2003; Ferrara et al., 1996). Therefore, the dosage of VEGF expression in the embryo is absolutely critical, as overexpression of Vegf leads to cardiac defects and developmental lethality (Miquerol et al., 2000). As a result, there are multiple mechanisms by which VEGF is controlled by hypoxia, including transcriptional upregulation,


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mRNA stabilization, and the presence of an internal ribosomal entry site to allow for maintenance of translation under hypoxic conditions (Forsythe et al., 1996; Levy et al., 1998; Stein et al., 1998). Although mature vessels are mostly quiescent, angiogenesis also occurs in the adult, for example, during wound healing and in response to hypoxic stress such as myocardial ischemia, and VEGF signaling is critical in triggering this process. In addition, abnormal angiogenesis is a feature of a number of pathological conditions including diabetes, hypertension, and cancer (Carmeliet, 2003). Dysregulation of the signals governing angiogenesis and the maturation of vessels can result either in insuYcient vascularization or in excessive formation of vessels that are often abnormal, immature, or leaky (Carmeliet, 2003). It has been suggested that a population of bone marrow‐ derived endothelial progenitor cells (EPCs) exists in the adult and is recruited by VEGF for neovascularization (Asahara et al., 1997; Grunewald et al., 2006; Khakoo and Finkel, 2005; Lyden et al., 2001; Rafii and Lyden, 2003; Reyes et al., 2002; Takahashi et al., 1999; Urbich and Dimmeler, 2004). In addition, hematopoietic cells of the myeloid lineage that express the angiopoietin receptor Tie‐2 (Tie2‐expressing monocytes or TEMs) have been shown to originate from the bone marrow and circulate in the peripheral blood (De Palma et al., 2003, 2005; Grunewald et al., 2006). Elimination of these TEM cells resulted in a lack of angiogenesis and tumor growth, suggesting that these cells may play an important role in tumor vascularization (De Palma et al., 2003, 2005; Lyden et al., 2001). The potential importance of bone marrow‐derived myeloid cells in tumor angiogenesis will be further discussed in Section VII.

III. The Hypoxia‐Inducible Factor Family A. Structure of HIF Proteins HIF proteins are members of a larger, evolutionarily conserved group of proteins known as bHLH‐PAS (basic helix loop helix‐Per ARNT Sim) proteins (Crews, 1998). These proteins function as sensors of environmental stimuli and activate the expression of genes important for angiogenesis, as well as circadian rhythms, xenobiotic detoxification, and adaptation to hypoxia (Crews and Fan, 1999; Gu et al., 2000; Kewley et al., 2004). Each member of this family contains an N‐terminal bHLH domain that mediates binding to consensus DNA sequences in the promoters of target genes (Murre et al., 1989). HIF proteins heterodimerize via their HLH and PAS domains in the center of each protein to form functionally active transcription factors (Wang and Semenza, 1995; Wang et al., 1995). The C‐terminus consists of one or two transactivation domains (TADs) that bind transcription

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Figure 2 Generalized structure of human HIF proteins. HIF proteins bind DNA at target gene promoters via the basic helix‐loop‐helix (bHLH) domain at the N‐terminus. The HLH and PAS (Per‐ARNT‐Sim) domains mediate heterodimerization of and subunits to form an active transcription complex. HIF1 and HIF2 each contain two transactivation domains (TADs); the C‐TAD binds to the coactivator proteins p300 and CBP. Stability of the HIF proteins is regulated by hydroxylation of two proline residues located within the oxygen‐ dependent degradation domain (ODD) via the prolyl hydroxylase (PHD) enzymes. In addition, hydroxylation of an asparagine residue located within the C‐terminus by the factor inhibiting HIF‐1 (FIH‐1) enzyme prevents association of HIF with p300/CBP.

cofactors such as p300 and CREB binding protein (CBP) (Fig. 2) (Arany et al., 1996; Carrero et al., 2000; Ebert and Bunn, 1998; Ema et al., 1999; Jiang et al., 1997b; O’Rourke et al., 1999; Pugh et al., 1997). The HIF family is comprised of three subunits: HIF1 [also known as MOP1 (member of PAS1)], HIF2 [also known as EPAS1 (endothelial PAS domain protein 1), MOP2, HLF (HIF1‐like factor), and HRF (HIF‐related factor)], and HIF3 (also known as MOP3); and three subunits: HIF / ARNT (aryl hydrocarbon nuclear translocator), ARNT2, and ARNT3 (Ema et al., 1997; Flamme et al., 1997; Gu et al., 1998; Tian et al., 1997; Wang and Semenza, 1995; Wang et al., 1995). HIF proteins form heterodimers of and subunits; HIF1/ARNT and HIF2/ARNT complexes have been shown to be primarily responsible for the hypoxic induction of angiogenesis (Jiang et al., 1996; Wenger and Gassmann, 1997). These heterodimeric protein complexes activate transcription by binding to hypoxia response elements (HREs), the first of which was identified in the Epo gene and contains a core sequence of CTACGTGCT (Semenza and Wang, 1992; Semenza et al., 1991; Wang et al., 1995). HIF1 and HIF2 are highly homologous proteins that share approximately 48% overall identity with the highest degree of similarity in the amino‐terminus (Ema et al., 1997; Tian et al., 1997). In contrast, there is some divergence in the C‐terminal sequences, which may help to explain diVerences in the genes activated by each subunit (see later). Expression of the principal subunit, ARNT, is constitutive, whereas subunit levels are tightly regulated and tissue‐restricted, allowing for spatial and temporal control of HIF‐dependent gene activation. HIF1 has been shown to be expressed nearly ubiquitously in humans and mice, whereas HIF2 expression is more spatially restricted. For example, HIF2 is highly


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expressed in tissues such as ECs, from which it was first identified (Ema et al., 1997; Jain et al., 1998; Tian et al., 1997). HIF2 mRNA is also detected in highly vascularized organs such as the heart, placenta, and lung (Ema et al., 1997; Tian et al., 1997). HIF2 protein is expressed during development in the lung and neural crest derivatives as well as in hypoxic tissues in the adult including bone marrow macrophages, kidney epithelial cells, liver parenchyma, cardiac myocytes, and pancreatic parenchymal cells (Ema et al., 1997; Talks et al., 2000; Wiesener et al., 2003).

B. Regulation of HIF Activity Activation of the HIF pathway is regulated at many levels, including mRNA expression, protein stability, subcellular localization, and activity, to ensure timely and eYcient induction of the hypoxic response. Changes in cellular oxygen concentrations are the major stimuli governing the regulation of HIF activity, but several nonhypoxic factors can also stimulate HIF. For example, a number of growth factors, such as insulin, epidermal growth factor, and PDGF, as well as cytokines, like tumor necrosis factor‐ and interleukin‐ 1 , can increase expression of the HIF1 protein (Haddad and Land, 2001; Stiehl et al., 2002; Treins et al., 2002; Zelzer et al., 1998; Zhou et al., 2003). Importantly, however, the amplitude of HIF induction by such stimuli is much lower than hypoxic activation of HIF (Arsham et al., 2002). In addition, oncogenes such as HER2neu, H‐ras, and v‐Src enhance HIF1 activity (Chen et al., 2001; Jiang et al., 1997a; Laughner et al., 2001). These results emphasize the importance of HIF activation during normal growth and development, as well as during the pathological processes of inflammation and tumorigenesis. The principal mechanism of HIF regulation is its posttranslational modification and subsequent degradation under normoxia. Both HIF and HIF subunits are constitutively transcribed and translated; however, regulation of the stability of the HIF subunits controls HIF‐dependent gene expression. Under normoxic conditions the subunits have a very short half‐life and are undetectable, whereas these proteins are rapidly stabilized in response to hypoxic stimulation (Salceda and Caro, 1997; Wang et al., 1995). The subunits contain an oxygen‐dependent degradation domain (ODD) (Fig. 2) that contains two conserved proline residues (Pro402 and Pro564 in humans) as part of an LXXLAP consensus motif. The ODD is suYcient for destabilization and degradation of HIF under normoxia (Huang et al., 1998; Kaelin, 2002). These proline residues are hydroxylated by a family of iron‐dependent dioxygenases in oxygenated cells known as the prolyl hydroxylase domain proteins (PHD1, ‐2, and ‐3 in humans) (Bruick and McKnight, 2001; Epstein

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et al., 2001; Ivan et al., 2001; Jaakkola et al., 2001; Masson et al., 2001; Yu et al., 2001). The C. elegans homolog of the PHD enzymes was first identified as a gene (egl‐9) associated with an abnormal egg‐laying phenotype, and orthologs have subsequently been described in D. melanogaster, mice, rats, and humans (Bruick and McKnight, 2001; Epstein et al., 2001). The hydroxylation reaction adds one atom of oxygen to form hydroxyproline with the production of succinate and carbon dioxide as by‐products. PHD enzymes function similarly to collagen prolyl hydroxylases and require molecular oxygen as a substrate in addition to iron, ascorbate, and 2‐oxoglutarate as cofactors. Therefore, these enzymes are catalytically inactive in the absence of oxygen or in the presence of iron chelators and cobalt ions that abolish iron binding (Masson and RatcliVe, 2003). In addition to the PHD enzymes, other studies have demonstrated that mitochondria act as cellular oxygen sensors that activate HIF through the generation of ROS (Brunelle et al., 2005; Chandel et al., 1998, 2000; Guzy et al., 2005; Mansfield et al., 2005). Hydroxylation of HIF1 and HIF2 subunits at these key proline residues under normal oxygen tensions provides a recognition site for binding of the von Hippel‐Lindau protein (pVHL), which is the substrate‐recognition component of a complex consisting of elongins B and C, Cul2, and Rbx‐1 that mediates the polyubiquitination of HIF proteins (Fig. 3) (Cockman et al., 2000; Ivan et al., 2001; Iwai et al., 1999; Jaakkola et al., 2001; Kamura et al., 1999; Kibel et al., 1995; Lisztwan et al., 1999; Lonergan et al., 1998; Maxwell et al., 1999; Pause et al., 1997; Yu et al., 2001). Following binding of pVHL, HIF subunits are rapidly degraded via the 26S proteasome, thereby preventing transactivation of HIF target genes under normoxia (Kim and Kaelin, 2003). PHD enzymatic activity is inhibited in hypoxic cells and unhydroxylated HIF proteins escape recognition by pVHL. Stabilized HIF then translocates to the nucleus, where it binds to its dimerization partner ARNT to form an active transcription complex (Fig. 3) (Kallio et al., 1999; Tanimoto et al., 2000). pVHL is encoded by the VHL tumor suppressor gene, which is mutated in a hereditary cancer syndrome known as VHL disease (see Section VII for more detail) (Gnarra et al., 1997; Iliopoulos et al., 1995; Latif et al., 1993). Briefly, germline mutation of VHL results in predisposition to the development of a spectrum of tumors including clear cell renal cell carcinoma (RCC), cerebellar and retinal hemangiomas, and pheochromocytomas (Ivan and Kaelin, 2001). These tumors arise on loss or inactivation of the second allele of VHL in somatic tissues and are highly vascularized with elevated levels of VEGF and Epo, two important HIF target genes. Loss of pVHL activity mimics a hypoxic state and allows for constitutive HIF activity under normoxic conditions (Iliopoulos et al., 1996; Maxwell et al., 1999). A second level of HIF regulation occurs through the hydroxylation of asparagine 803 in the C‐terminal activation domain in the presence of


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Figure 3 Regulation of HIF activity. Under normoxic conditions, HIF subunits are polyubiquitinated at two proline residues within the oxygen‐dependent degradation domain (ODD) by a family of enzymes known as prolyl hydroxylases (PHDs). This promotes recognition by the pVHL E3 ubiquitin ligase complex and subsequent degradation of HIF via the proteasome. In addition, hydroxylation of a C‐terminal asparagine residue of HIF by factor inhibiting HIF‐1 (FIH‐1) prevents binding of cofactors required for HIF activity. However, hypoxia inhibits the activity of the PHD and FIH‐1 enzymes, allowing HIF proteins to escape recognition by pVHL, be stabilized, and translocate to the nucleus. There they dimerize with HIF /ARNT and bind hypoxia response elements (HREs) within the promoters of target genes. Together with the coactivator proteins p300 and CBP, the HIF complex activates the transcription of a panel of genes required for the response to hypoxia.

oxygen. This reaction is accomplished by a dioxygenase known as factor inhibiting HIF‐1 (FIH‐1) (Mahon et al., 2001). Hydroxylation of this asparagine residue inhibits HIF activity by preventing the recruitment of the coactivators p300 and CBP (Hewitson et al., 2002; Lando et al., 2002; McNeill et al., 2002). Since FIH‐1 requires oxygen for catalytic activity, the C‐terminus of HIF remains unmodified under hypoxia and can interact with its cofactors to productively activate transcription of its target genes.

C. HIF Target Genes The first HIF target gene identified was Epo, which stimulates red blood cell production and is markedly upregulated on hypoxic exposure (Semenza and Wang, 1992; Wang et al., 1995). It was soon realized, however, that HIF functions as a master regulator of the long‐term response and adaptation to

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hypoxia by activating the transcription of close to 200 genes (Semenza, 2003). These genes are involved in a broad range of cellular processes as shown in Table I. For example, HIF induces the expression of a number of metabolic genes that maintain glucose uptake and ATP production through glycolysis in the face of oxygen deprivation. These genes include, but are not limited to, Glut‐1, lactate dehydrogenase A (LDHA), phosphoglycerate kinase (PGK), and aldolase A (Hu et al., 2003, 2006; Semenza, 2003). HIF can also enhance the expression of genes that modulate cell proliferation, viability, and apoptosis, such as insulin‐like growth factor‐2 (IGF‐2), the cyclin dependent kinase inhibitor p21, and the proapoptotic gene Bnip3 (Bruick, 2000; Feldser et al., 1999; Goda et al., 2003; Hu et al., 2003, 2006; Sowter et al., 2001). Importantly, a large percentage of genes induced in the hypoxic response is involved in various steps of angiogenesis. One of the most important of such genes is VEGF, which is a primary regulator of the formation of new blood vessels (Forsythe et al., 1996). HIF also induces the expression of VEGF‐R1 (Flt‐1), VEGF‐R2 (Flk‐1), plasminogen activator inhibitor‐1 (PAI‐1), Ang‐1 and ‐2, the Tie‐2 receptor, and MMP‐2 and ‐9 (Ben‐Yosef et al., 2002; Currie et al., 2002; Elvert et al., 2003; Gerber et al., 1997; Kietzmann et al., 1999, 2003; Melillo et al., 1997; Tian et al., 1997). Furthermore, hypoxic stabilization of HIF results in the induction of genes that control vascular tone and blood flow, such as nitric oxide synthases (iNOS) and adrenomedullin (ADM), as well as inflammatory cytokines such as interleukin‐8 (IL‐8) (Cejudo‐Martin et al., 2002; Desbaillets et al., 1999; Garayoa et al., 2000; Melillo et al., 1995, 1997). The net result of the activation of these genes is the stimulation of increased vessel formation and remodeling to provide adequate oxygen delivery to hypoxic tissues. HIF‐ dependent expression of these genes is essential for developmental angiogenesis and the response to ischemic insults, and upregulation of proangiogenic HIF target genes has been demonstrated to occur in tumorigenesis. As stated earlier, HIF1 and HIF2 are the primary regulators of genes essential for the hypoxic response. In addition to the tissue‐specific expression patterns of HIF1 and HIF2, there are diVerences in the target genes activated by each subunit. These two proteins share a number of targets, suggesting that there may be redundancy in the function of HIF subunits (Hu et al., 2003). However, there are likely diVerences in the genes activated by each protein depending on the cell type, even when both subunits are present. HIF1 has been shown to uniquely activate the glycolytic enzymes, including PGK and aldolase A, while Oct‐4, a transcription factor involved in maintaining pluripotentiality of ES cells, appears to be regulated by HIF2 but not HIF1 (Covello et al., 2005, 2006; Hu et al., 2003). The mechanisms governing target gene specificity are the subject of investigation.


