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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.

Frank Antonicelli (99), Faculty of Medicine, Extracellular Matrix and Cell Signaling—Reims University, UMR 6198 CNRS, 51095 Reims Cedex, France C. W. Archer (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Georges Bellon (99), Faculty of Medicine, Extracellular Matrix and Cell Signaling—Reims University, UMR 6198 CNRS, 51095 Reims Cedex, France Laurent Debelle (99), Faculty of Science, Extracellular Matrix and Cell Signaling—Reims University, UMR 6198 CNRS, 51095 Reims Cedex, France Nino Devidze (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 Patricia Donahoe (37), Department of Pediatric Surgery, Massachusetts General Hospital, Harvard Medical School, Boston, Massachusetts 02114 G. P. Dowthwaite (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Cristina Florea (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 David R. Harder (75), Department of Physiology, Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, Wisconsin 53226 William Hornebeck (99), Faculty of Medicine, Extracellular Matrix and Cell Signaling—Reims University, UMR 6198 CNRS, 51095 Reims Cedex, France Marshall S. Horwitz (157), Department of Medicine, Division of Medical Genetics, University of Washington School of Medicine, Seattle, Washington 98195 Danica Jakovcevic (75), Department of Physiology, Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, Wisconsin 53226 Hosein Kami Kia (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 ix

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I. M. Khan (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Lee-Ming Kow (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 Vera P. Krymskaya (61), Department of Medicine, Pulmonary, Allergy, and Critical Care Division, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Anna Lee (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 David MacLaughlin (37), Department of Pediatric Surgery, Massachusetts General Hospital, Harvard Medical School, Boston, Massachusetts 02114 S. F. Oldfield (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Reynold A. Panettieri, Jr. (61), Department of Medicine, Pulmonary, Allergy, and Critical Care Division, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Donald Pfaff (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 S. N. Redman (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Stephen J. Salipante (157), Department of Genome Sciences, University of Washington School of Medicine, Seattle, Washington 98195 Marlene Schwanzel-Fukuda (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 R. Williams (1), CardiV School of Biosciences, CardiV University, CardiV CF103US, Wales, United Kingdom Jin Zhou (37), Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021

Contents

Contributors

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1 The Development of Synovial Joints I. M. Khan, S. N. Redman, R. Williams, G. P. Dowthwaite, S. F. Oldfield, and C. W. Archer I. II. III. IV. V. VI.

Introduction 2 Specification and Patterning of the Joint 5 Joint Cavitation and the Role of Mechanical Stimuli 13 Cell–Matrix Interactions During Joint Formation 19 The Development of Articular Cartilage: Overview 22 Aspects of Postnatal Joint Biology 23 References 28

2 Development of a Sexually Differentiated Behavior and Its Underlying CNS Arousal Functions Lee-Ming Kow, Cristina Florea, Marlene Schwanzel-Fukuda, Nino Devidze, Hosein Kami Kia, Anna Lee, Jin Zhou, David MacLaughlin, Patricia Donahoe, and Donald Pfaff I. Introduction 38 II. Sexual DiVerentiation of Brain Mechanisms Producing Lordosis Behavior 41 III. Development of Brain Mechanisms Underlying Arousal 53 IV. Outlook for New Work on Sexually DiVerentiated Behaviors References 56

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3 Phosphodiesterases Regulate Airway Smooth Muscle Function in Health and Disease Vera P. Krymskaya and Reynold A. Panettieri, Jr. I. II. III. IV. V.

Introduction 61 Developmental and Physiological Responses Regulated by PDEs PDEs and Airway Smooth Muscle Function 67 Cytokine and Chemokine Secretion by ASM: A Role for PDEs Conclusions 70 Acknowledgments 70 References 71

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4 Role of Astrocytes in Matching Blood Flow to Neuronal Activity Danica Jakovcevic and David R. Harder I. Introduction 76 II. Neurovascular Unit 76 III. Neuron–Astrocyte Interactions and Regulation of Synaptic Transmission in the Brain 78 IV. Neurovascular Coupling: Role of Astrocytes 82 V. Summary 89 References 90

5 Elastin-Elastases and Inflamm-Aging Frank Antonicelli, Georges Bellon, Laurent Debelle, and William Hornebeck I. Introduction 100 II. Elastic Fibers: Formation and Degradation 103 III. Biological Activities of Elastin Peptides (EPs): The ‘‘Elastin Receptor System’’ 111 IV. Elastolysis, Aging, and AAAs 117 V. The Elastin Connection in Melanoma 127 VI. Concluding Remarks 135 References 138

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6 A Phylogenetic Approach to Mapping Cell Fate Stephen J. Salipante and Marshall S. Horwitz I. II. III. IV. V. VI. VII.

Development 158 Fate Maps 159 Limited Information from Cell Marking 165 Stochastic Nature of Development in Higher Organisms A New Approach: Phylogenetic Fate Mapping 171 Challenges Dealing with Randomness in Development Extended Applications of Phylogenetic Fate Mapping Acknowledgments 180 References 180

Index 185 Contents of Previous Volumes

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168 177 179

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The Development of Synovial Joints I. M. Khan, S. N. Redman, R. Williams, G. P. Dowthwaite, S. F. Oldfield, and C. W. Archer CardiV School of Biosciences, CardiV University CardiV CF103US, Wales, United Kingdom

I. Introduction A. Joint Morphogenesis: Overview II. Specification and Patterning of the Joint A. The Limb Field and Joints B. Formation of the Joint Interzone III. IV. V. VI.

Joint Cavitation and the Role of Mechanical Stimuli Cell–Matrix Interactions During Joint Formation The Development of Articular Cartilage: Overview Aspects of Postnatal Joint Biology References

During vertebrate evolution, successful adaptation of animal limbs to a variety of ecological niches depended largely on the formation and positioning of synovial joints. The function of a joint is to allow smooth articulation between opposing skeletal elements and to transmit biomechanical loads through the structure, and this is achieved through covering the ends of bones with articular cartilage, lubricating the joint with synovial fluid, using ligaments to bind the skeletal elements together, and encapsulating the joint in a protective fibrous layer of tissue. The diversity of limb generation has been proposed to occur through sequential branching and segmentation of precartilaginous skeletal elements along the proximodistal axis of the limb. The position of future joints is first delimited by areas of higher cell density called interzones initially through an as yet unidentified inductive signal, subsequently specification of these regions is controlled hierarchically by wnt14 and gdf5, respectively. Joint‐forming cell fate although specified is not fixed, and joints will fuse if growth factor signaling is perturbed. Cavitation, the separation of the two opposing skeletal elements, and joint morphogenesis, the process whereby the joint cells organize and mature to establish a functional interlocking and reciprocally shaped joint, are slowly being unraveled through studying the plethora of molecules that make up the unique extracellular matrix of the forming structure. The joint lining tissue, articular cartilage, is avascular, and this limits its reparative capacity such that arthritis and associated joint pathologies are the single largest cause of disability in Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)79001-9

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the adult population. Recent discoveries of adult stem cells and more specifically the isolation of chondroprogenitor cells from articular cartilage are extending available therapeutic options, though only with a more complete understanding of synovial joint development can such options have greater chances of success. ß 2007, Elsevier Inc.