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Table I Representative Examples of Genes that Are Regulated by HIF Signaling Under Hypoxia Partial List of Genes Transcriptionally Activated by HIF Cell proliferation and apoptosis Insulin‐like growth factor‐2 (IGF‐2) Transforming growth factor‐ (TGF‐) Transforming growth factor‐ (TGF‐ ) p21 Bnip3 Glucose metabolism Phosphoglycerate kinase 1 (PGK) Glucose transporter‐1 (Glut‐1) Glucose transporter‐3 (Glut‐3) Lactate dehydrogenase A (LDHA) Aldolase A Phosphofructokinase L (PFKL) pH regulation Carbonic anhydrase IX (CAIX) Erythropoiesis Erythropoietin (Epo) Iron metabolism Transferrin Transferrin receptor Extracellular matrix metabolism Matrix metalloproteinase‐2 (MMP‐2) Fibronectin Inflammation Interleukin‐8 (IL‐8) Chemokine receptor CXCR4 Stromal cell‐derived factor‐1 (SDF‐1) Transcription factors Oct‐4/Pou5F1 Angiogenesis and control of vascular tone Vascular endothelial growth factor (VEGF) VEGF‐receptor 1 (VEGF‐R1/Flt‐1) VEGF‐receptor 2 (VEGF‐R2/Flk‐1) Platelet‐derived growth factor‐ (PDGF‐ ) Angiopoietin‐2 (Ang‐2) Tie‐2 Endothelin‐1 (ET‐1) Plasminogen activator inhibitor‐1 (PAI‐1) Inducible nitric oxide synthase 2 (iNOS2) Adrenomedullin (ADM)

Although the primary role of HIF is the direct activation of target genes, it has been appreciated that there is substantial cross‐talk between the HIF pathway and other signaling pathways. This interaction between HIF and

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other cellular signal transduction molecules may serve to amplify and broaden the hypoxic response. For example, hypoxia has been shown to inhibit the diVerentiation of cell types including myogenic cells and neural stem cells through enhanced Notch signaling in a manner that is dependent on HIF1 function (Gustafsson et al., 2005). HIF also has an eVect on genes regulated by c‐Myc; HIF1 inhibits the ability of c‐Myc to repress p21, resulting in hypoxia‐induced cell cycle arrest (Goda et al., 2003; Koshiji et al., 2004; Mack et al., 2005). HIF1 interacts directly with and stabilizes the tumor suppressor p53, whereas p53 targets HIF1 for ubiquitination (An et al., 1998; Ravi et al., 2000). Furthermore, links between HIF and other transcription factors, including c‐Jun and NF‐B, have been suggested (Alfranca et al., 2002; Bracken et al., 2005; Gerald et al., 2004; Jung et al., 2003a,b; Laderoute et al., 2002; Walmsley et al., 2005; Zhou et al., 2003). It is likely that the eVect of hypoxia on gene expression and the role of HIF in regulating transcription will become increasingly complex as additional mechanisms of cross‐talk are elucidated.

IV. The Role of HIF in Developmental Angiogenesis Hypoxia is an important feature of embryonic development: as the embryo grows and develops, it quickly outstrips the oxygen and nutrients provided by diVusion alone (Maltepe and Simon, 1998). These naturally occurring hypoxic gradients trigger the expression of genes critical for the formation of a complex network of blood vessels to provide adequate oxygenation of the dividing tissues. A number of HIF target genes has been shown to be essential for proper vascular development, including Vegf and Epo (Carmeliet et al., 1996; Ferrara et al., 1996; Kieran et al., 1996; Lin et al., 1996; Wu et al., 1995, 1999). Mice lacking these genes die during embryonic development with severe vascular and hematopoietic defects. Furthermore, gene dosage of Vegf in the yolk sac and embryo is critical for proper angiogenesis and development (Damert et al., 2002; Duan et al., 2003; Miquerol et al., 2000). These results demonstrate the importance of the expression of HIF targets during development. In agreement with these findings and with the role of HIF as the central mediator of the hypoxic response, inactivation of HIF subunits in mouse models also results in embryonic lethality. Targeted deletion of the Arnt gene encoding HIF1 /ARNT leads to lethality between days 9.5 and 10.5 of embryonic development (E9.5–10.5). The absence of Arnt expression resulted in defective blood vessel formation in the yolk sac and ineYcient vessel remodeling and maturation in the embryo (Kozak et al., 1997; Maltepe et al., 1997). Loss of Arnt resulted in significantly reduced hematopoietic progenitor development in the yolk sac; this defect was cell extrinsic


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and could be rescued with exogenous VEGF (Adelman et al., 1999). Explants from Arnt/ embryos also displayed abnormal hematopoiesis as well as a failure to undergo vasculogenesis or angiogenesis (Ramirez‐Bergeron et al., 2006). Again, administration of recombinant VEGF rescued these vascular defects, confirming that VEGF secreted by hematopoietic cells is necessary for vessel development (Ramirez‐Bergeron et al., 2006). Furthermore, Arnt deficiency resulted in aberrant architecture of the placenta, the vascular tissue that provides nutrients to the growing embryo (Adelman et al., 2000). Aggregation of Arnt/ ES cells with tetraploid wild‐type embryos rescued the placental defect; however, these embryos still exhibited defects in yolk sac vascularization and malformations in the endocardial cushion, leading to impaired cardiac function (Adelman et al., 2000). These phenotypes strongly support the essential role of HIF signaling, through the activation of targets such as VEGF and Epo, in both extraembryonic and embryonic tissues for the establishment of the vasculature and the hematopoietic system. Another important molecule in the oxygen‐sensing pathway, VHL, also plays an important role in angiogenesis. Vhl / mice die between E10.5 and E12.5 due to defects in placental vascularization with an absence of embryonic blood vessels in the placental labyrinth, hemorrhaging, and necrosis (Gnarra et al., 1997). EC‐specific deletion of the Vhl gene is embryonic lethal at E12.5 with significant hemorrhaging in the head and heart regions, a disorganized and dilated yolk sac vasculature, and defective embryonic vasculogenesis in the placenta (Tang et al., 2006). Deletion of Hif1a expression in these mice did not rescue this phenotype, suggesting that the role of Vhl in embryonic angiogenesis is independent of HIF1. pVHL has been shown to bind to fibronectin, an ECM protein that is necessary for proper vessel formation (Ohh et al., 1998; Stickle et al., 2004). Tang et al. (2006) showed that loss of Vhl expression in ECs resulted in reduced fibronectin deposition around vessels both in the yolk sac and embryo, leading to increased EC permeability and impaired EC migration. Therefore, pVHL regulation of fibronectin assembly, in addition to its role in negatively regulating the HIF pathway, is required for embryonic angiogenesis. Deficiency in Hif1a results in embryonic lethality during midgestation at around E10.5 similar to loss of Arnt (Iyer et al., 1998; Ryan et al., 1998). Hif1a/ ES cells expressed decreased levels of glycolytic enzymes and VEGF mRNA in response to hypoxia, confirming the role of HIF1 in regulating hypoxic gene expression. In addition, loss of Hif1a resulted in decreased ES cell proliferation. Hif1a/ embryos were developmentally delayed with fewer somites than wild‐type embryos, and displayed a disorganized yolk sac vasculature lacking branching. Mutant embryos developed severe defects in neuronal development, such as failure of the neural tube to close, and significant cardiovascular abnormalities, including defective

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ventricle formation (Compernolle et al., 2003; Iyer et al., 1998; Ryan et al., 1998). These phenotypes correlated with increased hypoxia in the neural ectoderm and enhanced apoptosis of cephalic mesenchymal cells compared to wild‐type embryos (Iyer et al., 1998; Ryan et al., 1998). Moreover, although vascularization of the embryo initiated properly in Hif1a‐deficient embryos, these vessels, especially those in the cephalic region, regressed by E9.5 and did not form a complete capillary network (Iyer et al., 1998; Ryan et al., 1998). In agreement with these results, neural cell‐specific deletion of Hif1a led to reduced expression of HIF targets, including VEGF, and decreased vessel density in the brain at E19, which was accompanied by enhanced apoptosis (Tomita et al., 2003). These findings demonstrate that HIF1 is necessary for cardiovascular development, neural crest cell migration, and embryonic angiogenesis. Four independent mouse models for Hif2a deficiency have been generated, each of which is embryonic lethal but with dramatically diVerent phenotypes (Compernolle et al., 2002; Peng et al., 2000; Scortegagna et al., 2003a; Tian et al., 1998). In the first of these models, Hif2a‐deficient embryos died by E16.5 due to bradycardia and cardiac failure. This was attributed to decreased production of catecholamines from the Hif2a‐expressing organ of Zuckerkandl, possibly due to reduced tyrosine hydroxylase expression (Tian et al., 1998). Despite the initial identification of the Hif2a gene in ECs, no vascular defects were observed in these embryos. Mutant embryos in the second model died between E9.5 and E13 with abnormal vessel remodeling postvasculogenesis and hemorrhaging in the yolk sac and embryo (Peng et al., 2000). The third model demonstrated 50% lethality by E13.5 as a result of cardiac failure, while the Hif2a/ mice that survived died later as neonates of respiratory distress syndrome (Compernolle et al., 2002). This phenotype was found to be the result of reduced VEGF levels and consequently decreased surfactant production by alveolar type 2 cells within the lung. This result suggests a role for HIF2 in the regulation of VEGF expression in and development of the embryonic lung, which expresses high levels of HIF2 (Ema et al., 1997). In contrast to these previous models, Scortegagna et al. (2003a) were able to obtain viable Hif2a‐deficient adult mice, although at a significantly reduced frequency, by intercrossing mice from two genetic backgrounds. These mice presented with multiple organ pathology, the phenotype of which is discussed in greater detail in Section V. The diVerences in these phenotypes are likely attributable to modifying genes in the various genetic backgrounds used in each model. Nevertheless, the impaired embryonic survival observed in the absence of HIF2 expression emphasizes that HIF1 and HIF2 have distinct and nonoverlapping, essential functions in developmental angiogenesis. Further evidence of the importance of HIF2 in development comes from recent work that identified Oct‐4/Pou5F1 as a HIF2‐unique target gene


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(Covello et al., 2006). Using a knock‐in approach, the Hif1a locus was replaced with Hif2a, resulting in expanded HIF2 expression and a corresponding increase in Oct‐4 levels. Enhanced HIF2 expression is embryonic lethal, suggesting that there is a dose‐dependent requirement for HIF2‐controlled Oct‐4 expression during hematopoietic stem cell diVerentiation and for the survival of primordial germ cells (Covello et al., 2006).

V. The Role of HIF in Adult Tissues In addition to its role in development, HIF is important for the physiological hypoxic response in the adult. Mice heterozygous for Hif1a have an impaired response to chronic hypoxic exposure, including delayed erythropoiesis and decreased pulmonary vascular remodeling (Yu et al., 1999). HIF1 is induced in keratinocytes during wound healing of the skin, overlapping to some extent with VEGF expression (Elson et al., 2000). Several groups have used conditional gene targeting of a floxed Hif1a allele to study the function of HIF1 in specific adult tissues. For example, deletion of Hif1a in the brain resulted in decreased numbers of cells in the cortex and defective spatial memory (Tomita et al., 2003). Hif1a has also been shown to be necessary for chondrocyte survival and for mammary gland diVerentiation and lactation, independent of its role in inducing VEGF expression and vessel formation (Schipani et al., 2001; Seagroves et al., 2003). Deletion of Hif1a in cardiac myocytes impaired contraction of cardiac muscle and decreased ATP levels in the heart. Hif1a deficiency in the heart also resulted in a reduction in VEGF expression and in the number of vessels in the left ventricle (Huang et al., 2004). Loss of Hif1a is also detrimental for the function of macrophages, as HIF1 is required for glycolysis and ATP production in these cells (Cramer et al., 2003). In the absence of Hif1a, the expression of genes, including PGK and VEGF, is significantly decreased and the migration and invasion of macrophages is impaired both in vitro and in physiologic assays. A number of the molecules released by macrophages are proangiogenic, such as VEGF, angiopoietins, IL‐8, MMP‐2, and MMP‐9, emphasizing the importance of these cells in stimulating the formation of new blood vessels in hypoxic tissues (Pollard, 2004). Hypoxia and HIF have been shown to regulate the expression of many of these factors, suggesting a role for the HIF pathway in macrophage‐induced angiogenesis (Semenza, 2003). The potential implications of HIF1 and HIF2 expression in macrophages in the context of tumor angiogenesis will be discussed more thoroughly in Section VII. Importantly, an EC‐specific deletion using Tie2‐Cre demonstrated a role for Hif1a in angiogenesis (Tang et al., 2004). Mice lacking Hif1a expression in ECs developed normally, suggesting that HIF2 may be important or

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may compensate for loss of HIF1 during formation of the embryonic vasculature. However, ECs lacking Hif1a displayed reduced proliferation and decreased tubular network formation under hypoxia, as well as an inability to eYciently form new capillaries in wound healing assays. Furthermore, mutant ECs exhibited impaired ECM penetration in matrigel and defective chemotactic migration, which was attributable to decreased VEGF expression (Tang et al., 2004). These results support the importance of HIF1 in the function of ECs in response to hypoxia and the potential role of HIF1 in neovascularization during wound healing and tumor formation (Elson et al., 2000). It remains to be determined whether these angiogenic phenotypes are secondary to the glycolytic defects that are observed in the absence of HIF1. As described earlier, Hif2a deficiency was found to be embryonic lethal in several independent studies. However, by mating mice of two diVerent backgrounds (129 and C57), Scortegagna et al. (2003a) generated viable homozygous Hif2a‐null mice, although at less than the expected frequency. Mice lacking Hif2a were smaller in size and had shorter lifespans due to pathological phenotypes in multiple organs. For example, mutant mice developed cardiac hypertrophy, steatosis of the liver, and retinopathy. In addition, mitochondrial dysfunction was observed with increased oxidative stress, as measured by elevated levels of ROS. This finding suggests that HIF2 may regulate the expression of antioxidant enzymes that eliminate ROS. These mutant mice also had a hypocellular bone marrow with significantly reduced hematocrit levels and red blood cell numbers, likely due to a defect in the bone marrow stroma (Scortegagna et al., 2003b). In addition, Epo production was decreased in Hif2a/ kidneys, demonstrating that HIF2 is essential for the regulation of renal Epo expression and thus for global hematopoiesis (Scortegagna et al., 2005). Future studies using conditional, tissue‐specific Hif2 knockout models will likely reveal additional functions for this protein in the adult.