I. Introduction A central characteristic of the vertebrate form is the possession of an internal skeleton that first comprises a supporting structure and, together with articulations, provides a means of both movement and locomotion. Joints have been classified according to their structures that, in turn, reflect the degree of movement that any one joint aVords. Earlier investigators described four types of joint. First, a synarthrosis where the skeletal elements are joined to each other by cartilage, fibrocartilage, or other connective tissues. Second, a schizarthrosis where the space between the skeletal elements (referred to as an interzone) comprises connective tissue containing a single or a number of cavities that are usually small. Third, a hemiarthrosis or periarthrosis that has a single joint cavity but the skeletal elements remain united around the periphery. Fourth and finally, a eudiarthrosis (more recently referred to as a diarthrosis) that comprises separate articulating elements and a joint cavity that is limited peripherally by synovial tissues (Haines, 1941, 1942). More recently, this classification has been refined to include the degree of movement that the joint allows. Consequently, the synarthrosis is maintained and the bones are in almost direct contact but interspersed by connective tissue and there is no appreciable movement as, for example, between cranial bones. Second, an amphiarthrosis that has contiguous osseous surfaces connected by flattened disks of fibrocartilage as in the vertebrae and pubic symphysis that allow for limited movement. Third and last is the diarthrosis that is freely movable possesses a cavity that is lined by a synovial membrane. This type of joint may or may not possess ligamentous and meniscal structures internally (Gray, 1988). This chapter will restrict itself to considering synovial joints that are mainly found within the appendicular skeleton. Synovial joints permit a wide range of movement at the interface of two opposing skeletal elements and can withstand loads of up to 10 times the body weight with a very low‐friction coeYcient (RatcliVe and Mow, 1996). As mentioned above, synovial joints are composed of a variety of tissues that all contribute to the function of the joint (Fig. 1). A fibrous articular capsule, that is continuous with the periosteum, encapsulates and stabilizes the joint. Occasionally, the joints are also stabilized by the presence of accessory ligaments within the joint space linking the skeletal elements such as the cruciate ligament of the knee joint. A synovial membrane lines the capsule, which is

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Joint capsule

Articular cartilage

Synovium

Epiphyses

Figure 1 Generalized diagram of a synovial joint. Articular cartilage covers the skeletal elements. The bones are separated by the synovial joint cavity. A fibrous capsule that encapsulates the joint connects the skeletal elements. This capsule is lined by a synovial membrane that secretes synovial fluid which enables smooth articulation of the joint.

responsible for secreting the synovial fluid. The synovial fluid provides the lubrication, required for low‐friction articulation, and nutrition for the avascular articular cartilage. The surfaces of the two opposing skeletal elements are covered by hyaline articular cartilage or occasionally fibrocartilage (reviewed by Benjamin, 1999). It is the hyaline articular cartilage that contributes most toward the functional ability of the joint providing a durable load‐bearing surface that permits pain‐free movement. Because of the variety of tissues comprising the synovial joint, it may also be considered an organ. Consequently, disease or trauma aVecting one tissue of the joint has wide‐ranging implications on the health and function of the joint as a whole. Osteoarthritis and inflammatory rheumatoid arthritis are debilitating conditions that aVect joint function and while our understanding

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of the pathology of these diseases has greatly improved, the only truly successful treatment restoring joint function is total joint replacement. Trauma to the articular cartilage fails to heal spontaneously due, in part, to its avascular nature and will often progress to osteoarthritis with time. While most current articular cartilage repair strategies provide symptomatic relief and improved joint function, most repair tissues degenerate with time due to the nature of the tissue formed and the lack of integration between the host and repair tissue (Redman et al., 2005). A greater understanding of the mechanisms involved in the development of synovial joints and the development of articular cartilage may enable the elucidation of novel repair strategies to restore long‐term joint function following trauma and disease.

A. Joint Morphogenesis: Overview Long bone formation initiates as uninterrupted mesenchyme condensations in the early limb buds, which undergo cytodiVerentiation to chondrocytes (Fell, 1925; Thorogood and HinchliVe, 1975). The proximal region of the mesenchymal condensations gives rise to the humerus or femur with the more distal regions forming the radius/ulna or tibia/fibula and digits. Initially, these contiguous condensations chondrify. Subsequently, at the sites of joint formation, the resident cells flatten and become nonchondrogenic to form what is known as an interzone (Archer et al., 2003; Craig et al., 1987) (Fig. 2). In this respect, in long bone elements at least, the formation of the joint is a secondary segmentation event. The mechanisms underlying joint site determination within the early embryonic cartilage, however, remain poorly understood and will be considered below. Depending on species, the structure can vary between a distinct three‐layered structure comprising two chondrogenous layers separated by a flattened cell central layer as seen in chick long bone elements (Craig et al., 1987) to a thin flattened cell layer (two to three cells) as seen in humans (Edwards et al., 1994). The tissues of the joint are believed to derive from the cells of the interzone (Mitrovic, 1977). In the chick, the two outer layers diVerentiate into chondrocytes and are incorporated into the epiphyses of the lengthening anlagen. It has been suggested that cells from the intermediate cell layer in fact give rise to the articular cartilage that covers the ends of the opposing bones (Archer et al., 1994; Ito and Kida, 2000). The origin of the cells within the interzone that gives rise to the joint ligaments and synovial lining remains to be fully elucidated (Pacifici et al., 2005). In contrast, if we consider smaller elements such as carpal or tarsal elements, then these initiate as individual condensations that on chondrification and expansion abut each other leaving a layer of flattened cells. Whether these cells constitute an interzone is still open to question and will be discussed later (Storm and Kingsley, 1999). While the

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1. The Development of Synovial Joints Mesenchymal condensation

Joint specification

Chondrogenic differentiation

BMPs Chordin Noggin Initiation

Wnt14

Cavitation

Joint formation

Outer layer Interzone formation

Interzone

Intermediate layer

Articular cartilage Synovial cavity

Gdf5

Figure 2 Interzone specification, formation, and cavitation during synovial joint morphogenesis. Within an initial mesenchymal condensation, an unknown trigger stimulates wnt14 expression at the site of incipient joint formation. Gdf5 is, thereafter, expressed and the cells take on an elongated morphology and significantly reduce sox9 and collagen type II expression. BMP antagonists chordin and noggin are expressed in the interzone cells and act to stabilize joint‐ inducing positional cues. The interzone adopts a three‐layered structure (in the case of long bone elements) that undergoes separation or cavitation on mechanically induced synthesis of hyaluronan. The morphogenesis of the functional joint organ results in articular cartilage lining the ends of skeletal elements, which are bathed in synovial fluid, produced by a synovial membrane, and encased within a fibrous capsule.

cell lineage of the joints structure remains rather obscure, we know more about the initial specification of the joints and some of the signaling molecules involved. The next phase of development is the formation of the joint cavity, thus providing functionality to the structure concomitant with the initiation of muscular activity (Fig. 2). The process of cavitation is rather counterintuitive since it involves the generation of a cavity between two cartilaginous elements that are growing against each other through forces largely generated by hypertrophy and matrix secretion within the element. It is achieved by cells of the interzone, articular surface, and synovium through massive upregulation of hyaluronan (HA) synthesis that is movement (mechanically) dependent since paralysis of the embryo results in failure to cavitate or secondary cartilaginous fusion in previously cavitated joints.