VI. HIF and Ischemic Injury Levels of HIF1 mRNA and protein increase following ischemic insults in a number of tissues, including the retina, heart, brain, and lung, in both mice and humans (Bergeron et al., 1999; Lee et al., 2000; Ozaki et al., 1999; Yu et al., 1998). This finding supports a role for the HIF pathway in pathophysiological angiogenesis. For example, HIF activity has been associated with ischemia in the retina that is associated with diabetic retinopathy and retinopathy of prematurity (ROP), in which retinal vessels become ischemic due to occlusion, resulting in neovascularization. This increased angiogenesis can lead to blindness if uncontrolled, and is dependent on


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enhanced VEGF expression, possibly mediated by HIF1 (Ferrara, 1999; Ozaki et al., 1999). In order to study HIF2 function in the retina, Morita et al. (2003) generated a hypomorphic Hif2a allele via the insertion of a floxed neomycin gene into exon 1. Mice homozygous for the interrupted Hif2a allele were viable and fertile without developmental abnormalities. Using an animal model of ROP in which mice were exposed to hyperoxia followed by a return to room air, these authors demonstrated that mice with reduced Hif2a expression did not undergo retinal angiogenesis and displayed a lower induction of Epo compared to wild‐type mice (Morita et al., 2003). This finding suggests that HIF2 is an important mediator of the vascular phenotype associated with ROP and therefore may represent a useful therapeutic target. In the case of myocardial tissue, blockage of coronary arteries inhibits blood flow to the heart, resulting in ischemia and induction of HIF1 expression (Lee et al., 2000). HIF1 expression correlates with enhanced VEGF production and neovascularization in animal models and in humans (Lee et al., 2000; Martin et al., 1998). Similarly, increased HIF1 mRNA is associated with expression of HIF target genes, such as glycolytic enzymes and VEGF, in brain tissue surrounding the site of ischemia (Bergeron et al., 1999; Marti et al., 2000). In patients with chronic obstructive lung disease, hypoxia results in vascular remodeling that reduces blood flow and causes heart failure. HIF1 appears to play a role in this process, as mice heterozygous for this gene display reduced pulmonary remodeling (Yu et al., 1999). HIF1 signaling has also been implicated in the pathogenesis of preeclampsia, in which placental trophoblasts fail to invade the maternal decidua, leading to defective vascular remodeling of the uterine arteries. Hypoxia in early pregnancy stimulates proliferation of placental trophoblasts through HIF1‐dependent activation of TGF‐ 3 expression (Caniggia et al., 1999, 2000). Downregulation of HIF activity is thought to be necessary for trophoblasts to become invasive and to allow for perfusion of the placenta through maternal blood vessels. Administration of angiogenic factors, including VEGF and FGF, and transplantation of bone marrow cells have been tested as potential treatments to enhance vascularization in ischemic disease (Rafii and Lyden, 2003; Simons, 2005; Tateishi‐Yuyama et al., 2002). The above‐mentioned findings suggest that increasing HIF1 levels may also be of therapeutic value in treating ischemic disorders through the induction of neovascularization. Evidence supporting this hypothesis comes from the finding that transgenic mice overexpressing active HIF1 in the skin have a marked increase in VEGF expression and vascularization; importantly, these vessels were functionally sound and leakage‐resistant (Elson et al., 2001). Treatment with DNA encoding the N‐terminus of HIF1 fused to the VP16 transactivation domain stimulated angiogenesis in both a rabbit hindlimb ischemia model

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and a model of myocardial infarction in the rat (Shyu et al., 2002; Vincent et al., 2000). In an alternative approach, a macrophage‐derived peptide, PR39, that inhibits HIF degradation by interacting with the proteasome has been shown to increase angiogenesis in ischemic mouse cardiac tissue (Li et al., 2000). In addition, overexpression of peptides containing the HIF prolyl hydroxylation sites impeded degradation of endogenous HIF1, resulting in induction of HIF target genes and angiogenesis (Willam et al., 2002). Prolyl hydroxylase inhibitors may also prove to be useful in this regard (Ivan et al., 2002; Jaakkola et al., 2001). Activation of the HIF pathway may also be beneficial in ischemic preconditioning. Preconditioning by exposure to mild hypoxia has been suggested as a means to provide protection against subsequent challenge by stimulating angiogenesis or by preventing apoptosis, as is the case in the retina (Grimm et al., 2002, 2005). Exposure to lower oxygen levels or treatment with cobalt chloride induces HIF1 protein expression and oVers protection to the brain, heart, and retina in various animal models (Bergeron et al., 2000; Bolli et al., 1997; Gidday et al., 1994, 1999; Kietzmann et al., 2001; Ozaki et al., 1999; Schulz et al., 2001).

VII. HIF and Cancer A. Tumor Angiogenesis and Hypoxia The induction of angiogenesis is essential for tumor growth, survival, and progression to an invasive phenotype (Hanahan and Folkman, 1996). One of the critical factors responsible for mediating the activation of the so‐called angiogenic switch is the presence of hypoxia within solid tumors. As tumors proliferate beyond 1–2 mm3, oxygen and nutrients become limiting, resulting in the development of hypoxia. Tumor hypoxia correlates with aggressive disease and a poor prognosis in patients, and leads to resistance to radiation therapy, since this treatment requires oxygen free radicals to induce cell death (Hockel and Vaupel, 2001; Hockel et al., 1996, 1999). Cancer cells adapt to this hypoxic environment through the activation of a number of cellular pathways that stimulate glycolysis, genetic instability, invasion, and neovascularization. These processes provide the tumor with adequate energy and blood supply to allow for continued growth in the face of lower oxygen. As the master regulator of the hypoxic transcriptional response, HIF has been implicated as an important player in tumor growth and angiogenesis. HIF activates the expression of a broad range of genes that can contribute to tumorigenesis such as glucose transporters that support the enhanced rate of glycolysis in tumors, growth factors that stimulate proliferation, and metalloproteinases and chemokines that may influence metastasis and EC


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recruitment (Ben‐Yosef et al., 2002; Ceradini et al., 2004; Dang et al., 1997; Harris, 2002; Staller et al., 2003; Zagzag et al., 2005). Importantly, as mentioned earlier, HIF induces multiple genes involved in angiogenesis, most notably VEGF, which is essential for initiating and maintaining new vessel formation (Forsythe et al., 1996; Semenza, 2000).

B. HIF Expression in Tumors HIF1 is overexpressed in a variety of human cancers and correlates with highly vascular tumors (Brahimi‐Horn and Pouyssegur, 2005; Semenza, 2002). For example, HIF1 protein levels were increased in glioblastoma multiforme, prostate cancer, breast cancer, lung cancer, and pancreatic cancer (Bos et al., 2001; Giatromanolaki et al., 2001; Talks et al., 2000; Zagzag et al., 2000; Zhong et al., 1999). In addition, HIF1 is induced in a transgenic mouse model of epidermal carcinogenesis (Elson et al., 2000). HIF2 overexpression is also detected in these tumor types, with more pronounced expression than HIF1 in hepatocellular carcinoma (Bangoura et al., 2004; Talks et al., 2000). Furthermore, both subunits are expressed in RCC (see later), with progressively higher expression in more advanced tumors and a bias toward HIF2 expression (Mandriota et al., 2002; Turner et al., 2002; Wiesener et al., 2001). Immunostaining for both HIF subunits is predominantly nuclear and generally strongest in necrotic regions; there is often significant colocalization with increased levels of VEGF mRNA, especially in the case of HIF2 (Favier et al., 2001; Talks et al., 2000; Zhong et al., 1999). In several studies, HIF1 expression is associated with decreased survival and resistance to radiation therapy, suggesting that evaluation of HIF levels may have prognostic value (Aebersold et al., 2001; Brahimi‐Horn and Pouyssegur, 2005; Unruh et al., 2003). In addition to enhanced expression in tumor parenchymal cells, HIF2 is also upregulated in tumor‐associated macrophages (TAMs) within the stroma (Talks et al., 2000). TAMs are recruited to hypoxic, avascular regions of many diVerent tumors by growth factors and chemokines, and the degree of TAM infiltration correlates with tumor progression and poor prognosis (Leek and Harris, 2002; Leek et al., 2002; Pollard, 2004). For example, high expression of HIF2 in TAMs was shown to significantly correlate with high tumor vascularity and tumor grade in breast cancer and bladder cancer (Leek et al., 2002; Onita et al., 2002). Macrophages likely promote tumor growth and angiogenesis by secreting growth factors that stimulate tumor cell proliferation, MMPs that induce invasion and metastasis, and proangiogenic cytokines (Pollard, 2004). These angiogenic factors include basic fibroblast growth factor (bFGF), iNOS, angiopoietins, IL‐8, and VEGF, many of which are HIF target genes (Pollard, 2004). In particular, macrophage infiltration

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has been associated with increased VEGF expression, potentially through HIF activation, and tumor angiogenesis in several studies both in vitro and in vivo (Bingle et al., 2006; Leek et al., 1996, 2000; Lewis et al., 2000). VEGF in turn is a chemotactic factor for macrophages and monocytic endothelial precursor cells (TEMs), which may serve to further enhance vascularization within the tumor (Barleon et al., 1996; Grunewald et al., 2006; Leek et al., 2000). There are several mechanisms by which HIF is overexpressed in tumors, the primary of which is hypoxic activation. In addition, mutations in tumor suppressor genes, such as VHL (Section VII.C), p53, and PTEN, result in increased HIF1 and HIF2 (Maxwell et al., 1999; Ravi et al., 2000; Zundel et al., 2000). Conversely, activation of oncogenes, including v‐SRC, EGFR, and HER2neu, and subsequent signaling through the phosphatidylinositol‐3‐ kinase (PI‐3K) and mitogen‐activated protein (MAP) kinase pathways mediate HIF1 accumulation (Chen et al., 2001; Jiang et al., 1997a; Laughner et al., 2001). In each of these cases, activation of the HIF pathway promotes the production of angiogenic factors, such as VEGF, and therefore stimulates tumor angiogenesis (Harris, 2002; Semenza, 2000).

C. von Hippel‐Lindau Disease One of the principal lines of evidence supporting a role for HIF in tumor angiogenesis comes from analysis of the eVect of mutations in the VHL tumor suppressor gene. The hereditary cancer syndrome known as VHL disease is caused by germline mutations in VHL. Patients inherit an inactivating mutation in one allele of VHL and develop tumors on mutation or loss of the remaining allele in somatic tissue (Kaelin, 2002). The disease is characterized by a defined spectrum of highly vascular tumors that arise within specific tissues, including hemangioblastomas of the central nervous system and retina, pheochromocytoma aVecting the adrenal medulla, and RCC. In addition, VHL is also inactivated in a large percentage of sporadic hemangioblastomas and RCC through mutations, deletions, loss of heterozygosity, or methylation (Kaelin, 2002). Loss of pVHL function results in constitutive HIF activation throughout RCC tumors, even under normoxia (Wiesener et al., 2001). Enhanced expression of HIF target genes, including VEGF and TGF‐, mediates proliferation, angiogenesis, and increased vascular permeability in these tumors (de Paulsen et al., 2001; Gunaratnam et al., 2003; Kaelin, 2002). Cell lines derived from RCC tumors display high levels of HIF activity (especially HIF2), and the introduction of wild‐type VHL into these cells restores proper regulation of the HIF pathway and suppresses tumor growth (Iliopoulos et al., 1995).


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Most of the tumor‐associated mutations in VHL dysregulate HIF activity, either by disrupting binding of pVHL to HIF subunits or by impairing the formation of the complete E3 ubiquitin ligase complex (CliVord et al., 2001; Duan et al., 1995; Stebbins et al., 1999). Hydroxylated HIF is bound by the ‐domain of pVHL through several conserved residues; mutations of these amino acids abrogate this interaction with HIF and are associated with tumor development. The ‐domain of pVHL is responsible for binding to elongins B and C and formation of an active E3 complex, and mutations in this domain are also found in VHL disease (Duan et al., 1995; Kamura et al., 2000; Ohh et al., 2000; Stebbins et al., 1999; Tanimoto et al., 2000). In addition, an ‐helix at the C‐terminus of pVHL is important, although not required, for stabilization of substrate binding and thus for complete ubiquitination of HIF (Lewis and Roberts, 2004). However, additional mutations are required for Vhl‐mediated tumorigenesis, since although loss of Vhl in these cells resulted in constitutive HIF stabilization, Vhl/ tumors displayed a growth disadvantage (Mack et al., 2003, 2005). In addition, HIF‐ independent functions of pVHL, such as fibronectin matrix assembly, likely play an important role in tumor development (CliVord et al., 2001; HoVman et al., 2001; Ohh et al., 1998; Stickle et al., 2004). Several groups have tried to model VHL disease in the mouse in order to gain a better understanding of the mechanisms associated with tissue‐specific tumor development. Although mice completely lacking Vhl die during embryogenesis, targeted inactivation of one allele of Vhl in the liver resulted in the formation of cavernous hemangiomas that resemble hemangioblastomas as well as polycythemia (Gnarra et al., 1997; Haase et al., 2001). HIF2, but not HIF1, expression was detectable in these tumors, and VEGF mRNA levels were significantly elevated (Haase et al., 2001). HIF signaling was required for the development of both hemangiomas and polycythemia in this model. However, inactivation of Hif1a alone did not suppress these phenotypes, further suggesting the importance of HIF2 in VHL disease (Rankin et al., 2005). Another mouse model has been generated in which kidney‐specific deletion of Vhl enhanced renal cyst development (Rankin et al., 2006). Furthermore, mice expressing various mutant forms of Vhl associated with VHL disease have been generated; analysis of this allelic series should help elucidate the relative contributions of HIF‐dependent and HIF‐independent functions of pVHL to tumorigenesis (Rathmell et al., 2004; Rathmell, W. K., Hickey, M. M., and Simon, M. C., unpublished observations). D. The Role of HIF1a and HIF2a in Tumor Angiogenesis Inactivation of VHL and subsequent HIF activation is detected in the early stages of RCC development and is associated with increased vascularization

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in early foci, suggesting that HIF promotes angiogenesis in these tumors (Mandriota et al., 2002). In agreement with this, loss of Vhl in murine ES cells and murine embryonic fibroblasts (MEFs) resulted in constitutive HIF activation and increased VEGF expression. As a result, Vhl/ teratocarcinomas and fibrosarcomas derived from these cells were more hemorrhagic and exhibited a higher microvessel density compared to tumors derived from wild‐type cells (Mack et al., 2003, 2005). A number of mouse xenograft studies have investigated the role of the HIF1 subunit in tumor growth and angiogenesis. These studies obtained conflicting results with regard to the role of HIF1 in tumor cell proliferation and apoptosis, probably due to diVerences in the cell types used. Some groups found that tumors lacking HIF1 grew more slowly (Maxwell et al., 1997; Ryan et al., 1998, 2000; Tang et al., 2004), while others reported that loss of HIF1 enhanced tumor growth and decreased apoptosis (Blouw et al., 2003; Carmeliet et al., 1998). However, in each case HIF1 inactivation resulted in impaired tumor angiogenesis. For example, hepatoma cells deficient for HIF1 /ARNT formed less vascular, slow‐growing tumors with reduced VEGF levels compared to tumors derived from wild‐type cells (Maxwell et al., 1997). Deletion of HIF1 in ES cells also led to decreased VEGF expression and defective vascularization of tumors in nude mice (Carmeliet et al., 1998; Ryan et al., 1998). Similarly, Lewis lung carcinoma cells gave rise to necrotic tumors with significantly reduced microvessel density when injected into mice lacking HIF1 in ECs (Tang et al., 2004). Therefore, HIF1 likely promotes tumor vascularization by inducing the expression of VEGF as well as other angiogenic factors such as the angiopoietins and VEGF receptors. Importantly, this eVect is dependent on the tumor microenvironment; subcutaneous tumors derived from Hif1a‐ deficient astrocytes were poorly vascularized with extensive necrosis, while brain tumors derived from these cells exhibited increased microvessel density compared to wild‐type tumors (Blouw et al., 2003). HIF2 is highly expressed in ECs and neural crest cells, from which pheochromocytoma originates, as well as a number of tumors and RCC cell lines, suggesting that HIF2 may be the more important subunit in the pathogenesis of VHL disease and tumorigenesis in a broader sense. Several genes involved in tumor growth and angiogenesis, including cyclin D1, Glut‐1, TGF‐, and VEGF, were found to be specifically regulated by HIF2, not HIF1, in RCC cell lines (Raval et al., 2005). Expression of a constitutively stable form of HIF2, in which one of the critical proline residues was mutated to alanine, in VHL‐rescued RCC cells resulted in the formation of larger tumors in nude mice as compared to controls (Kondo et al., 2002, 2003). In contrast, stabilization of HIF1 alone by similar means did not induce tumor formation (Maranchie et al., 2002). Furthermore, downregulation of HIF2 with short hairpin RNAs (shRNAs) blocked the ability of VHL‐deficient RCC cells to


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form tumors in nude mice (Kondo et al., 2003). These findings demonstrate that HIF2 signaling is necessary and suYcient to induce tumorigenesis in the case of RCC cells. Further experiments are necessary to address the role of HIF2 in other types of tumors. Additional studies have reported that HIF2 contributes more significantly to tumor angiogenesis. Teratocarcinomas generated from ES cells in which Hif1a was replaced with Hif2a were larger in size with increased vascularization. This phenotype correlated with enhanced expression of genes such as VEGF, cyclin D1, and TGF‐ (Covello et al., 2005). Similarly, Acker et al. (2005) found that overexpression of HIF2 in glioma tumors stimulated VEGF expression, while inhibition of HIF2 resulted in decreased VEGF levels and significantly less vascular teratocarcinomas. Therefore, both HIF1 and HIF2 regulate tumor angiogenesis in response to hypoxia, although their respective contributions in the context of diVerent cell types needs to be further explored.