II. Specification and Patterning of the Joint A. The Limb Field and Joints Direct evidence for patterning of joints within mesenchymal condensations in developing limbs is unavailable; in its stead is a plethora of data linking the growth of the developing limb and molecules that govern growth and

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patterning, to specification of synovial joints. Limbs develop from small protrusions arising from the somatopleural mesoderm from the main trunk of the embryo. T‐box transcription factors Tbx5 and Tbx4 specify the developing forelimb and hind limb, respectively (Agarwal et al., 2003; Naiche and Papaioannou, 2003; Takeuchi et al., 2003) and act in concert with downstream wnt signaling to activate fibroblast growth factor 10 (fgf10) expression in the developing limb bud. Positioning and patterning of the limb involves cellular interactions between the ectoderm surrounding the limb and mesenchymal cells that form the core of the limb bud. Cells within the limb bud acquire positional identities along each of the three axes of the limb, proximal–distal, anterior–posterior, and dorsal–ventral and this determines how cells diVerentiate. The apical ectodermal ridge (AER) is a specialized epithelial structure induced at the junction between dorsal and ventral ectoderm at the distal end of the limb, slightly after limb bud formation, and expresses fgf8 and fgf4 to promote growth along the proximal– distal axis. Underlying the AER is a region (
250 mm) of undiVerentiated, proliferating mesenchyme called the progress zone where cells acquire their positional identity such that cells leaving early will form proximal structures and those leaving later form distal ones (Summerbell et al., 1973). The signaling center for anterior–posterior patterning resides in the zone of polarizing activity (ZPA) in the mesenchyme at the posterior distal margin of the limb bud. Grafting the ZPA to the anterior distal margin generates mirror image limb duplication. Secreted morphogen sonic hedgehog (shh) is produced in the ZPA and it can duplicate the eVects of ZPA transplantation. The molecular mechanism of shh involves regulating the positive and negative transcriptional activity of Gli3, high levels of shh promote transcriptional activation and low levels produce transcriptional repression. The dorsal– ventral polarity of the limb bud is controlled by the surface ectoderm by wnt7a that induces the expression of Lmx1, a LIM homeodomain transcription factor, in the underlying dorsal mesenchyme. Ectopic expression of Lmx1 in the ventral mesenchyme induces double dorsal limbs (Riddle et al., 1995; Vogel et al., 1995) and further, mice with null mutations of wnt7a display dorsal to ventral transformation of cell fate, that is, footpads appear on the dorsal part of the foot, indicating wnt7a is a dorsalizing signal (Parr and McMahon, 1995). Coordinated and integrated activities of these three signaling centers drive patterned limb growth (Niswander, 2002). Continual fgf8 expression at the AER promoting growth along the proximal–distal axis would be expected to result in extended digits, and this is exactly what occurs when beads soaked in shh, that maintain fgf8 expression in the AER in nearby digits, are placed in interdigital spaces, penultimate phalanges were duplicated (Sanz‐Ezquerro and Tickle, 2003). But, why are only penultimate phalanges duplicated? It appears that terminal phalanges are formed by a diVerent mechanism than proximal ones, switching oV fgf8 activates an

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alternative pathway specifying terminal phalanges that appears to be heavily dependent on wnt signaling. In their experiments, Sanz‐Ezquerro and Tickle found that formation of a new joint may involve a ‘‘threshold mechanism’’ related to the length of mesenchymal condensation below which no segmentation occurs; also, the increase in length of each respective phalange prior to segmentation occurring is intrinsically diVerent for each digit. Fgf8 is initially regulated by shh in the ZPA and is thought to be later regulated by Indian hedgehog (ihh) in the condensing mesenchyme of the digits. In their analysis of the short digits radiation‐induced mouse mutant (Dsh), Niedermaier et al. (2005) found that a chromosomal inversion on chromosome 5 caused shh expression to replace ihh expression in the developing phalanges. In the heterozygote Dsh/þ proximal and medial phalanges are fused, shh replaces ihh expression in the cartilage analagen resulting in disruption of an ihh– pthrp feedback loop (Niedermaier et al., 2005) that delays chondrocyte diVerentiation and hypertrophy causing shortening of digits. Gdf5 expression, a key marker of incipient joint formation, is absent in Dsh/þ mutant digits and conversely ihh is not expressed in the digits of brachypodism (gdf5 null) mice. Additionally, mutations in ihh cause human brachydactyly type A1 and ihh mutant mice have grossly shortened limbs and partially fused joints between the humerus and ulna, and between the carpals, as well as unsegmented and uncalcified mutant digits (St‐Jacques et al., 1999). The implication is that disruption of the physical distance between ihh and pthrp expression domains interferes with joint formation and function. Although the most recent emphasis on joint formation has focused on secreted signaling molecules, earlier work suggested that Hox genes, homeodomain‐containing transcription factors that pattern the body axis of animal embryos, could also be responsible for specification of segmentation boundaries in developing limbs. Hox genes are organized in clusters and are expressed along the body axis in a manner corresponding to their position within the cluster (Krumlauf, 1994). In higher vertebrates, there are 39 Hox genes found on 4 clusters (a, b, c, and d) on diVerent chromosomes (Koussoulakos, 2004). Each Hox gene has a specific anterior limit of expression with no posterior limit within a morphogenetic field. All four Hox groups play a role in limb development; however, the gene expression patterns of Hoxa9–13 and Hoxd9–13 imply specific roles in limb development. Experimental evidence shows that expression of Hoxa9 and its paralog Hoxd9 specify the scapula, paralogs 10 specify the humerus, 11 the radius and ulna, 12 the carpals, and 13 specify metacarpals and phalanges (Zakany and Duboule, 1999). It has been hypothesized that Hoxa‐d9–13 expression patterns along the proximal–distal axis determines the position of the joints (Duboule, 1992; Yokouchi et al., 1991). Null mutations of paralogs can result in the fusion of some bones, suggesting a role in joint formation, but such analyses are

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complicated by the fact that there is functional cooperation not only between paralogous but also nonparalogous Hox genes (Favier et al., 1996). Skeletal defects due to null mutations of Hox genes are not attributable to changes in patterning but, rather, to changes in chondrogenesis, specifically changes in cell adhesion and proliferation both critical determinants of chondrocyte condensations (Hall and Miyake, 2000). In addition, Nelson et al. (1996) have demonstrated that there is no obvious correlation between Hox gene expression between the hand and foot. The latter report also demonstrated that sequential activation of Hox genes directed by mechanisms acting at the distal tip of the limb bud were not solely responsible for establishing Hox gene expression boundaries in the developing limb bud. Although Hox homeobox‐containing genes are not specifically expressed within the forming joint, Cux1, a vertebrate homologue of the Drosophila cut gene possessing a single homeobox and several novel DNA‐binding motifs (cut repeats), is restricted to the joint interzone in mesenchymal condensations (Lizarraga et al., 2002). The Drosophila cut gene is required for the formation of the fly wing margin and it is also found in the homologous structure in vertebrates, the AER. Cux1 is highly expressed at sites of incipient joint formation; it is present in stage 25 chick wing buds separating the humerus, radius, and ulna rudiments at the site where the elbow joint will subsequently form. Adenoviral expression of Cux1 in diVerentiated cultures of limb mesenchyme results in suppression of chondrocyte diVerentiation. Crucially, Cux1 does not induce either wnt14 or gdf5 expression; therefore, it is probably a downstream target of wnt signaling and regulates a restricted but important aspect of joint interzone formation. Generation of mice null mutants for
1‐crystalline enhancer‐binding factor (
EF1), that contains two Kruppel zinc finger clusters flanking a homeodomain (Funahashi et al., 1993), causes widespread skeletal defects such as hyperplasia of Meckel’s cartilage, shortened mandible and long bones and fused ribs, but also results in fusion of carpal/tarsal bones and joints, fusions of humerus to either radius or ulna, and fusion of the femur and os coxae (hip) (Takagi et al., 1998). Interestingly, skeletal abnormalities only occur when the homeodomain is deleted, and as it does not contribute to DNA binding its functions are thought to be primarily through epistatic interactions with other homeodomain proteins. Notable exceptions to the role of the distal limb bud structure in specifying positional identities have been published demonstrating firstly that the AER maintains proliferation and inhibits cell death of underlying mesenchyme but does not control cell fate (Dudley et al., 2002) and secondly, that in experiments to knock out fgf4 and fgf8 function in mouse fore and hind limbs, normal proximal structures along with abnormally sized distal ones were observed, contrary to what would be expected from the positional identity model of limb development (Sun et al., 2002). One way to reconcile the experimental evidence was to propose that precursors to limb elements

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are prespecified early in limb bud development due to their proximity to proximal or distal signaling centers, which then multiply and survive under the influence of the AER to generate the requisite skeletal structures (Richardson et al., 2004), though, to date, direct evidence for precursor elements is lacking (Wolpert, 2002). The model also proposed that specification of stylopod, zeugopod, and autopod (fore, mid, and distal, respectively) limb domains are uncoupled from segmentation. Here, periodicity of segmentation would be controlled by continual outgrowth within a domain and presumably joint specification through intrinsic gradients of agonistic and antagonistic signaling molecules such that reiterative segmentation within the distal autopodal domain could result in hyperphalangy as is observed in dolphin digits (Fedak and Hall, 2004). The situation is rather diVerent in the axial skeleton where synovial (zygapophysial) joints separate vertebrae. An intervertebral disk composed of an outer external annulus of collagen organized as thick fibrous lamellae and an inner annulus, the nucleus pulposus, populated with chondrocytes expressing collagen type II (Col2) and aggrecan connects adjacent vertebrae. Careful removal of somitocoele cells from somites of 2‐day‐old chick embryos leads to fusion of vertebral bodies due to the absence of a joint, the posterior articular process and intervertebral disk. It is thought that somitocoele cells represent a joint‐forming compartment, the ‘‘arthrotome,’’ whose removal does not aVect the development of any other part of the vertebrae other than the joint (Mittapalli et al., 2005).

B. Formation of the Joint Interzone Wnt14 (also known as wnt9a) plays a central role in initiating synovial joint formation in the chick limb (Hartmann and Tabin, 2001). Wnt14 gene expression is initially detected as a transverse stripe in the presumptive joint regions of future joints at stage 27 in the autopod of developing chick limbs (Fig. 3). Expression of wnt14 then separates to form bipartite stripes that probably demarcate the limits of the forming joint interzone. In later stages of development, wnt14 is observed in parts of the fibrous capsule as well as the synovial lining of the joint capsule. Injection of wnt14 retroviral constructs into the posterior region of developing chick wings results in the formation of multiple gaps comprising condensed mesenchyme within the cartilage elements. The cells within these gaps exhibit weak Safranin‐O staining, are densely packed, and express growth factor gdf5 and collagen type III, a characteristic matrix component of the interzone (Fig. 3). Further analysis showed that genes that are downregulated during normal interzone formation such as sox9, noggin, col2, col9, and aggrecan are similarly downregulated in interzone‐like areas within wnt14 virally infected limbs. In order to

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Figure 3 Expression of wnt14 (A) and gdf5 (B) during phalangeal joint formation in the chick embryo. In situ hybridization of stage 33 autopods with antisense probes to wnt14 describes expression in the presumptive joint regions (arrowhead) and interdigital mesenchyme (arrowed). Expression of gdf5 in the autopod is expressed in the joint region (arrowhead) and also in the adjoining perichondrium that is absent in wnt14‐screened sections (red arrowhead). [Reprinted from Hartmann and Tabin (2001) with kind permission from Elsevier. Copyright (2001).]

confirm that the condensed mesenchyme had adopted the interzone phenotype, in situ hybridization was used to show that various markers previously known to be expressed in developing joints but not in prechondrogenic mesenchyme such as wnt4, CD44rel and chordin were expressed in the gap regions. Hartmann and Tabin also discovered that not only did wnt14 arrest chondrocyte diVerentiation in retrovirally infected chick limb micromass cultures, but crucially, chondrocyte diVerentiation in prediVerentiated micromass cultures could be reversed, reflecting events that occur during in vivo interzone formation. While these marker correlations are persuasive, it would be interesting to test if these cells were also mechanically sensitive in relation to HA synthesis. Of the 19 wnt family members, 3 wnts are expressed in overlapping and complementary patterns in the joint interzones of developing limbs, wnt14, wnt4, and wnt16, and all signal through the canonical wnt‐signaling pathway. Wnt ligands bind to receptors encoded by the Frizzled (Fz) gene family and LRP5/6 that result in the stabilization of cytosolic ‐catenin that is then translocated to the nucleus, where it activates downstream gene expression by binding LEF/TCF transcription factors. Guo et al. (2004) performed micromass assays in which wnt14 adenovirus was added to mouse limb mesenchymal cells that had a conditional targeted allele of ‐catenin. They found that wnt14 adenovirus infection inhibited chondrogenic diVerentiation; however, this eVect was reversed when cultures were serially infected first with Cre‐adenovirus to inactivate ‐catenin followed by wnt14 adenovirus. They also found that serial infection of micromass cultures with wnt14 and dominant‐negative LEF1 that blocks transcriptional activity of ‐catenin also reversed the negative eVects of wnt14 on chondrogenic diVerentiation. Guo et al. (2004) also created transgenic mice expressing a constitutively