E. Therapeutic Implications The use of antiangiogenic agents to treat cancer was first proposed in 1971 by Judah Folkman, sparking a period of intense research in this area (Folkman, 1971). Despite some initial setbacks, the results of clinical trials have given new life to this idea and suggest that the inhibition of angiogenesis may in fact be a promising therapeutic strategy. For example, a monoclonal antibody against VEGF known as bevacizumab or Avastin has been shown to increase overall survival in metastatic colorectal and non‐small‐cell lung cancer patients when administered together with standard chemotherapy (Hurwitz et al., 2004; Kim et al., 1993). In addition, small molecule inhibitors, such as Sorafenib, have been developed that target multiple receptor tyrosine kinases, including the VEGF‐Rs and PDGF‐R, in both ECs and tumor cells. Treatment with Sorafenib has been shown to be eVective in patients with advanced RCC, resulting in a significant increase in progression‐free survival (Carmeliet, 2005; Ferrara and Kerbel, 2005). Other approaches to targeting tumor angiogenesis are also being developed and are in the early phases of clinical trials, including a soluble, chimeric VEGF receptor known as the VEGF‐trap, and recombinant human endostatin, an endogenous inhibitor of angiogenesis (Herbst et al., 2002; Holash et al., 2002). The success of these studies has led to the suggestion that the inhibition of upstream regulators of angiogenesis may also be eVective in the treatment of cancer. Given the significant role of HIF activity in promoting tumor growth and angiogenesis, this pathway represents an exciting target for therapeutic intervention. Modulation of HIF signaling, perhaps in combination with

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more traditional chemotherapy and/or other more direct antiangiogenic treatments may provide eYcient inhibition of tumor vascularization and proliferation. However, it will be essential to take into account the varied and complicated eVects of HIF on gene expression, such as its role in regulating apoptosis, when designing novel cancer therapies. One possibility that has been proposed is the use of HIF1 antisense therapy, which blocked T‐cell lymphoma growth in combination with T‐cell‐specific activation (Sun et al., 2001). Antisense therapy against HIF also synergized with overexpression of VHL in gliomas to increase tumor cell apoptosis and decrease tumor angiogenesis (Sun et al., 2006). In addition, blockade of the interaction between HIF and its coactivators CBP/p300 by polypeptides corresponding to the HIF1 transactivation domain suppressed hypoxia‐inducible gene expression and attenuated tumor growth in a xenograft model (Kung et al., 2000). Tumor‐specific death may be achieved by using gene therapy systems in which HREs drive the expression of proapoptotic or antiproliferation genes (Dachs et al., 1997). Alternatively, hypoxic tumor cells can be specifically targeted by anaerobic bacteria expressing prodrugs that inhibit tumor growth (Agrawal et al., 2004; Bettegowda et al., 2003; Dang et al., 2001; Lemmon et al., 1997). Macrophages engineered to express adenoviral vectors may also be useful for targeting therapeutic genes and drugs to hypoxic tumors (GriYths et al., 2000). Inhibition of HIF could also enhance the response of tumors to radiation therapy. Irradiation has been shown to induce HIF1 through the production of ROS, resulting in the expression of VEGF and enhanced survival of ECs (Moeller et al., 2004). Inhibition of HIF1 with a small molecule, YC‐1, slowed tumor growth following irradiation (Moeller et al., 2004; Yeo et al., 2003). However, HIF can also sensitize tumors to radiation through the activation of p53‐dependent apoptosis, emphasizing the complex eVects of HIF on tumors. Therefore, radiation treatment followed by HIF blockade may prove to be an eVective cancer therapy (Moeller et al., 2005).

VIII. Conclusions The HIF pathway is an essential eVector of the cellular response to changes in oxygen concentration. Activation of HIF under hypoxia results in the induction of a broad program of gene expression that is necessary for adaptation to low oxygen. HIF signaling mediates the hypoxic response through cell autonomous mechanisms, for example, by regulating proliferation, apoptosis, and metabolism, as well as through cell nonautonomous eVects on angiogenesis. HIF‐stimulated expression of proangiogenic factors, such as VEGF, angiopoietins, VEGF receptors, MMPs, and IL‐8, is critical


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for the formation of new blood vessels in both physiological and pathophysiological settings. HIF signaling is required for proper embryonic development, particularly in the formation of the vasculature, and is important for the function of a variety of cell types in the adult. In addition, HIF expression and activity are increased under conditions of pathological hypoxia thereby leading to enhanced vascularization. Therefore, modulation of HIF represents a potential therapeutic target that may have benefits in the treatment of both ischemic disease and cancer. It is hoped that ongoing and future studies will help to further elucidate the mechanisms of HIF regulation and the importance of the hypoxic response in both normal physiology and disease.

Acknowledgments We thank members of the lab, especially Dr. Brian Keith, for helpful discussions and proofreading assistance. We apologize to those colleagues whose work could not be cited directly.

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Index A Acetyltransferase activity, 150 Achondroplasia (ACH), 6 Acidic-Aromatic motif, in Bik1p/CLIP-170 proteins, 58–59 Activin, 13 pathways, 16, 18, 20 Activin receptor-like kinase (ALK) 1, 9–10 complexed with TGF- RII, 10 ActRI, 9 ActR-IB, 9 Adenamotous polyposis coli (APC), 105 Adenoviral overexpression of noggin, 11 Adipocytes, 116 progenitors, 117 reduction of, 117 Adrenomedullin (ADM), 226 Aldolase A, 226 Alf1p, 51 ALK1. See Activin receptor-like kinase (ALK) 1 ALK2, 9–10 loss of, 25 ALK3, 9 ALK4, 9 ALK5, 9 ALK6, 9 ALK7, 9 Alkaline phosphatase (ALP), 117 activity, 118 Alzheimer’s disease, 89–90 Angiogenesis, 217–218 control of, 220 mechanisms of, 219 Angiogenic factors, 233, 235 Angiogenic switch, 220, 234 Angiopoietin(s), 220, 231, 235 receptor tie-2, 221 Angiopoietin-1 (Ang-1), 220 Antagonists, 11 Antiapoptotic consequences, 95 Antiapoptotic function, 117

Antiresorptive treatments, 104 Antisense therapy, 240 AP-1, 8 Apoptosis, 2, 28 proliferation in, 117 Apoptotic osteoblasts, 117 Archaea, 139 Arnt gene, 228 Arrow, 106–107 ARS1, 142 Articular cartilage, 14, 32, 35 Articular chondrocytes, 31–32 Articular region, 32 Ase1p protein, 77 ASF, 159 ASF-mediated nucleosome deposition, 159 ASF1 proteins, 160 ATDC5 cells, 24–25, 28, 34 chondrosarcoma cells, 21 ATP hydrolysis, 132 ATX-2 activity, 209 Autophagic activity, 96, 98 Autophagic processes, 92 Autophagosome-like structures, 97 Autophagosomes, 91, 97 Autophagy, 90–92 in cell culture, 94 chaperone-mediated, 91 induction, 95 lysosomal pathways, 89 lysosome routes, 91 regulation, 96 upregulation, 90, 97 Autosomal dominant brachydactyly type C, 22 Autosomal dominant HBM trait, 113 Autosomal dominant sclerosteosis, 111 Avastin, 239 Axin, 105 protein, 108 259

260 B BAC transgenic approach, 16 Basic fibroblast growth factor (bFGF), 235 Basic helix loop helix-Per ARNT Sim (bHLH-PAS) proteins, 221–222 Bevacizumab, 239 BFA1 gene, 64, 76 Bik1p-GFP localization, 70 Bik1 protein, from Saccharomyces cerevisiae, 50. See also Microtubule plus-end tracking proteins activities of, 61–63 microtubule-binding activity, 62 microtubule-stabilizing activity, 62 domains of, 51 genomic studies of, 76 gene expression levels, 78 quantitation of Bik1p molecules in cell, 78 synthetic lethal interactions, 77–78 interactions with other proteins, 60–61 at kinetochore, 76 phenotypes of, 61–63 regulation by phosphorylation, 75 role in spindle positioning, 63–64 in dynein pathway, 64–66 in Kar9p pathway, 66–69 in mating cells, 69–70 structure of, 50 cargo-binding domain, 56–60 coiled-coil domain, 55, 56 head domain, 51–55 Bik1 protein, head domain of CAP-Gly domain, 51–55 serine-rich region, 55 Bim1p protein, 60 in Kar9p pathway, 66–67 Blast colony-forming cell (BL-CFC), 219 BMP. See Bone morphogenetic proteins BMP2, 14 activity of, 12 -mediated cartilaginous nodule, 19 promoter activity, 118 sequestering ligands, 11 BMP4, 14, 34 binding to BMPRIA, 10 binding to BMPRIB, 10 promoter activity, 28 BMP5, 14 expression of, 16 BMP7, 12, 14 promoter activity, 28

Index Bmp2 expression, 31 Bmp4 expression, 30 Bmp6 expression, 31 Bmpr1b-null mice, 19 Bmpr1b transcription, 16 BMP receptors, 12 interactions of, 32 type I, 15 type IA, 9 type II, 15 BMPRIA, 9–10, 15, 19, 23–25 allele of, 20 loss of, 25 BMPRIB, 9, 13, 15, 19–20, 25 expression, 16 loss of, 25 mutations in, 22 signaling, 22 BMPs 5, 6, and 7, interrelated, 10 BMPs 2 and 4, interrelated, 10 BMPs chondrogenesis, 16 BMP signaling, 12–13, 18–21, 29, 32, 34 activity, 31 cascades, 8, 32 coreceptors for, 10 downstream eVectors of, 25 extracellular modulators of, 11 pathways, 8, 12, 23, 31, 33 regulation of, 10 transduction pathways, 8 BMP type I receptors, 13 Bone formation, 104, 121 rate, 114 fractures, susceptibility to, 114 geometry, 119 growth, 104, 109 homeostasis, 115, 118 mass, 105, 114–115, 117 matrix-secreting activity, 105 mineral density, 113 remodeling, 104, 118–119 repair, 104 shape, 119 Wnt Signaling in, 120 Bone marrow, 221 macrophages, 223 Bone morphogenetic proteins (BMPs), 2–3, 5, 7–8, 10, 13, 103–104

Index antagonists, 11 in chondrogenesis, 19 coreceptors for, 10 -dependent activation, 27 in early chondrocyte diVerentiation, 21 high-aYnity complexes with, 11 -induced gene expression, 27 ligands of, 9 mediated chondrogenesis, 11 -mediated signaling, 26 morphogens of, 118 pathways, 10, 16, 18, 20, 25, 28, 32 antagonistic interactions, 30 FGF-mediated antagonism of, 31 intracellular elements of, 34 maturation, 24 proliferation, 24 role for, 14 Smad pathways, 26–27 Bone morphogens, 117 Brachydactyly, 13 type A2, 22 Bradycardia, 230 Bromodomain motif, 132 B-type cyclin. See Clb5p BUB2 gene, 64 C Caenorhabditis elegans, 186–187, 192, 218 germ line, 194, 197 CAF-1, 159–160, 162 activity, 158 CAG trinucleotide repeat expansion, 90 Cancer, 217, 221 Canonical GLP-1 Notch cascade, 210 Canonical Notch pathway, 194 Canonical pathways, 7, 105–106 Canonical Smad pathway, 8 Canonical Wnt cascade, 115 Canonical Wnt pathway, 111 CAP-Gly domain of BiK1p and CLIP-170 proteins, 51–55 structure and conservation of, 54 Carbamazepine (CBZ), 96 Cardiac failure, 230 Cardiac myocytes, 223, 231 Cardiovascular abnormalities, 229–230 Cargo-binding domain, of Bik1p and CLIP-170, 56–60, 76

261 Cartilage(s) articular, 14–15 condensations, 20 hypertrophic, 14 overgrowth, prevention, 23 Ca2þ signaling pathway, 106 Catecholamines, production of, 230 -Catenin, 7, 105, 114 inactivation complex, 108 phosphorylation, 105, 108 signaling, 106 stabilization, 120 CBP/p300, 9 C2C12 culture, 12 CD44, 12, 34 cytoplasmic domain of, 12 Cdc6, 149 phosphorylation of, 146 Cdc45, 152 complexes, 153 CDK, 146 pathway, 152 phosphorylation by, 152 Cdt1, 149 Cell–cell interactions, 19 Cell fusion, in mating, 70 Cellular morphology, 4 Cellular organelles, 218 Cephalic mesenchymal cells, apoptosis of, 230 Cerebellar hemangioma, 237 Chaperone-mediated autophagy, 91 Chaperone protein Mesd, 112 Chick chondrocytes, 24 Chondroblasts, 4 Chondrocyte(s), 2–3, 6, 10, 13, 18, 115–117 BMP signaling in, 23 characteristic of, 21 development of, 15 diVerentiation, 7, 25, 27, 31 progression of, 24 hypertrophic, 14 life cycle, 8 markers, 6 hypertrophic, 24 phenotype, 20 proliferating, 24, 33 proliferation, 5, 14, 28–29, 31 control of, 28 PZ, 14 terminal diVerentiation of, 24

262 Chondrocytic genes, 3 Chondrodysplasia, 6, 22, 31 acromesomelic, 22 Grebe-type, 22 Chondrodysplasias, human analysis of, 22 Chondrogenesis, 2–3, 6–7, 13, 16–19, 23, 25, 31 BMP mediated, 11 control of, 33 early stages of, 21 Smad-mediated signaling in, 25 Chondrogenic condensations, 21–22 Chondrogenic diVerentiation, induction of, 28 Chondrogenic lineage, 20, 23 Chondrogenic markers, inhibits expression of, 11 Chondroitin sulfate (CS), 12 Chondroitin-4-sulfotransferase 1 gene, 12 Chondroprogenitor cells, 22 Chordin, 11 expression levels, 11 Chordin-null mice, 11 Chromatin, 148–149, 154, 158 accessibility complex (CHRAC), 132 assembly, 160 assembly factor-1 (CAF-1), 157 binding, 131 protein, 135 protein HP1/Swi6, 135 fiber, 130, 134 prereplication complex, 144 proteins, 135, 156 remodelers, 133–135, 139 remodeling, 130–131, 133, 143 complexes CHD, 156 CHRAC, 142 ISWI, 142 remodeling ATPases, 157 template, 155–156 Chromodomains, 132, 136 Chromosomal proteins, 139 Chromosome segregation, 130 C3H10T1/2 cells, 25 Ci, mammalian orthologs of, 118 Circadian rhythms, 221 Clb5p and Kar9p pathway, 67 Clb5p/Cdc28p kinase, 67 Cleidocranial dysplasia, 104

Index CLIP-170 proteins, mammalian, 70. See also Bik1 protein; Microtubule plus-end tracking proteins CAP-Gly domains of, 51–55 C-terminal domain of, 65 interactions with EB1 proteins and, 74–75 interactions with other proteins, 60–61 regulation by phosphorylation, 75 CLIPR59 protein, 51 Cohesion proteins, 132 Coiled-coil domain, in Bik1 protein, 55–56 Col2a1-Cre transgene, 17, 20 Col2a1 gene, 34 Col2a1 promoter, 17 Col2-Cre phenotypes, 18 Collagen(s), 11 expression type II, 24 type X, 24 I and osteopontin, 117 II, 4 IX, 4 production, type II, 21 promoter, type X, 26 X, 4, 6 XI, 4 Columnar zone (CZ), 4–5, 32 ColX promoter, 28, 30 Condensed chondroprogenitors, 23 Constitutively active (CA), 6 Copolymerization-based treadmilling mechanism, 73 Coronary heart disease (CHD), 135 Craniofacial bones, 2 Craniofacial dysostosis, 104 CREB binding protein (CBP), 222 CSL transcription factors, 192–193 Cubitus interrupts (Ci), 118 Cyclin-dependent kinases (CDKs), 129, 144, 151 Cytokines, 63–64, 223, 226, 235 Cytoplasmic complex, 105 Cytoplasmic domain of CD44, 12 Cytoplasmic polyadenylation element-binding (CPEB) protein, 205 Cytoplasmic vacuoles, 109 Cytoprotective eVects, 95 Cytoskeleton, 13 Cytosolic peptidases, 93 CZ. See Columnar zone