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active N‐terminally truncated form of ‐catenin under the control of a Col2a1 promoter/enhancer. Cartilage formation was significantly reduced in mutant mice, and this impairment also aVected endochondral ossification, though not intramembranous ossification, and contributed to perinatal lethality. Chondrocytes in developing limbs lost their characteristic chondrocytic phenotype, as exemplified by loss of sox9 expression and adoption of a joint interzone phenotype characterized by the expression of interzone‐ specific gene products such as collagen types I and III, gdf5, and chordin. As stated earlier, localized production of wnt14 induces the expression of gdf5. Gdf5, gdf6, and gdf7 were cloned in response to data, suggesting further bone morphogenetic protein (BMP) family members than currently known were responsible for normal skeletal development. BMP5 mutations result in the short ear mouse phenotype that aVect the size and shape of axial skeletal components, whereas appendicular elements such as limbs and digits are unaVected (Storm et al., 1994). Conversely, the brachypodism mutation in mice results in altered bone lengths and number of segments in all four limbs but leave the axial skeleton unaltered (Storm et al., 1994). Degenerate PCR was used to identify a new subgroup of BMP proteins including gdf5, gdf6, and gdf7 that shared
83% homology and between 46% and 57% homology with related subgroups. Subsequently, it was shown that only gdf5 mapped to the brachypod disease locus on chromosome 2. Premature translational stop codons caused by frameshift mutations were identified in the gdf5 open reading frame of three independent brachypod mutant mouse strains. Detailed expression analysis of gdf5 gene expression early in limb development shows sequential appearance of transcripts in the proximodistal manner consistent with normal joint formation. Bands of expression are present between 10 days‐postcoitus (dpc) and 15.5 dpc in the shoulder, elbow, and developing digit‐ray, including regions of future joints between the metacarpal and phalanges, a pattern that is also replicated in the hind limb. Initial onset of gdf5 expression precedes by 24–36 hours, the morphological appearance of joint interzones (Fig. 3), and expression continues for up to a further 3 days at a particular site (Storm and Kingsley, 1996). Specifically, mutations in gdf5 cause fusion of all proximal and medial phalanges, fusion of sesamoid bones in digits, fusion of bones in the wrists and ankles, and abnormalities in the structure of the knee, all limb‐specific abnormalities. Cartilage condensations in mutant mice are indistinguishable from condensations in wild‐type mice but segmentation through the formation of interzone structures does not occur. It has been conjectured that one reason why short limbs occur is the reduction in epiphyseal elements resulting in the absence of structures to elongate the bone. Work by Francis‐West et al. (1999) has also shown that gdf5 is expressed in two distinct phases in skeletal development, firstly at

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cartilage condensation where it acts to promote cell adhesion, and secondly by stimulating proliferation of epiphyseal chondrocytes adjacent to the joint interzone. Application of exogenous gdf5 beads into the interdigital region of embryonic chick forelimbs stimulated cartilage growth and consequently inhibited synovial joint formation and caused localized fusion of skeletal elements, syndactylism, between adjacent digits and metacarpal and proximal phalanges (Storm and Kingsley, 1999). The eVect of exogenous gdf5 application was stage dependent and accompanied by an upregulation of proteins associated with cartilage diVerentiation such as collagen type II and ihh and downregulation of markers of developing joints such as gli3 and gdf5 itself. Interestingly, gdf5 downregulation was specific to the joint‐forming regions and not to gdf5 expression in the surrounding perichondrium. Thus, initial gdf5 expression in the digital‐ray is broader than observed in later stages and acts to stimulate cartilage development and then restricts its expression to a narrow stripe between developing cartilage elements. Similar studies in the chick limb also demonstrated that ectopic gdf5 application in the interdigital mesenchyme inhibits joint formation and that this inhibition is a secondary consequence of the action of gdf5 on epiphyseal chondrocytes in the cartilage condensation (Storm and Kingsley, 1999). Most importantly, this study served to show that gdf5 alone is not suYcient to induce joint formation (unlike wnt14) and that joint defects may be a secondary consequence of stimulatory eVects of gdf5 on cartilage development. Gdf6 and gdf7 are also expressed in stripes across developing skeletal condensations prior to segmentation but in a more spatially restricted pattern when compared to gdf5. Gdf6 null mutant mice have joint fusions between specific bones in the wrists and ankles (Settle et al., 2003). Gdf7 mutation in mice leads to defects in seminal vesicle formation and no skeletal defects are present (Settle et al., 2001). The gdf subfamily of growth factors is also capable of inducing synovial joint‐associated structures such as tendons and ligaments. Gdf7 is expressed at the distal tips of digits adjacent to where tendons are formed. Subcutaneous implantation of gdf7 and BMP2 in rats results in formation of neotissues such as ligament, tendon, as well as cartilage and bone (Wolfman et al., 1997). In order to discount redundancy among diVerent gdfs, a gdf5:gdf6 double mutant was made and showed dramatic failures of joint formation (synovial and fibrous) and bone growth aVecting both the appendicular and axial skeletons. Wolfman et al. (1997) had previously discovered that gdf5 is expressed between mesenchymal condensations within the axial skeleton that constitute the vertebral column and that functional redundancy by related BMP family members may mask mutant phenotypes in the axial skeleton. As gdf6 mutant mice still express gdf5 in the interzone, gdf6 is thought to be required at the execution phase of segmentation rather than initiation of

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interzone formation. It has been proposed that some of the skeletal defects in gdf5 or gdf6 mutant mice may be due to reduced movement of aVected joints, it is known that immobilization of joints results in fusion even after cavitation has begun (Pitsillides, 2003), and this may partly explain some of the observable phenotypes. Indeed, recent work has clarified this somewhat by demonstrating that gdf5 expression is not aVected by immobilization of joints in ovo, whereas fgf2 expression bordering the joint forming cavity is selectively diminished (Kavanagh et al., 2006). It is clear that coordination between diVerent chondrogenic signals acting in the developing limb specifies the establishment of joints, epiphysis, and diaphysis. Mutation of noggin, a secreted antagonistic of BMPs and gdfs, results in inhibition of joint formation in mice homozygous for the null mutation (Brunet et al., 1998). Mutant limbs are shorter and broader along the proximodistal axis and joint fusions extend from the elbow down through to the digits. This single‐extended cartilage element expresses a normal suite of BMPs and proteins involved in cartilage maturation such as ihh and pthrp and the element matures synchronously. Joint failure was seen as a failure to induce gdf5 in the presumptive joint region. Work by Merino et al. (1999) showed that ectopic noggin application induces gdf5 expression in the adjacent interdigital mesenchyme, and ectopic BMP7 in the interdigital mesenchyme inhibited gdf5 expression in presumptive joints. The implication of this work is that there is a position within noggin and BMP concentration gradients within which gdf5 expression is induced and disruption of these concentration gradients disrupts normal gdf5 expression in incipient joints causing fusions. Given the conserved nature of BMPs in evolution, it is not surprising that gdf5 homologues such as contact have been discovered in lower vertebrate species. Crotwell et al. (2001) demonstrated that zebrafish gdf5 (contact) expression is first associated with cartilage growth and diVerentiation of mesenchymal condensations in developing endoskeletal supports of the dorsal, anal, and caudal fins. Later, contact is expressed in the segmenting regions of the dorsal and anal fin radials, and is probably required for the restriction of segmentation to appropriate locations, as is the case for tetrapods (Fig. 4).

III. Joint Cavitation and the Role of Mechanical Stimuli The next stage of joint development is the formation of the fluid‐filled joint space separating the opposing skeletal elements, cavitation, followed by the morphogenesis of these elements into interlocking structures. Classical studies have demonstrated the necessity of mechanical stimuli in developing joints. Using a number of in ovo methods including the addition of decamethonium bromide, botulinum, and succinylcholine or simply

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Khan et al. A Dorsal fin

Distal radial Proximal radial

B

Figure 4 Expression of gdf5 homologue contact in zebrafish (Danio rerio) fin radials. The dorsal fin rays (A) are supported by endoskeletal rods of cartilage, known as radials (blue) [adapted from Bird and Mabee (2003)]. Gdf5 is expressed in the mesenchyme between the youngest developing radials (B). Segmentation begins in the most anterior radials and continues posteriorly. Later in development, radials segment proximodistally into two parts, large proximal and smaller distal radials. Gdf5 expression is restricted along the anterior length of proximal and distal radials prior to segmentation, the distal radials articulate directly with fin rays. [Reprinted from Crotwell et al. (2001) with kind permission of Springer Science and Business Media. Copyright (2001).]

by the maintenance of limbs in organ culture as a means of removing mechanical stimuli and, in already cavitated joints, the removal of the mechanical stimulus resulted in cartilaginous fusions at the articular surfaces (Drachmann and SokoloV, 1966; Fell and Canti, 1934; Hamburger and Waugh, 1940; Mitrovic, 1982; Osborne et al., 2002). These studies indicate that embryonic limb movement plays an essential role in joint cavity formation and maintenance but they do not demonstrate mechanistically how movement contributes to joint cavitation. Movement may result in creating diVerential growth patterns, by physically disrupting tissue or by altering local extracellular matrix (ECM) composition at the developing joint line.

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Work has shown that there is very little evidence for proliferation in interzonal areas of the developing joint compared to that in cartilaginous areas, thus suggesting that mechanical stimuli are required for joint cavitation and are modulated by local diVerential patterns of growth (Edwards, 1994; Kavanagh et al., 2002; Lewis, 1977; Wolpert et al., 1975). Other work, however, where immobilization has been induced, shows mild eVects on the growth of limbs with only slight delays observed in development (Drachmann and SokoloV, 1966; Murray and Drachman, 1969). Thus, it seems that diVerential growth is insuYcient to act solely as a mediator of mechanical stimuli in joint cavitation. The disruption of cell–cell cohesion at the joint line arising from mechanical forces generated by muscle activity could allow separation to occur. This process, however, seems unlikely as joint development is precisely controlled up until this point and to disrupt the precise mechanisms with imprecise mechanical forces as a means of tissue separation seems unlikely. In addition, Hasty et al. (1993) have shown that in myogenin‐deficient mice, the joint forms even though there is no contracting skeletal muscle to generate mechanical forces. At present, evidence points toward movement‐induced mechanical stimuli being responsible for altering the synthesis and degradation of ECM components along the joint line resulting in cavitation. There is strong evidence to suggest that HA is one of these components. HA is classed as an unsulfated GAG, first discovered as the metachromatic substance at the joint line by Anderson and Brorasmussen (1961) and was confirmed to be HA (Anderson, 1961). Originally considered to act as a space‐filling molecule, further evidence now suggests HA plays a much more critical role in joint development, especially when Craig et al. (1990) demonstrated the presence of HA at the joint line concomitant with the first signs of cavitation. The enzyme uridine diphosphoglucose dehydrogenase (UDPGD) is essential in providing the precursors for HA synthesis, and research using enzyme histochemistry has demonstrated the presence of UDPGD activity in cells of the interzone just prior to cavitation and within the articular surfaces and synovium during joint cavitation (HinchcliVe, 1977; Pitsillides et al., 1995) (Fig. 5). Presence of the HA‐binding protein CD44 on the interzonal cells emphasizes the importance of HA in joint cavitation. High levels of HA observed along the joint line would, therefore, promote cell separation by saturation of HA‐ binding proteins (HABP) accounting for the loss of tensile properties in the ECM (Toole, 1981) (Fig. 6). Indeed, the administration of exogenous HA‐ oligosaccharides which are able to disrupt HA–HABP interactions in ovo blocked joint cavitation with fused joints resembling those in immobilization studies (Dowthwaite et al., 1998; Fell and Canti, 1934). Further analysis of joints disrupted with HA oligosaccharides revealed interzone regions of these limbs with decreased CD44 levels, decreased UDPGD activity, and changes in alcian blue staining, strongly suggesting that mechanical influences are

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CD44 UDPGD HAS Free HA Cell-bound HA

Uncavitated

Cavitated

Figure 5 Schematic representation of hyaluronan‐related events during joint cavitation. Before cavitation, both the interzone (gray) and the articular surfaces express CD44. There is little UDPGD activity within the interzone, although activity is present in the articular surfaces. HAS is also expressed in the articular surfaces before cavitation. Both free and bound HA are present in the interzone at this stage. After cavitation, the articular surfaces and the synovium bind HA via CD44, while the cavity contains only free HA. UDPGD activity and HAS expression are maintained in the synovium and the articular surfaces. [Reprinted from Archer et al. (2003) with kind permission of Wiley. Copyright (2003).]