263

Index D DAN, 11 DDK, 151 DDK pathway, 152 Dementia tauopathy, 90 Dentatorubral pallidoluysian atrophy (DRPLA), 90 Deubiqitinating enzymes, 93 Deubiquitinylating enzyme, 134 Diabetes, 221 4,6-Diamidino-2-phenylindole (DAPI), 161 Dickkopf (Dkk) proteins, 106–107, 110 DiVerentiating proliferative cells, 196 Distal tip cells (DTCs), 188 DNA binding, 131 DN-ALK2, 19 DNA polymerase, 155 DN-type I BMP receptor, 21 Dominant HBM trait, 113 Dominant negative (DN)-BMPRIB, 19 DRAGON, 10 Drc1/Sld2 protein, 151 Drosophila, 139, 141, 143, 148, 158, 160, 186, 197 germ lines, 187, 196 homologue, 107. See also Drosophila (Arrow) melanogaster, 53, 218, 224 models, 89, 94 mutant, 107 Nanos, 194 Rpd3 HDAC, 143 translational regulator, 202 Drosophila (Arrow), 107 DTC, 192 membrane, 199 Du Pan syndrome, 22 Dynein pathway, in spindle positioning, 63–66 E ‘‘Early pathway.’’ See Kar9p pathway EB1 homologue. See Bim1p protein EB1 proteins, interactions with CLIP-170 proteins, 74–75 ECM, 4, 12, 32 proteins, 220 Ectodermin, 13 Ectopic cartilage, 7, 21

Ectopic chondrogenesis, 19 Ectopic joint formation, 21 EGF pathways, antagonistic interactions, 30 Embryogenesis, 186 LRP6, 109 of the skeletal system, 104 Embryonic angiogenesis, 229–230 Embryonic development, 109–110, 116–117, 217, 219, 228, 241 Embryonic lethality, 17 Embryonic tissues, 229 Endochondral ossification, 27 Endogenous inhibitors, 105 Endothelial cell (EC) precursors, 220 Endothelial progenitor cells (EPCs), 221 Enzyme, deubiquitinylating, 134 Epidermal growth factor (EGF), 223 domain, 107 Epigenetic inheritence, 135 Epo expression, 232 ERK1/2, 10 activity, 31 inhibition of, 30 phosphorylation by, 30 ERK1/2-mediated inhibition, 31 E3 ubiquitin ligase complex, 237 Euchromatin, 130, 135 Excitotoxicity, 95 Exon 1 fragment, 93 Expression of marker genes, 4 Extracellular matrix (ECM), 4, 11, 220 mineralization of, 115 Extraembryonic tissues, 229 Eye abnormality, 109 Eye vascularization, 112

F Fab1p, 78 FAST-1, 8 Fbf-1 fbf-2 double-mutant males, 204 FBF functions, 202 Fbf mutant, 205 Fbf mutations, 207 FBF-1 protein, 206 FBF-2 protein, 206 Fbf-1 single mutants, 204 Fem-3 bindinding factor (FBF)-1, 202 FGF, 5 FGF2, 6 FGF activity, 31

264 Fgf18 expression, 30 FGF-mediated inhibition, 6 FGF pathways, 7 FGFR3, 6 Fgfr3, 6 expression, 30 Fgfr3ACH, 6 FGF signaling, 6, 30 Fibrillin-1 deficient mice, 12 mutations in, 12 Fibrillin-2, 12 Fibrillins sequester, 12 Fibroblast growth factor (FGF), 2, 220 Fibronectin, 229, 237 FOG-1 protein, 205 Follistatin, 11 Fracture healing, 118 Frizzled (Fz) receptors, 105–106 Frontotemporal dementia, 94 Frontotemporal tauopathy, 90, 94 G GADD45 , inhibition of, 27 GBP/Frat1 protein, 108 Gcn5, 149–150 Gcn5 HAT, 143 GDF5, 14 Gdf5, mutation in, 22 Gdf6, expression of, 16 GDF5 protein, mutated, 22 GDF5 signaling, 13 Gene encoding sclerostin (Sost), 105 Genes, for regulating exit from mitosis, 64 GINS, 153 Gld-1; fbf-1 fbf-2 null triple mutants, 206–208 GLD-1 activity, 205 Gld-1 gld-2 null mutant adults, 208 Gld-1 gld-2 null mutant larvae, 208 Gld-2; glp-1 double mutant, 205 GLD-1 levels, 201–202 Gld-1 null mutant animals, 199 Gld-2 pathway, 204 GLD-1 pathways, 186, 195–196, 200, 202, 208 activity, 199 GLD-2 pathways, 186, 195–196, 200, 202, 206–208 activity, 199 GLD-1 protein, 207–208 levels, 204

Index Gli1, mammalian orthologs of, 118 Gli3, mammalian orthologs of, 118 Gli proteins, 30, 118 Gli3 proteins, 30 GLP-1 receptor, 199 signaling, 195 Glp-1 activity, 209 GLP-1 Notch signaling, 200, 202, 204, 206, 208–210 ternary complex, 199 Glucose transporter-1 (Glut-1), 218 Glycine/serine-rich (GS) receptor, type I, 8 Glycogen synthase kinase (GSK)-3beta, 105 Glycolytic enzymes, 218, 226, 229, 233 Glycoproteins, 105 GNAT, 149 G-protein–coupled receptor (GPCR), 106 Gremlin, 11 Growth and diVerentiation factors (GDFs), 10 Growth plate BMP actions, 23 maturation, 8 proliferation, 8 GSK-3 , 109 inhibition of, 105 inhibitor lithium chloride, 117 inhibitors of, 115, 120 H Hair follicle morphogenesis, 118 HAT, 134, 144, 149–150 HBM. See High bone mass Hbo1, 149 HD, 95 HDAC Rpd3, 143 HDAC Sir2, 148 HD pathogenesis, 90 HD patients, 97 Hedgehog (Hh) family of proteins, 104 morphogens of, 118 Hedgehog (Hh) signaling, 103, 118 Hemangioblasts, 219 Hematopoiesis, abnormal, 229 Hematopoietic cells, 219, 221, 229 Hematopoietic defects, 228 Hematopoietic stem cell diVerentiation, 231 Heparan sulfate, 12 Heparin sulfate proteoglycans (HSPGs), 11

Index Hermaphrodite germ line, 188, 195, 208, 210 Heterochromatin, 130, 135, 138–139, 158–160, 162 protein 1 (HP1), 135 proteins, 137, 151 replication of, 132 Heterochromatization, 136 Heterodimeric protein complexes, 137 Heterozygous humans, 104 HFE2, 10 H3-H4 chaperone, 159 HIF. See Hypoxia-inducible factor Hif1a, 231 in angiogenesis, 231–232 deficiency in adult tissues, 231 deletion, in cardiac myocytes, 231 HIF1, 222, 231, 235 in adult tissue, role of, 231–232 subunit, hydroxylation of, 224 Hif2, 232 deficiency, in mice, 232 HIF2, 231–232, 235, 238–239 mRNA, detection of, 223 in renal Epo expression and hematopoiesis, role of, 232 subunit, hydroxylation of, 224 Hif2a deficiency, 230 Hif1a-deficient embryos, 230 HIF proteins, polyubiquitination of, 224 HIF signaling, 227, 229, 237–240 HIF-stimulated erythropoietin (Epo) expression, 218 High bone mass (HBM), 105 human mutations, 115 mutation, 111–112 phenotype, 106, 113 trait, 103 HIR proteins, 159–160 Histone(s), 133, 154, 156, 158 acetylated, 134 chaperone-deposition complex, 159 chaperones, 157 deposition, 130 methylated, 135 methylation of, 134 modifications, 130, 133, 135 modifiers, 139 octamer, 132 phosphorylated, 133 Histone acetyltransferases (HATs), 133 Histone-binding proteins, 157

265 Histone deacetylases (HDACs), 134 Histone–DNA interactions, 131 Histone methyltransferases (HMTs), 134 H3K9, methylated, 136 H3K9Me residue, 136 H3K9 methyltransferase, 162 Homeobox transcription factors, 3 Hormone-related peptide (PTHrP), 2 Hoxc-8, 8 HP1/Swi6, 136 H3S10, phosphorylation of, 136 Human Orc1, 144 Humans, phenotypes of, 17 Huntingtin, 90, 93 Huntington’s disease (HD), 89–90 Hypertension, 221 Hypertrophic cartilage, 14 Hypertrophic cells, 5 BMPs in, 24 Hypertrophic chondrocytes, 5, 7, 14–15, 26–27 apoptosis of, 33 Hypertrophic diVerentiation, 5, 25 promotion, 24 Hypertrophic zone (HZ), 4–5, 7 Hypertrophy, 28 Hypoxia, 217, 219 Hypoxia-inducible factor (HIF), 217–218 activity, regulation of, 223–225 and cancer HIF1 and HIF2 in tumor angiogenesis, role of, 237–239 HIF expression in tumors, 235–236 therapeutic implications, 239–240 tumor angiogenesis and hypoxia, 234–235 von Hippel-Lindau disease, 236–237 in developmental angiogenesis, 228–231 hypoxic activation of, 223 and ischemic injury, 232–234 pathway, 218 proteins, 221 role in adult tissues, 231–232 subunits, 222 target genes, 219, 224–228 Hypoxia response elements (HREs), 222 Hypoxic regions, 218 Hypoxic response, 218, 223, 226, 228, 231, 240 Hypoxic stimulation, 223 HZ. See Hypertrophic zone

266 I IGF, 5 IHH. See Indian hedgehog Ihh, 6, 11, 27, 34 expression, 30 IHH/PTHrP pathways, 28 IL-8, 231, 235 Imitation SWItch (ISWI), 132 Immunoprecipitation, 29 Immunosuppression, 96 IMPase inhibition, 96 Indian hedgehog (IHH), 2, 4–6, 118 receptors, 34 Inducible nitric oxide synthase 2 (iNOS), 226–227, 235 Inflammatory cytokines, 226 Inhibitor Smads (I-Smads) 6, 13 Inhibitor Smads (I-Smads) 7, 13 Inhibitors of TGF- superfamily members, 14 INO80 family, 133 Inositol monophosphatase (IMPase), 96 Inositol monophosphate (IP1), 96 Insulin, 223 Insulin-like growth factor (IGF), 2 Insulin-like growth factor-2 (IGF-2), 226 Interleukin-1 , 223 Interleukin-8 (IL-8), 226 Interphalangeal joints, loss of, 21 Intracellular protein aggregation, 89–90 misfolding, 89–90 Intracellular proteinopathy substrates, 90 Intracellular regulation, 12 Intracytoplasmic aggregate-prone proteins, 93 Ischemia, 217 Ischemic injury, 219 Ischemic injury and HIF, 232–234 J JNK. See Jun kinase Joint formation, 8 Jun kinase (JNK), 10, 13 K Kaposi’s sarcoma, 143 Kar3p-GFP localization, 70 Kar9p pathway, in spindle positioning, 63–64, 66–69

Index Kidney epithelial cells, 223 Kielin, 11 Kinase inhibitor, 95 p21, 226 Kinetochores, Bik1 protein at, 76 KIN4 gene, 64 Kip2p protein, 65–66, 73

L Lactate dehydrogenase A (LDHA), 226 LAG-1-binding sites, 202, 206 LAG-2 ligand, 200 LAG-1 transcription factor, 197 ‘‘Late pathway.’’ See Dynein pathway Lateral plate mesoderm, 2 LDLR family, 107 Lewis lung carcinoma cells, 238 Limb mesenchyme, 20 Lipoprotein receptor-related protein (LRP)5, 103, 105 overexpression of, 113 LIS1 gene, mutations in, 65 Lithium chloride, GSK- inhibitor, 117 Live cell analyses of Bik1p, 69 of dynein mutants, 64 Liver parenchyma, 223 Low-density lipoprotein receptor (LDLR), 106 Lower bone mass, 111 LRP5. See Lipoprotein receptor-related protein 5 LRP5/6, 107 function, 107 mutants, 112 receptors, 111 structure, 107 Lrp5 deficiency, 109 Lrp6 deficiency, 109 Lrp5-deficient mice, 112 Lrp6-deficient mice, 112 Lrp5 gene, 112 Lrp5 G171V, 111 transgenic mice, 117, 119 Lrp6 haploinsuficiency, 112 L-Sox5, 20 LTE1 gene, 64 Lymphoid enhancing factor (Lef) transcription factors, 105 Lysosomal hydrolases, 92

Index M Macroautophagy, 89, 91 MAML transcriptional coactivator, 192 Mammalian autophagy, 96 Mammalian target of Rapamycin (mTOR) pathway, 92 MAPK/ERK kinase 1 (MEK1), 6 MAPK pathways, 6, 8 Marfan syndrome, 12 Mating cells, Bik1 protein in, 69–70 Matrix metalloproteinase 13 (MMP13), 5 Matrix metalloproteinases (MMPs), 220 MC615 chondroprogenitor cells, 25 MCM. See Mini chromosome maintenance complexes MCM-Hbo1, 150 Mechanostransduction, 115 Meiosis-promoting activity, 195 Meiotic prophase progression, 195 MEN. See Mitotic exit network Mesenchymal cells, 2, 19 apoptosis of, 22 condensation, 7 lines, 17 C3H10T1/2, 117 Mesenchymal condensations, 7, 109 Mesenchymal pluripotent cell lines, 117 Mesenchymal pluripotent cells, 119 Mesenchymal precursors, 104 Mesodermal cells, 220 Mesodermal progenitor cells, 219 Metabolic disorders, 109 Metal-binding motifs. See Zinc knuckle motif Metalloproteinase, 11 Metazoan Hbo1, 149 Mice -catenin–deficient, 114 chordin-null, 11 fibrillin-1–deficient, 12 with genetrap mutations, 12 heterozygous, 231, 233 lacking Bmpr1a, 24, 26 lacking Bmpr1b, 21–22, 24, 26 lacking Gdf5, 21 lacking noggin, 11 lacking Smad2, 26 lacking Smad4, 26 lacking Tgf r2, 26 Lrp5 deficient, 112 Lrp6 deficient, 112 Lrp5 G171V transgenic, 117

267 phenotypes of, 17 Tcf-deficient, 114 Microautophagy, 91 Micromass cultures, 28 Microtubule-associated genes, in Kar9p pathway, 77 Microtubule-binding activity, of Bik1p, 62, 70 Microtubule depolymerization, in mating cells, 69–70 Microtubule plus-end tracking proteins (þTIP), 74–75. See also Bik1p protein; CLIP-170 protein, mammalian behavior of, 70–71 mechanisms of behavior of, 71–74 end-loading, 71, 72, 73 hitchhiking, 73 motor-based transport, 71, 72, 73 surfing, 71, 72, 73 Microtubule-stabilizing activity of Bik1p, 62 Mineralization, 117 of MC3T3-E1 cells, 118 Mini chromosome maintenance (MCM) complexes, 146, 148–149 binding to chromatin, 147 DDK phosphorylation of, 152 helicase, 146 helicase complex, 154 proteins, 151 Mitochondrial-dependent cytochrome c, 95 Mitochondrial dysfunction, 232 Mitochondrial pathway, 95 Mitogen-activated protein kinase (MAPK), 6 Mitotic exit network (MEN), 64 MMP. See Matrix metalloproteinases MMP-2, 231 MMP-9, 231 MMP13, 6 MOG proteins, 209 Mouse Genome Informatics, 4 Mouse models, 89, 94 mRNA levels, 202 metabolism, 209 regulators, 195, 210 regulatory pathway GLD-1, 194 GLD-2, 194 regulatory pathways, 186, 195 stabilization, 221 mTOR, 94 enzyme, in CLIP-170 phosphorylation, 75