required for HA synthesis and thus functional joint formation (Bastow et al., 2005; Dowthwaite et al., 1998). Interestingly, it has also been shown that gdf5 protein expression is moderately reduced on mechanical stimulation or UDPDG overexpression and allied to the fact that antibody localization of gdf5 appears to show a broad band of expression which suggests the function of gdf5 may be prior to cavitation (Kavanagh et al., 2006). Joint tissue during development is, however, subjected to several mechanical strains. In ovo work by Osborne et al. (2002) has demonstrated the eVects of blocking sustained physiological static loading of joints due to the sustained contraction of particular muscle groups (static) and blocking stimuli from the intermittent and discontinuous contractions of the same muscle groups (dynamic). Neuromuscular blockers decamethonium bromide and pancuronium bromide were administered to induce rigid or flaccid paralysis, respectively. The removal of both static and dynamic stimuli prevented cavity formation and reduced epiphyseal breadth and longitudinal length by decreasing cartilage volume but not bone volume. Reductions in HA were observed in the interzone regions of the joint with menisci and ligaments failing to develop. When rigid paralysis was induced in already cavitated joints, fewer and less severe fusions were observed compared to those

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1. The Development of Synovial Joints

HA

CD44

Cell aggregation (condensation)

Cell separation (cavitation)

Figure 6 Schematic representation of cell surface hyaluronan (HA) binding and its eVect on cell aggregation (condensation) and cell separation. Cells expressing CD44 and surrounded by HA maintain their position relative to one another (top). A decrease in HA concentration between the two cells will cause cell aggregation as occurs during cartilage condensation and an increase in HA concentration between the cells will cause separation of the cells as in cavitation. [Reprinted from Archer et al. (2003) with kind permission of Wiley. Copyright (2003).]

embryos subjected to flaccid paralysis. These data suggest that mechanical stimuli from static loading contributes diVerently to joint maintenance and, once established, the joint cavity can maintain its function if only subjected to static loads. With time, the embryo becomes more restricted and so signals from dynamic stimuli would decrease. It is, therefore, important that stimuli from static movement are able to maintain a functioning joint cavity. The above studies have demonstrated the eVects of reducing mechanical stimuli on developing joints and, more recently, evidence is accumulating which reports on the eVects of the application of mechanical strain to cells of the developing joint. When chick fibrocartilage cells were subjected to a 10‐min period of 3600me, a strain magnitude that is equivalent to those engendered by normal load‐bearing on bone, significant increases in HA release were demonstrated along with increased UDPGD activity and HABP expression (Dowthwaite et al., 1998; Lanyon, 1996). Momberger et al. (2005) have also shown that, in isolated rabbit synoviocytes, a short period (10min) of mechanical stretch was just as eVective in producing a significant increase in HA release as a sustained stretch of 180 min.

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An important gene involved in HA synthesis is HA synthase (HAS) which exists in three isoforms, derived from three separate genes HAS1, HAS2, and HAS3 (Spicer and McDonald, 1998) which have been shown in vitro to be responsible for synthesizing HA of diVerent chain lengths and at diVerent rates. In synoviocytes, it has been demonstrated that HAS1 and HAS2 are in similar abundance, whereas HAS3 is the least abundant (Recklies et al., 2001). DiVerential expression of the three HAS genes suggest that each plays an important role in maintaining joint function. The application of mechanical strain to both chick fibrocartilage cells and rabbit synoviocytes showed no increase in HAS2 expression yet an increase in HAS3 expression was induced on mechanical strain in chick fibrocartilage cells (Dowthwaite et al., 1998). It does need to be taken into consideration, however, that levels of media HA concentrations have been shown to correlate with HAS mRNA expression levels in some studies while in others this is not the case (Jacobson et al., 2000; Recklies et al., 2001). The evidence presented strongly suggests that mechanical cues stimulate cells of the developing joint to release HA and recent data suggests that de novo synthesis of HA occurs as a result of these mechanical stimuli. Indeed, it is unlikely that all of the observed HA increase is released from the preformed pericellular HA coat as this would not account for the high levels of HA seen after cells have been subjected to stretch (Anggiansah et al., 2003; Coleman et al., 1997; Price et al., 1996). In order to determine the eVects of diVerential gene expression on de novo HA synthesis, addition of the translational inhibitor, cycloheximide and transcriptional inhibitor, actinomycin D to stretched synoviocytes resulted in a loss of the observed increases in HA release that were associated with stretch (Momberger et al., 2005). Whether these inhibitors are targeting the HAS genes or others involved in HA synthesis remains to be determined at this present time, but it is clear that changes in gene expression are necessary for mechanical stimuli to result in increased HA synthesis. Protein kinase C has been shown to increase HAS activity through phosphorylation and it is interesting to note that activation of PKC in rabbit knee joints in vivo and in rabbit synoviocytes in vitro results in increased HA release, whereas blocking PKC activity results in a decrease in HA release (Anggiansah et al., 2003; Klewes and Prehm, 1994; Momberger et al., 2003). Additionally, the activation of both PKC and PKC
by stretch supports a role for mechanical stimuli in HA release. Taking all these data into consideration, it has been suggested that mitogen‐activating protein kinase (MAPK) signaling could be involved in the mechanotransduction pathway which results in HA synthesis. Further to this suggestion, Ca2þ channel activators were used to demonstrate activation of PKC resulting in subsequent increases in HA, suggesting Ca2þ ion flux might be responsible for the phosphorylation and subsequent activation of PKC which then leads to

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HA release (Momberger et al., 2003). Conversely, the addition of Ca2þ chelators resulted in decreased HA release (Momberger et al., 2003). Downstream of the MAPK‐kinase pathway, evidence exists to suggest that extracellular‐signal‐related kinases (ERK1/2) are important in the mechanotransduction pathway (Bastow et al., 2005; Momberger et al., 2005). Phosphorylated ERK1/2 has been found to be present at the sites of cavity formation and is also preserved after cavitation (Bastow et al., 2005; Ward et al., 1999). The immobilization of limbs reduced phosphorylated ERK1/2 at the developing joint, whereas mechanical strain induced more cytoplasmic phosphorylated ERK1/2 (Bastow et al., 2005). On addition of Ca2þ chelators, which block PKC activity as mentioned previously, downstream ERK1/2 activity is subsequently reduced and as a consequence decreased HA release is observed due the lack of ERK1/2 phosphorylation. If indeed the MAP‐kinase pathway is necessary for the transduction of mechanical stimuli to increase HA release, then it seems likely that phosphorylation of PKC via increased Ca2þ ion flux is necessary for the activation of ERK1/2. Activation through phosphorylation of ERK1/2 could result in its translocation to the nucleus allowing further phosphorylation of gene regulatory proteins. Subsequent changes in gene expression and possibly HAS gene expression could follow leading to the observed increases in HA synthesis on mechanical stimuli. Following cavitation, the skeletal elements undergo morphogenesis. The mechanisms underlying morphogenesis of the opposing skeletal elements into the diverse interlocking structures remains to be fully elucidated and is the most poorly understood process of joint development (Pacifici et al., 2005).