268 mTOR-independent autophagy enhancing pathway, 96 mTOR-independent autophagy pathway, 95 Multicellular organism, growth of, 218 Murine embryonic fibroblasts (MEFs), 238 Murine fibroblasts, 11 Mutant(s) gld-1; fbf-1 fbf-2 null triple, 206–207 huntingtin, 89–91, 94 polyglutamine proteins, 98 Mutations in fibrillin-1, 12 Myocardial ischemia, 221 Myocytes, 116 Myo-inositol-1,4,5-triphosphate (IP3) levels, 96 Myo2p, 66 Myosin-guidance model, 67 MYST, 144 N N-cadherin, 19 inhibitors, 19 Ndel1 protein, 65 Ndel2 protein, 65 Ndl1p protein, 65 Neoplastic disease, 218 Neurodegeneration, 89 Neurodegenerative diseases, 89–91, 95 Neurodegenerative disorders, 96 Nip100p protein, 51 Nitric oxide synthases (iNOS), 226 Nodal pathways, 16, 18, 20 Noggin, 11, 23 adenoviral overexpression of, 11 mice lacking, 11 Noncanonical pathway, 105 Nonhypoxic factors, 223 NOS-3 functions, 204 NOS-3 levels, 202 Notch receptor, 200 Notch signaling, 208 pathway, 194 Nuclear translocation, 12 Nucleosome, 132 Nucleosome assembly, 156 O OAZ, 8 Oct-4 expression, 231

Index Oct-4 levels, 231 Ocular structure, disruption of, 111 Okazaki fragments, 154 OPPG. See Osteoporosis pseudoglioma OPPG human mutations, 115 OPPG mutations, 111 OPPG phenotypes, 106, 113 ORC. See Origin recognition complex Origin recognition complex (ORC), 138–139, 149, 151, 160 Ossification, 8 Osteoarthritic articular chondrocytes, 32 Osteoarthritis, 31–32 progressive, 18 Osteoblasts, 2, 4–5, 7, 103, 111, 115 diVerentiation, 115, 117–118 diVerentiation pathway, 104 eVects of Wnt, 116 function, 117–118 lineage, 115 lineage, development of, 118 lineage, marker genes of, 104 maturation, 115 mature, 118 precursors, 104 progenitors, 120 proliferation, 117 sensitivity of, 13 Osteoblast-specific APC deficiency, 114 Osteocalcin expression, 29 Osteochondral progenitor(s), 2 cells, 3 Osteochondroprogenitor lineage, 20 Osteoclasts diVerentiation, 114–115 Osteocytes, 115 Osteogenesis, 119 Osteopenia, 114 Osteopetrosis type I syndromes, 111 Osteopontin (OPN), 6 Osteoporosis disorders, 109 susceptibility to, 104 treatment, 120 treatment of, 104 Osteoporosis pseudoglioma (OPPG) syndrome, 103, 105, 109 Osteoporotic patients, bone mass in, 105 Osteoprogenitor cells, 118 Osteoprogenitors, 118 Osteoprotegrin (OPG), 115 Osteosclerotic phenotypes, 105

269

Index Overt chondrocytes, maturation of, 17, 22 Oxidative phosphorylation, 218 Oxygen-dependent degradation domain (ODD), 223, 225 Oxygen homeostasis, 217 Oxygen-sensing pathway, 229 P Pac1p, 60 Pac2p, 51 Pancreatic parenchymal cells, 223 Parathyroid, 2 Paraxial mesoderm, 2 Parkinson’s disease, 89–90, 94 PCAF, 150 PCNA, 154, 159 PDGF, 220 Periarticular cells, 5 Pericentromeric heterochromatin, 142 Perichondrial cells, 4 Perichondrial signaling, 18 Perichondrium, 3, 5–6, 14, 17, 25, 33 Perichondrium flanking, 5 Perinatal lethality, 109 Periosteum, 5, 14 Phalangeal elements, loss of, 22 PHD. See Prolyl hydroxylase domain PHD enzymes, 222, 224 Pheochromocytomas, 224 Phosphoglycerate kinase (PGK), 226 Phosphorylate receptor, type II, 8 Phosphorylation, 31, 67 bik1p protein regulation by, 75 CLIP-170 proteins regulation by, 75 of p38, 28 Plasminogen activator inhibitor-1 (PAI-1), 226 Platelet-derived growth factor (PDGF), 220 Pluripotent mesenchymal cell lines, 19 p38 MAPK kinase pathways, 10 Polyalanine-containing proteins, 94 Polyglutamine aggregates, 94, 98 -containing proteins, 94 diseases, 89 expansion, 89 tract, 94 Polyglutamine (polyQ) tract, 90 Postnatal bone accrual, 118 Posttranscriptional modes of action, 34

Postvasculogenesis, 230 PR39, 234 Precartilaginous cells, 21 Prechondrogenic cells, 7 Prechondrogenic condensations, 20, 25 Prechondrogenic mesenchyme, 18 Prehypertrophic cells, 4 Prehypertrophic chondrocytes, 4–5, 26–28 Premature diVerentiation, 28 Preosteoblasts, 5 preRC. See Prereplication complex preRC proteins, 148 Prereplication complex (preRC), 129, 144 acetylation of, 150 Proangiogenic HIF target genes, 226 Proangiogenic macrophages, 231 Proapoptotic gene Bnip3, 226 Proapoptotic signals, 6 Prodrugs, 240 Proliferating cells, 197 Proliferative zone (PZ), 4 Prolyl hydroxylase domain (PHD) proteins, 223–224 -Propeller, 111 Proteasome pathways, 89 Protein conformational diseases, 95 Protein kinase A (PKA), phosphorylation by, 75 Proteinopathies, 89–90 Proteins, cohesion, 132 Proteoglycan aggrecan, 4 Proteoglycan ECM, 12 Proteolysis, susceptibility to, 17 Ptc, 34 Pthlh, 4 PTHrP, 4–6 expression, 27, 29 Pulmonary vascular remodeling, 231 PVHL, 237 PZ, 7, 18 R RANK ligand (RANKL), 115, 120 Rapamycin, 94 ester, CCI-779, 94 protective eVect of, 95 therapy, 96 Rapamycin (mTOR) pathway, 92 Rapamycin-sensitive kinase. See mTOR enzyme

270 RCC, 236 Reactive oxygen species (ROS), 218 Receptors diVerent combinations of, 16 of TGF- superfamily members, 14 Remodelers, 132 Renal cell carcinoma (RCC), 224 Replication factor C (RFC), 154 Replication protein A (RPA), 151 Respiratory distress syndrome, 230 Restin, 53 Resting zone (RZ), 4 Retinal hemangiomas, 224 Retinoic acid (RA), 187 Retinopathy of prematurity (ROP), 232–233 RGMa, 10 RNA-binding protein, 194 RNA polymerase, 154 R-Smad, 8 activity, 12 phosphorylation, 13 R-Smads and Smad4 (Smad6), inhibiting interactions, 13 Rudimentary condensations, 20 Runt-related, 104 Runx2, 8 RZ, 5, 11 S Saccharomyces cerevisiae, 50, 136–137, 139, 141, 144, 147, 149 cdc7, mutants of, 151 Dbf4, 150 ORC, 142 SAMP6 mice, treatment of, 117 SCA3, 89 SCF E3 ubiquitin ligase complex, 200 Schizosaccharomyces pombe, 137, 141, 149, 160 Orc4, 139 Sclerosteosis, 103, 111 Sclerostin, 103, 119 Secreted frizzled-related proteins (Sfrps), 106 Serine-rich region, in Bik1p protein, 55, 56 Serine-threonine kinase receptors type I, 8 type II, 8 Sex determination pathway, 210 Sfrp1 deletion, 114

Index Shc1p, 78 Shmoos. See Mating cells, Bik1p protein in Short hairpin RNAs (shRNAs), 238–239 Signaling pathways interactions, 8 regulating chondrogenesis chondrocytes, 5–8 Signal transducer and activator of transcription (STAT), 6 Single-span transmembrane proteins type I, 107 Sir1, 160 SiRNA, 17, 149 Sir1 protein, 136 Skeletal abnormality, 109, 111 Skeletal system, embryogenesis, 104 Ski, 9 Smad 1, 9–10 detection, 15 Smad1/5, 15 Smad2, 9, 32 levels, 35 negative form of, 18 Smad2/3, Sox9 associated with, 17 Smad3, 9, 32 negative form of, 18 Smad4, 26 mutants, cartilage-specific, 26 Smad5, 9–10 detection, 15 phosphorylation, BMP7-induced, 31 Smad4;Col2a1-Cre phenotype, 26 Smad4-deficient cells, 26 Smad-mediated pathways, 25 Smad-mediated signaling, 26 Smad (non)-mediated pathways, 25 Smad1 phosphorylation, 11–12 Smads 8, 9 detection, 15 Smads receptor, DNA, 8 Smads receptors, type I, 8 Smad ubiquitination regulatory factors (Smurfs) 1 (Smurf1), 13 2 (Smurf2), 32 Smurfs. See Smad ubiquitination regulatory factors SNF2h, 142 Sno, 9 Sorafenib, 239 Sox6, 20

Index Sox9, 3, 7 expression, 20 Smad2/3 associated with, 17 Sox genes, 20 Spc110p, 78 Spindle pole body (SPB), 61, 67 Spindle positioning, Bik1 protein role in, 63–64 dynein pathway in, 63–66 Kar9p pathway in, 63–64, 66–69 in mating cells, 69–70 Spinobulbar muscular dystrophy (SBMA), 90 Spinocerebellar ataxias (SCAs), 89–90 Split ATPase, 133 STAT, 8 Sternal chondrocytes, 31 Superhelical torsion, 132 Swi6, 160 SWI/SNF proteins, 132 SWI/SNF remodeling factor, 142 Symphalangism, 22–23 Syndecan-3, 11 Synostosis syndrome, multiple, 23 Synthetic lethal interactions, in Kar9p pathway, 77–78 factors and chaperones involved in alpha- and beta-tubulin folding, 77 MEN and/or spindle assembly checkpoint genes, 77 microtubule-associated genes, 77 RNA polymerase, RNA splicing, and ribosomal functions, 77 transcription factors, 78 Synthetic lethality, 63 -Synuclein, 90, 96 mutants, 94 T Tandem aYnity purification (TAP) tag, 78 TAP tag. See Tandem aYnity purification tag Tauopathies, 89–90 T- R-I, 9 T cell factor (Tcf), 105 TEM1 gene, 64 Teratocarcinomas, 239 TGF- , 5, 7, 9, 13, 17–18, 25 responsiveness to, 26 Tgf 1, 17, 34 TGF- s 1–3, detection, 14 Tgf 2, 18 Tgf 2, 17

271 Tgf 3, 17 TGF- –activated kinase, (TAK1), 27 TGF- /activin signaling, 14 TGF- and BMP pathways, balance between, 31 TGF- chondrogenesis, 16 TGF- ligands, 10 TGF- pathways, 10, 16–18, 20, 32 Tgf r2, alleles of, 17 TGF- receptors interactions of, 32 Type I, 17 type II (TGF- RII), 9 TGF- RII, 18 negative form of, 18 receptor, type II, 17 TGF- s 1–3, coexpression, 14 TGF- signaling, 34–35 pathways, 12, 17, 31 regulation of, 10 TGF- specific Smads, 18 TGF- superfamily members, 8 expression of, 14 inhibitors of, 14 Tgf 1upregulation, 35 TGIF, 9 Tie2-Cre, 231 Tie2-expressing monocyte (TEM), 221 Tip1p, from Schizosaccharomyces pombe, 50, 53, 62 Tolloid, 11 TopBP1/Dpb11 complexes, 153 TOPGAL transgene, 115 Trabecular bone, 114 Transactivation domains (TADs), 221 Transcriptional activation complex, 194 Transcriptional activators, recruitment of, 9 Transcriptional activity, 31, 108 Transcriptional coactivator for Smads (Runx2), 26 Transcriptional dysregulation, 91 Transcriptional regulators, 218 Transcriptional repressors, 9 Transcriptional upregulation, 220 Transcription complex, 224 Transcription factor 2 (Runx2), 104 Transcription factors, 141, 221 Abf1, 141–142 Barx2, 21 Mcm1, 141 Osterix, 4

272 Transcription regulators, 217 Transforming growth factor- (TGF- ), 2, 220 superfamily, 8 Transgenic mice, 22, 113 Tub1p protein, 59 interaction with Bik1p, 61 Tubulin cofactors, 51 folding, 77 Tumor-associated macrophages (TAMs), 235 Tumor growth, 219 Tumorigenesis, 223, 226, 234–235, 238 Tumor necrosis factor-, 223 Tumorous germ line, 195 Tumorous mutant, 207 Tumorous phenotype, 209 Tumors, growth of, 218 Twisted gastrulation (Tsg), 11 Type II collagen, 3–4, 27, 34 induction of, 28 Type X collagen, 26 Tyrosine hydroxylase expression, 230 Tyrosine kinase receptor Ror2, 13 U Ubiquitin conjugation of, 93 ligases, 13 moieties, 93 Ubiquitin-conjugating enzyme, 93 Ubiquitin-proteasome pathway, 91, 93 V Valproic acid (VPA), 96 Van Buchem disease, 111 Van Buchem syndrome, 103 Vascular defects, 229–230 Vascular endothelial growth factor (VEGF), 219 Vasculogenesis, 218–220, 229 VEGF, 219, 231, 235–236 expression, 233 VEGF-R2, 220 VEGF-R2þ angioblasts, 220

Index VEGF signaling, 221 VEGF-trap, 239 VHL, 236–237 overexpression of, 240 Von Hippel-Lindau disease, 236–237 Von Hippel-Lindau protein (pVHL), 224, 229 W Wg-dependent mutants, 107 Wnt, 2, 5 Wnt10a, overexpression of, 117 Wnt antagonists, 106, 113–114 Dkk1, 111, 114 WIF-1, 114 Wnt/ -catenin signaling, 107 Wnt canonical pathway, 117 Wnt canonical signaling, 106 Wnt cascade, 112 Wnt coreceptor, 103 Wnt gene family, 109 Wnt morphogens, 118 Wnt pathway, 7, 105 Wnt signaling, 4, 107, 111–112, 114–115, 117, 119 in bone, 120 cascade, 115 pathway, 105 Wound healing, 217 X Xenobiotic detoxification, 221 Xenopus, 139, 142–143, 147–148, 160 embryos, 107 extracts, 155 oocytes, 138 Y YC-1, 240 Yeast, 137 YWTD- propeller domains, 107–108 Z Zinc knuckle motif of CLIP-170, 58

Contents of Previous Volumes Volume 47 1 Early Events of Somitogenesis in Higher Vertebrates: Allocation of Precursor Cells during Gastrulation and the Organization of a Moristic Pattern in the Paraxial Mesoderm Patrick P. L. Tam, Devorah Goldman, Anne Camus, and Gary C. Shoenwolf

2 Retrospective Tracing of the Developmental Lineage of the Mouse Myotome Sophie Eloy-Trinquet, Luc Mathis, and Jean-François Nicolas

3 Segmentation of the Paraxial Mesoderm and Vertebrate Somitogenesis Olivier Pourqule´

4 Segmentation: A View from the Border Claudio D. Stern and Daniel Vasiliauskas

5 Genetic Regulation of Somite Formation Alan Rawls, Jeanne Wilson-Rawls, and Eric N. Olsen

6 Hox Genes and the Global Patterning of the Somitic Mesoderm Ann Campbell Burke

7 The Origin and Morphogenesis of Amphibian Somites Ray Keller

8 Somitogenesis in Zebrafish Scott A. Halley and Christiana Nu¨sslain-Volhard

9 Rostrocaudal Differences within the Somites Confer Segmental Pattern to Trunk Neural Crest Migration Marianne Bronner-Fraser

273

274

Contents of Previous Volumes

Volume 48 1 Evolution and Development of Distinct Cell Lineages Derived from Somites Beate Brand-Saberi and Bodo Christ

2 Duality of Molecular Signaling Involved in Vertebral Chondrogenesis Anne-He´le`ne Monsoro-Burq and Nicole Le Douarin

3 Sclerotome Induction and Differentiation Jennifer L. Docker

4 Genetics of Muscle Determination and Development Hans-Henning Arnold and Thomas Braun

5 Multiple Tissue Interactions and Signal Transduction Pathways Control Somite Myogenesis Anne-Gae¨lle Borycki and Charles P. Emerson, Jr.

6 The Birth of Muscle Progenitor Cells in the Mouse: Spatiotemporal Considerations Shahragim Tajbakhsh and Margaret Buckingham

7 Mouse–Chick Chimera: An Experimental System for Study of Somite Development Josiane Fontaine-Pe´rus

8 Transcriptional Regulation during Somitogenesis Dennis Summerbell and Peter W. J. Rigby

9 Determination and Morphogenesis in Myogenic Progenitor Cells: An Experimental Embryological Approach Charles P. Ordahl, Brian A. Williams, and Wilfred Denetclaw

Volume 49 1 The Centrosome and Parthenogenesis Thomas Ku¨ntziger and Michel Bornens

2 g-Tubulin Berl R. Oakley


275

3 g-Tubulin Complexes and Their Role in Microtubule Nucleation Ruwanthi N. Gunawardane, Sofia B. Lizarraga, Christiane Wiese, Andrew Wilde, and Yixian Zheng

4 g-Tubulin of Budding Yeast Jackie Vogel and Michael Snyder

5 The Spindle Pole Body of Saccharomyces cerevisiae: Architecture and Assembly of the Core Components Susan E. Francis and Trisha N. Davis

6 The Microtubule Organizing Centers of Schizosaccharomyces pombe Iain M. Hagan and Janni Petersen

7 Comparative Structural, Molecular, and Functional Aspects of the Dictyostelium discoideum Centrosome Ralph Gra¨f, Nicole Brusis, Christine Daunderer, Ursula Euteneuer, Andrea Hestermann, Manfred Schliwa, and Masahiro Ueda

8 Are There Nucleic Acids in the Centrosome? Wallace F. Marshall and Joel L. Rosenbaum

9 Basal Bodies and Centrioles: Their Function and Structure Andrea M. Preble, Thomas M. Giddings, Jr., and Susan K. Dutcher

10 Centriole Duplication and Maturation in Animal Cells B. M. H. Lange, A. J. Faragher, P. March, and K. Gull

11 Centrosome Replication in Somatic Cells: The Significance of the G1 Phase Ron Balczon

12 The Coordination of Centrosome Reproduction with Nuclear Events during the Cell Cycle Greenfield Sluder and Edward H. Hinchcliffe

13 Regulating Centrosomes by Protein Phosphorylation Andrew M. Fry, Thibault Mayor, and Erich A. Nigg

14 The Role of the Centrosome in the Development of Malignant Tumors Wilma L. Lingle and Jeffrey L. Salisbury

15 The Centrosome-Associated Aurora/IpI-like Kinase Family T. M. Goepfert and B. R. Brinkley

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16 Centrosome Reduction during Mammalian Spermiogenesis G. Manandhar, C. Simerly, and G. Schatten

17 The Centrosome of the Early C. elegans Embryo: Inheritance, Assembly, Replication, and Developmental Roles Kevin F. O’Connell

18 The Centrosome in Drosophila Oocyte Development Timothy L. Megraw and Thomas C. Kaufman

19 The Centrosome in Early Drosophila Embryogenesis W. F. Rothwell and W. Sullivan

20 Centrosome Maturation Robert E. Palazzo, Jacalyn M. Vogel, Bradley J. Schnackenberg, Dawn R. Hull, and Xingyong Wu

Volume 50 1 Patterning the Early Sea Urchin Embryo Charles A. Ettensohn and Hyla C. Sweet

2 Turning Mesoderm into Blood: The Formation of Hematopoietic Stem Cells during Embryogenesis Alan J. Davidson and Leonard I. Zon

3 Mechanisms of Plant Embryo Development Shunong Bai, Lingjing Chen, Mary Alice Yund, and Zinmay Rence Sung

4 Sperm-Mediated Gene Transfer Anthony W. S. Chan, C. Marc Luetjens, and Gerald P. Schatten

5 Gonocyte–Sertoli Cell Interactions during Development of the Neonatal Rodent Testis Joanne M. Orth, William F. Jester, Ling-Hong Li, and Andrew L. Laslett

6 Attributes and Dynamics of the Endoplasmic Reticulum in Mammalian Eggs Douglas Kline

7 Germ Plasm and Molecular Determinants of Germ Cell Fate Douglas W. Houston and Mary Lou King


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Volume 51 1 Patterning and Lineage Specification in the Amphibian Embryo Agnes P. Chan and Laurence D. Etkin

2 Transcriptional Programs Regulating Vascular Smooth Muscle Cell Development and Differentiation Michael S. Parmacek

3 Myofibroblasts: Molecular Crossdressers Gennyne A. Walker, Ivan A. Guerrero, and Leslie A. Leinwand

4 Checkpoint and DNA-Repair Proteins Are Associated with the Cores of Mammalian Meiotic Chromosomes Madalena Tarsounas and Peter B. Moens

5 Cytoskeletal and Ca2+ Regulation of Hyphal Tip Growth and Initiation Sara Torralba and I. Brent Heath

6 Pattern Formation during C. elegans Vulval Induction Minqin Wang and Paul W. Sternberg

7 A Molecular Clock Involved in Somite Segmentation Miguel Maroto and Olivier Pourquie´

Volume 52 1 Mechanism and Control of Meiotic Recombination Initiation Scott Keeney

2 Osmoregulation and Cell Volume Regulation in the Preimplantation Embryo Jay M. Baltz

3 Cell–Cell Interactions in Vascular Development Diane C. Darland and Patricia A. D’Amore

4 Genetic Regulation of Preimplantation Embryo Survival Carol M. Warner and Carol A. Brenner

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Volume 53 1 Developmental Roles and Clinical Significance of Hedgehog Signaling Andrew P. McMahon, Philip W. Ingham, and Clifford J. Tabin

2 Genomic Imprinting: Could the Chromatin Structure Be the Driving Force? Andras Paldi

3 Ontogeny of Hematopoiesis: Examining the Emergence of Hematopoietic Cells in the Vertebrate Embryo Jenna L. Galloway and Leonard I. Zon

4 Patterning the Sea Urchin Embryo: Gene Regulatory Networks, Signaling Pathways, and Cellular Interactions Lynne M. Angerer and Robert C. Angerer

Volume 54 1 Membrane Type-Matrix Metalloproteinases (MT-MMP) Stanley Zucker, Duanqing Pei, Jian Cao, and Carlos Lopez-Otin

2 Surface Association of Secreted Matrix Metalloproteinases Rafael Fridman

3 Biochemical Properties and Functions of Membrane-Anchored Metalloprotease-Disintegrin Proteins (ADAMs) J. David Becherer and Carl P. Blobel

4 Shedding of Plasma Membrane Proteins Joaquıń Arribas and Anna Merlos-Sua´rez

5 Expression of Meprins in Health and Disease Lourdes P. Norman, Gail L. Matters, Jacqueline M. Crisman, and Judith S. Bond

6 Type II Transmembrane Serine Proteases Qingyu Wu

7 DPPIV, Seprase, and Related Serine Peptidases in Multiple Cellular Functions Wen-Tien Chen, Thomas Kelly, and Giulio Ghersi


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8 The Secretases of Alzheimer’s Disease Michael S. Wolfe

9 Plasminogen Activation at the Cell Surface Vincent Ellis

10 Cell-Surface Cathepsin B: Understanding Its Functional Significance Dora Cavallo-Medved and Bonnie F. Sloane

11 Protease-Activated Receptors Wadie F. Bahou

12 Emmprin (CD147), a Cell Surface Regulator of Matrix Metalloproteinase Production and Function Bryan P. Toole

13 The Evolving Roles of Cell Surface Proteases in Health and Disease: Implications for Developmental, Adaptive, Inflammatory, and Neoplastic Processes Joseph A. Madri

14 Shed Membrane Vesicles and Clustering of Membrane-Bound Proteolytic Enzymes M. Letizia Vittorelli

Volume 55 1 The Dynamics of Chromosome Replication in Yeast Isabelle A. Lucas and M. K. Raghuraman

2 Micromechanical Studies of Mitotic Chromosomes M. G. Poirier and John F. Marko

3 Patterning of the Zebrafish Embryo by Nodal Signals Jennifer O. Liang and Amy L. Rubinstein

4 Folding Chromosomes in Bacteria: Examining the Role of Csp Proteins and Other Small Nucleic Acid-Binding Proteins Nancy Trun and Danielle Johnston

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Volume 56 1 Selfishness in Moderation: Evolutionary Success of the Yeast Plasmid Soundarapandian Velmurugan, Shwetal Mehta, and Makkuni Jayaram

2 Nongenomic Actions of Androgen in Sertoli Cells William H. Walker

3 Regulation of Chromatin Structure and Gene Activity by Poly(ADP-Ribose) Polymerases Alexei Tulin, Yurli Chinenov, and Allan Spradling

4 Centrosomes and Kinetochores, Who needs ‘Em? The Role of Noncentromeric Chromatin in Spindle Assembly Priya Prakash Budde and Rebecca Heald

5 Modeling Cardiogenesis: The Challenges and Promises of 3D Reconstruction Jeffrey O. Penetcost, Claudio Silva, Maurice Pesticelli, Jr., and Kent L. Thornburg

6 Plasmid and Chromosome Traffic Control: How ParA and ParB Drive Partition Jennifer A. Surtees and Barbara E. Funnell

Volume 57 1 Molecular Conservation and Novelties in Vertebrate Ear Development B. Fritzsch and K. W. Beisel

2 Use of Mouse Genetics for Studying Inner Ear Development Elizabeth Quint and Karen P. Steel

3 Formation of the Outer and Middle Ear, Molecular Mechanisms Moise´s Mallo

4 Molecular Basis of Inner Ear Induction Stephen T. Brown, Kareen Martin, and Andrew K. Groves

5 Molecular Basis of Otic Commitment and Morphogenesis: A Role for Homeodomain-Containing Transcription Factors and Signaling Molecules Eva Bober, Silke Rinkwitz, and Heike Herbrand


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6 Growth Factors and Early Development of Otic Neurons: Interactions between Intrinsic and Extrinsic Signals Berta Alsina, Fernando Giraldez, and Isabel Varela-Nieto

7 Neurotrophic Factors during Inner Ear Development Ulla Pirvola and Jukka Ylikoski

8 FGF Signaling in Ear Development and Innervation Tracy J. Wright and Suzanne L. Mansour

9 The Roles of Retinoic Acid during Inner Ear Development Raymond Romand

10 Hair Cell Development in Higher Vertebrates Wei-Qiang Gao

11 Cell Adhesion Molecules during Inner Ear and Hair Cell Development, Including Notch and Its Ligands Matthew W. Kelley

12 Genes Controlling the Development of the Zebrafish Inner Ear and Hair Cells Bruce B. Riley

13 Functional Development of Hair Cells Ruth Anne Eatock and Karen M. Hurley

14 The Cell Cycle and the Development and Regeneration of Hair Cells Allen F. Ryan

Volume 58 1 A Role for Endogenous Electric Fields in Wound Healing Richard Nuccitelli

2 The Role of Mitotic Checkpoint in Maintaining Genomic Stability Song-Tao Liu, Jan M. van Deursen, and Tim J. Yen

3 The Regulation of Oocyte Maturation Ekaterina Voronina and Gary M. Wessel

4 Stem Cells: A Promising Source of Pancreatic Islets for Transplantation in Type 1 Diabetes Cale N. Street, Ray V. Rajotte, and Gregory S. Korbutt

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5 Differentiation Potential of Adipose Derived Adult Stem (ADAS) Cells Jeffrey M. Gimble and Farshid Guilak

Volume 59 1 The Balbiani Body and Germ Cell Determinants: 150 Years Later Malgorzata Kloc, Szczepan Bilinski, and Laurence D. Etkin

2 Fetal–Maternal Interactions: Prenatal Psychobiological Precursors to Adaptive Infant Development Matthew F. S. X. Novak

3 Paradoxical Role of Methyl-CpG-Binding Protein 2 in Rett Syndrome Janine M. LaSalle

4 Genetic Approaches to Analyzing Mitochondrial Outer Membrane Permeability Brett H. Graham and William J. Craigen

5 Mitochondrial Dynamics in Mammals Hsiuchen Chen and David C. Chan

6 Histone Modification in Corepressor Functions Judith K. Davie and Sharon Y. R. Dent

7 Death by Abl: A Matter of Location Jiangyu Zhu and Jean Y. J. Wang

Volume 60 1 Therapeutic Cloning and Tissue Engineering Chester J. Koh and Anthony Atala

2 a-Synuclein: Normal Function and Role in Neurodegenerative Diseases Erin H. Norris, Benoit I. Giasson, and Virginia M.-Y. Lee

3 Structure and Function of Eukaryotic DNA Methyltransferases Taiping Chen and En Li

4 Mechanical Signals as Regulators of Stem Cell Fate Bradley T. Estes, Jeffrey M. Gimble, and Farshid Guilak


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5 Origins of Mammalian Hematopoiesis: In Vivo Paradigms and In Vitro Models M. William Lensch and George Q. Daley

6 Regulation of Gene Activity and Repression: A Consideration of Unifying Themes Anne C. Ferguson-Smith, Shau-Ping Lin, and Neil Youngson

7 Molecular Basis for the Chloride Channel Activity of Cystic Fibrosis Transmembrane Conductance Regulator and the Consequences of Disease-Causing Mutations Jackie F. Kidd, Ilana Kogan, and Christine E. Bear

Volume 61 1 Hepatic Oval Cells: Helping Redefine a Paradigm in Stem Cell Biology P. N. Newsome, M. A. Hussain, and N. D. Theise

2 Meiotic DNA Replication Randy Strich

3 Pollen Tube Guidance: The Role of Adhesion and Chemotropic Molecules Sunran Kim, Juan Dong, and Elizabeth M. Lord

4 The Biology and Diagnostic Applications of Fetal DNA and RNA in Maternal Plasma Rossa W. K. Chiu and Y. M. Dennis Lo

5 Advances in Tissue Engineering Shulamit Levenberg and Robert Langer

6 Directions in Cell Migration Along the Rostral Migratory Stream: The Pathway for Migration in the Brain Shin-ichi Murase and Alan F. Horwitz

7 Retinoids in Lung Development and Regeneration Malcolm Maden

8 Structural Organization and Functions of the Nucleus in Development, Aging, and Disease Leslie Mounkes and Colin L. Stewart

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Volume 62 1 Blood Vessel Signals During Development and Beyond Ondine Cleaver

2 HIFs, Hypoxia, and Vascular Development Kelly L. Covello and M. Celeste Simon

3 Blood Vessel Patterning at the Embryonic Midline Kelly A. Hogan and Victoria L. Bautch

4 Wiring the Vascular Circuitry: From Growth Factors to Guidance Cues Lisa D. Urness and Dean Y. Li

5 Vascular Endothelial Growth Factor and Its Receptors in Embryonic Zebrafish Blood Vessel Development Katsutoshi Goishi and Michael Klagsbrun

6 Vascular Extracellular Matrix and Aortic Development Cassandra M. Kelleher, Sean E. McLean, and Robert P. Mecham

7 Genetics in Zebrafish, Mice, and Humans to Dissect Congenital Heart Disease: Insights in the Role of VEGF Diether Lambrechts and Peter Carmeliet

8 Development of Coronary Vessels Mark W. Majesky

9 Identifying Early Vascular Genes Through Gene Trapping in Mouse Embryonic Stem Cells Frank Kuhnert and Heidi Stuhlmann

Volume 63 1 Early Events in the DNA Damage Response Irene Ward and Junjie Chen

2 Afrotherian Origins and Interrelationships: New Views and Future Prospects Terence J. Robinson and Erik R. Seiffert

3 The Role of Antisense Transcription in the Regulation of X-Inactivation Claire Rougeulle and Philip Avner


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4 The Genetics of Hiding the Corpse: Engulfment and Degradation of Apoptotic Cells in C. elegans and D. melanogaster Zheng Zhou, Paolo M. Mangahas, and Xiaomeng Yu

5 Beginning and Ending an Actin Filament: Control at the Barbed End Sally H. Zigmond

6 Life Extension in the Dwarf Mouse Andrzej Bartke and Holly Brown-Borg

Volume 64 1 Stem/Progenitor Cells in Lung Morphogenesis, Repair, and Regeneration David Warburton, Mary Anne Berberich, and Barbara Driscoll

2 Lessons from a Canine Model of Compensatory Lung Growth Connie C. W. Hsia

3 Airway Glandular Development and Stem Cells Xiaoming Liu, Ryan R. Driskell, and John F. Engelhardt

4 Gene Expression Studies in Lung Development and Lung Stem Cell Biology Thomas J. Mariani and Naftali Kaminski

5 Mechanisms and Regulation of Lung Vascular Development Michelle Haynes Pauling and Thiennu H. Vu

6 The Engineering of Tissues Using Progenitor Cells Nancy L. Parenteau, Lawrence Rosenberg, and Janet Hardin-Young

7 Adult Bone Marrow-Derived Hemangioblasts, Endothelial Cell Progenitors, and EPCs Gina C. Schatteman

8 Synthetic Extracellular Matrices for Tissue Engineering and Regeneration Eduardo A. Silva and David J. Mooney

9 Integrins and Angiogenesis D. G. Stupack and D. A. Cheresh

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Volume 65 1 Tales of Cannibalism, Suicide, and Murder: Programmed Cell Death in C. elegans Jason M. Kinchen and Michael O. Hengartner

2 From Guts to Brains: Using Zebrafish Genetics to Understand the Innards of Organogenesis Carsten Stuckenholz, Paul E. Ulanch, and Nathan Bahary

3 Synaptic Vesicle Docking: A Putative Role for the Munc18/Sec1 Protein Family Robby M. Weimer and Janet E. Richmond

4 ATP-Dependent Chromatin Remodeling Corey L. Smith and Craig L. Peterson

5 Self-Destruct Programs in the Processes of Developing Neurons David Shepherd and V. Hugh Perry

6 Multiple Roles of Vascular Endothelial Growth Factor (VEGF) in Skeletal Development, Growth, and Repair Elazar Zelzer and Bjorn R. Olsen

7 G-Protein Coupled Receptors and Calcium Signaling in Development Geoffrey E. Woodard and Juan A. Rosado

8 Differential Functions of 14-3-3 Isoforms in Vertebrate Development Anthony J. Muslin and Jeffrey M. C. Lau

9 Zebrafish Notochordal Basement Membrane: Signaling and Structure Annabelle Scott and Derek L. Stemple

10 Sonic Hedgehog Signaling and the Developing Tooth Martyn T. Cobourne and Paul T. Sharpe

Volume 66 1 Stepwise Commitment from Embryonic Stem to Hematopoietic and Endothelial Cells Changwon Park, Jesse J. Lugus, and Kyunghee Choi


287

2 Fibroblast Growth Factor Signaling and the Function and Assembly of Basement Membranes Peter Lonai

3 TGF- Superfamily and Mouse Craniofacial Development: Interplay of Morphogenetic Proteins and Receptor Signaling Controls Normal Formation of the Face Marek Dudas and Vesa Kaartinen

4 The Colors of Autumn Leaves as Symptoms of Cellular Recycling and Defenses Against Environmental Stresses Helen J. Ougham, Phillip Morris, and Howard Thomas

5 Extracellular Proteases: Biological and Behavioral Roles in the Mammalian Central Nervous System Yan Zhang, Kostas Pothakos, and Styliana-Anna (Stella) Tsirka

6 The Genetic Architecture of House Fly Mating Behavior Lisa M. Meffert and Kara L. Hagenbuch

7 Phototropins, Other Photoreceptors, and Associated Signaling: The Lead and Supporting Cast in the Control of Plant Movement Responses Bethany B. Stone, C. Alex Esmon, and Emmanuel Liscum

8 Evolving Concepts in Bone Tissue Engineering Catherine M. Cowan, Chia Soo, Kang Ting, and Benjamin Wu

9 Cranial Suture Biology Kelly A Lenton, Randall P. Nacamuli, Derrick C. Wan, Jill A. Helms, and Michael T. Longaker

Volume 67 1 Deer Antlers as a Model of Mammalian Regeneration Joanna Price, Corrine Faucheux, and Steve Allen

2 The Molecular and Genetic Control of Leaf Senescence and Longevity in Arabidopsis Pyung Ok Lim and Hong Gil Nam

3 Cripto-1: An Oncofetal Gene with Many Faces Caterina Bianco, Luigi Strizzi, Nicola Normanno, Nadia Khan, and David S. Salomon

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4 Programmed Cell Death in Plant Embryogenesis Peter V. Bozhkov, Lada H. Filonova, and Maria F. Suarez

5 Physiological Roles of Aquaporins in the Choroid Plexus Daniela Boassa and Andrea J. Yool

6 Control of Food Intake Through Regulation of cAMP Allan Z. Zhao

7 Factors Affecting Male Song Evolution in Drosophila montana Anneli Hoikkala, Kirsten Klappert, and Dominique Mazzi

8 Prostanoids and Phosphodiesterase Inhibitors in Experimental Pulmonary Hypertension Ralph Theo Schermuly, Hossein Ardeschir Ghofrani, and Norbert Weissmann

9 14-3-3 Protein Signaling in Development and Growth Factor Responses Daniel Thomas, Mark Guthridge, Jo Woodcock, and Angel Lopez

10 Skeletal Stem Cells in Regenerative Medicine Wataru Sonoyama, Carolyn Coppe, Stan Gronthos, and Songtao Shi

Volume 68 1 Prolactin and Growth Hormone Signaling Beverly Chilton and Aveline Hewetson

2 Alterations in cAMP-Mediated Signaling and Their Role in the Pathophysiology of Dilated Cardiomyopathy Matthew A. Movsesian and Michael R. Bristow

3 Corpus Luteum Development: Lessons from Genetic Models in Mice Anne Bachelot and Nadine Binart

4 Comparative Developmental Biology of the Mammalian Uterus Thomas E. Spencer, Kanako Hayashi, Jianbo Hu, and Karen D. Carpenter

5 Sarcopenia of Aging and Its Metabolic Impact Helen Karakelides and K. Sreekumaran Nair

6 Chemokine Receptor CXCR3: An Unexpected Enigma Liping Liu, Melissa K. Callahan, DeRen Huang, and Richard M. Ransohoff


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7 Assembly and Signaling of Adhesion Complexes Jorge L. Sepulveda, Vasiliki Gkretsi, and Chuanyue Wu

8 Signaling Mechanisms of Higher Plant Photoreceptors: A Structure-Function Perspective Haiyang Wang

9 Initial Failure in Myoblast Transplantation Therapy Has Led the Way Toward the Isolation of Muscle Stem Cells: Potential for Tissue Regeneration Kenneth Urish, Yasunari Kanda, and Johnny Huard

10 Role of 14-3-3 Proteins in Eukaryotic Signaling and Development Dawn L. Darling, Jessica Yingling, and Anthony Wynshaw-Boris

Volume 69 1 Flipping Coins in the Fly Retina Tamara Mikeladze-Dvali, Claude Desplan, and Daniela Pistillo

2 Unraveling the Molecular Pathways That Regulate Early Telencephalon Development Jean M. He´bert

3 Glia–Neuron Interactions in Nervous System Function and Development Shai Shaham

4 The Novel Roles of Glial Cells Revisited: The Contribution of Radial Glia and Astrocytes to Neurogenesis Tetsuji Mori, Annalisa Buffo, and Magdalena Go¨tz

5 Classical Embryological Studies and Modern Genetic Analysis of Midbrain and Cerebellum Development Mark Zervas, Sandra Blaess, and Alexandra L. Joyner

6 Brain Development and Susceptibility to Damage; Ion Levels and Movements Maria Erecinska, Shobha Cherian, and Ian A. Silver

7 Thinking about Visual Behavior; Learning about Photoreceptor Function Kwang-Min Choe and Thomas R. Clandinin

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8 Critical Period Mechanisms in Developing Visual Cortex Takao K. Hensch

9 Brawn for Brains: The Role of MEF2 Proteins in the Developing Nervous System Aryaman K. Shalizi and Azad Bonni

10 Mechanisms of Axon Guidance in the Developing Nervous System Ce´line Plachez and Linda J. Richards

Volume 70 1 Magnetic Resonance Imaging: Utility as a Molecular Imaging Modality James P. Basilion, Susan Yeon, and Rene´ Botnar

2 Magnetic Resonance Imaging Contrast Agents in the Study of Development Angelique Louie

3 1H/19F Magnetic Resonance Molecular Imaging with Perfluorocarbon Nanoparticles Gregory M. Lanza, Patrick M. Winter, Anne M. Neubauer, Shelton D. Caruthers, Franklin D. Hockett, and Samuel A. Wickline

4 Loss of Cell Ion Homeostasis and Cell Viability in the Brain: What Sodium MRI Can Tell Us Fernando E. Boada, George LaVerde, Charles Jungreis, Edwin Nemoto, Costin Tanase, and Ileana Hancu

5 Quantum Dot Surfaces for Use In Vivo and In Vitro Byron Ballou

6 In Vivo Cell Biology of Cancer Cells Visualized with Fluorescent Proteins Robert M. Hoffman

7 Modulation of Tracer Accumulation in Malignant Tumors: Gene Expression, Gene Transfer, and Phage Display Uwe Haberkorn

8 Amyloid Imaging: From Benchtop to Bedside Chungying Wu, Victor W. Pike, and Yanming Wang

9 In Vivo Imaging of Autoimmune Disease in Model Systems Eric T. Ahrens and Penelope A. Morel


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Volume 71 1 The Choroid Plexus-Cerebrospinal Fluid System: From Development to Aging Zoran B. Redzic, Jane E. Preston, John A. Duncan, Adam Chodobski, and Joanna Szmydynger-Chodobska

2 Zebrafish Genetics and Formation of Embryonic Vasculature Tao P. Zhong

3 Leaf Senescence: Signals, Execution, and Regulation Yongfeng Guo and Susheng Gan

4 Muscle Stem Cells and Regenerative Myogenesis Iain W. McKinnell, Gianni Parise, and Michael A. Rudnicki

5 Gene Regulation in Spermatogenesis James A. MacLean II and Miles F. Wilkinson

6 Modeling Age-Related Diseases in Drosophila: Can this Fly? Kinga Michno, Diana van de Hoef, Hong Wu, and Gabrielle L. Boulianne

7 Cell Death and Organ Development in Plants Hilary J. Rogers

8 The Blood-Testis Barrier: Its Biology, Regulation, and Physiological Role in Spermatogenesis Ching-Hang Wong and C. Yan Cheng

9 Angiogenic Factors in the Pathogenesis of Preeclampsia Hai-Tao Yuan, David Haig, and S. Ananth Karumanchi

Volume 72 1 Defending the Zygote: Search for the Ancestral Animal Block to Polyspermy Julian L. Wong and Gary M. Wessel

2 Dishevelled: A Mobile Scaffold Catalyzing Development Craig C. Malbon and Hsien-yu Wang

3 Sensory Organs: Making and Breaking the Pre-Placodal Region Andrew P. Bailey and Andrea Streit

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4 Regulation of Hepatocyte Cell Cycle Progression and Differentiation by Type I Collagen Structure Linda K. Hansen, Joshua Wilhelm, and John T. Fassett

5 Engineering Stem Cells into Organs: Topobiological Transformations Demonstrated by Beak, Feather, and Other Ectodermal Organ Morphogenesis Cheng-Ming Chuong, Ping Wu, Maksim Plikus, Ting-Xin Jiang, and Randall Bruce Widelitz

6 Fur Seal Adaptations to Lactation: Insights into Mammary Gland Function Julie A. Sharp, Kylie N. Cane, Christophe Lefevre, John P. Y. Arnould, and Kevin R. Nicholas

Volume 73 1 The Molecular Origins of Species-Specific Facial Pattern Samantha A. Brugmann, Minal D. Tapadia, and Jill A. Helms

2 Molecular Bases of the Regulation of Bone Remodeling by the Canonical Wnt Signaling Pathway Donald A. Glass II and Gerard Karsenty

3 Calcium Sensing Receptors and Calcium Oscillations: Calcium as a First Messenger Gerda E. Breitwieser

4 Signal Relay During the Life Cycle of Dictyostelium Dana C. Mahadeo and Carole A. Parent

5 Biological Principles for Ex Vivo Adult Stem Cell Expansion Jean-François Pare´ and James L. Sherley

6 Histone Deacetylation as a Target for Radiosensitization David Cerna, Kevin Camphausen, and Philip J. Tofilon

7 Chaperone-Mediated Autophagy in Aging and Disease Ashish C. Massey, Cong Zhang, and Ana Maria Cuervo

8 Extracellular Matrix Macroassembly Dynamics in Early Vertebrate Embryos Andras Czirok, Evan A. Zamir, Michael B. Filla, Charles D. Little, and Brenda J. Rongish


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Volume 74 1 Membrane Origin for Autophagy Fulvio Reggiori

2 Chromatin Assembly with H3 Histones: Full Throttle Down Multiple Pathways Brian E. Schwartz and Kami Ahmad

3 Protein–Protein Interactions of the Developing Enamel Matrix John D. Bartlett, Bernhard Ganss, Michel Goldberg, Janet Moradian-Oldak, Michael L. Paine, Malcolm L. Snead, Xin Wen, Shane N. White, and Yan L. Zhou

4 Stem and Progenitor Cells in the Formation of the Pulmonary Vasculature Kimberly A. Fisher and Ross S. Summer

5 Mechanisms of Disordered Granulopoiesis in Congenital Neutropenia David S. Grenda and Daniel C. Link

6 Social Dominance and Serotonin Receptor Genes in Crayfish Donald H. Edwards and Nadja Spitzer

7 Transplantation of Undifferentiated, Bone Marrow-Derived Stem Cells Karen Ann Pauwelyn and Catherine M. Verfaillie

8 The Development and Evolution of Division of Labor and Foraging Specialization in a Social Insect (Apis mellifera L.) Robert E. Page Jr., Ricarda Scheiner, Joachim Erber, and Gro V. Amdam

Volume 75 1 Dynamics of Assembly and Reorganization of Extracellular Matrix Proteins Sarah L. Dallas, Qian Chen, and Pitchumani Sivakumar

2 Selective Neuronal Degeneration in Huntington’s Disease Catherine M. Cowan and Lynn A. Raymond

3 RNAi Therapy for Neurodegenerative Diseases Ryan L. Boudreau and Beverly L. Davidson

4 Fibrillins: From Biogenesis of Microfibrils to Signaling Functions Dirk Hubmacher, Kerstin Tiedemann, and Dieter P. Reinhardt

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5 Proteasomes from Structure to Function: Perspectives from Archaea Julie A. Maupin-Furlow, Matthew A. Humbard, P. Aaron Kirkland, Wei Li, Christopher J. Reuter, Amy J. Wright, and G. Zhou

6 The Cytomatrix as a Cooperative System of Macromolecular and Water Networks V. A. Shepherd

7 Intracellular Targeting of Phosphodiesterase-4 Underpins Compartmentalized cAMP Signaling Martin J. Lynch, Elaine V. Hill, and Miles D. Houslay

Developmental Vascular Biology (Current Topics in Developmental Biology)

Developmental Vascular Biology, Volume 62 (Current Topics in Developmental Biology)

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Current Topics in Developmental Biology