IV. Cell–Matrix Interactions During Joint Formation In conjunction with the signaling events occurring during the process of cavitation, there are a number of important cell–cell and cell–matrix interactions that are vital for the development and patterning of a fully functional diarthrodial joint. For normal joint formation and chondrogenesis to occur, the coordinated expression of a large number of ECM glycoproteins and cell surface receptors is required. The presence and distribution of ECM components during joint formation have, therefore, attracted much interest due to the dynamic nature in which these components are assembled and interact. More specifically, glycoproteins present in the ECM of joint‐associated tissues are involved in proliferation, migration, diVerentiation, and apoptosis during joint formation and are, thus, important regulators of cell behavior (Adams and Watt, 1993). During development, a number of ECM glycoproteins show specific spatial and temporal distributions that functionally contribute to the overall patterning and integrity of the joint.

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Studies have shown that tenascin‐C, a major extracellular glycoprotein, plays a vital role in skeletal development, in particular, during chondrogenesis (Koyama et al., 1995; Pacifici, 1995; Pacifici et al., 1993). Throughout development of the joint, the expression levels of tenascin alter with cell type and stage of diVerentiation of the joint‐associated tissues. Immunohistochemical studies have shown that tenascin is expressed in the developing articular cartilage, perichondrium, synovial lining, and junction between the growth plate and epiphysial bone in rat, mouse, and chick animal models (Chiquet and Fambrough, 1984; Mackie and Ramsey, 1996; Pacifici et al., 1993). Within the joint specifically, tenascin is intimately involved in articular cartilage development with expression first observed in the chondrogenic mesenchyme of early developing mouse and avian limb buds (Pacifici et al., 1993). As development proceeds, the expression of tenascin becomes more localized to the territorial matrix of the superficial articular cartilage and then to a thin strip on the articular surface. This pattern of expression, however, recedes over time and is absent with the onset of maturity and is, therefore, not retained in mature articular cartilage (Chiquet and Fambrough, 1984; Mackie and Ramsey, 1996; Melnick et al., 1981; Pacifici et al., 1993). There is functional evidence for the role of tenascin throughout joint development. With the appearance of tenascin during the onset of early cartilage diVerentiation, studies suggest that tenascin is involved in the stimulation of chondrogenesis and the establishment of the chondrogenic phenotype by facilitating the detachment of cells from the fibronectin‐containing ECM thereby causing cell rounding and condensation (Chiquet and Fambrough, 1984; Mackie and Murphy, 1998; Mackie et al., 1987). Studies have also shown the ability to inhibit avian wing bud cells from attaching to fibronectin‐coated substrates in the presence of tenascin (Mackie et al., 1987). Reports also suggest that the role of tenascin‐C during cartilage formation is to establish a barrier between skeletal elements (Chiquet and Fambrough, 1984; Erickson and Bourdon, 1989). The antiadhesive nature of tenascin has been extensively reported during cartilage formation especially in the formation of the interzone between developing joints resulting in the creation of two contiguous but distinct skeletal elements. Additionally, Salter (1993), reported an increase in expression of tenascin within the joint with the onset of diseases such as OA and RA. A recapitulation of expression levels similar to that seen during development in the surface zone of the articular cartilage and in the vasculature of the synovium suggests, therefore, a possible reparative response by the joint to disease (Salter, 1993). Thrombospondins are a family of five distinct but related adhesive glycoproteins that are thought to play a role in chondrogenesis and joint formation. Although each member, which includes cartilage oligomeric matrix protein (COMP), is expressed in locations complementary to regions of

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the limb involved in chondrogenesis and joint formation, the functional significance of the presence of these proteins has not yet been elucidated despite the belief that they influence these processes (Mackie and Murphy, 1998; Tucker et al., 1997). Thrombospondins 1, 2, 3, 4 and COMP generally appear sequentially as development proceeds. Thrombospondins 1 and 2 are expressed in early chondrogenic mesenchyme with expression patterns similar to that of tenascin‐C, later becoming restricted to peripheral regions of cartilage (O’Shea and Dixit, 1988; Tucker, 1993; Tucker et al., 1995, 1997). Thrombospondins 3 and 4 are expressed later in development in regions of proliferating chondrocytes of the growth plate (Iruela‐Arispe et al., 1993; Tucker et al., 1997). It appears that the remaining thrombospondin family member, COMP, is expressed when chondrogenesis is well advanced and is, therefore, absent in regions of cartilage formation prior to cavitation (Franzen et al., 1987). Once expressed, COMP is abundant in the territorial matrix surrounding the maturing articular chondrocytes and may enhance the load‐bearing characteristics of the matrix (Hedbom et al., 1992; Murphy et al., 1999). Syndecan‐3, a member of the heparin sulfate family of transmembrane proteoglycans, plays a major role in limb morphogenesis. Although unclear, the spatial and temporal distribution of syndecan‐3 in regions of nascent joint development suggests a role in the onset of joint diVerentiation (Gould et al., 1995; Koyama et al., 1995). Syndecan‐3 is also expressed in the chondrogenic mesenchyme and may also play a role in regulating chondrogenesis, although this expression is lost in early cartilage models (Gould et al., 1995). More specifically, syndecan‐3 is a cell surface receptor and coreceptor for growth factors and signaling molecules (Kosher, 1998; Shimazu et al., 1996). Studies suggest that syndecan‐3, therefore, may modulate signaling molecules such as gdf5 (a heparin‐binding BMP‐like signaling molecule) that promote cell adhesion and proliferation indicative of initial stages of condensation and joint specification (Buxton et al., 2001; Francis‐West et al., 1999; Storm and Kingsley, 1996). Fibronectin, an ECM glycoprotein, is expressed in early chondrogenic mesenchyme (Melnick et al., 1981) and is thought to play an inhibitory role during chondrogenesis (Pennypacker et al., 1979; Swalla and Solursh, 1984; West et al., 1979) although it has been suggested that fibronectin is possibly required for the induction of precartilage condensations (Frenz et al., 1989; Mackie and Murphy, 1998). Although similar expression patterns are observed during early chondrogenesis between tenascin‐C and fibronectin, fibronectin is expressed in surrounding areas of nonchondrogenic mesenchyme and is also retained in mature cartilage (Dessau et al., 1980; Melnick et al., 1981). A paper by Garciadiego‐Cazares et al. (2004) may reignite interest in the role of cell–matrix interactions in joint development and the fate of

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chondrocytes since this topic has evoked little interest in recent years due partly to the current interest in signaling molecules expressed during these developmental events. Integrins, ECM cell surface receptors, are emerging as key players in the fate of diVerentiating chondrocytes. Although the role of integrins in skeletal development is poorly understood, a number of studies have highlighted the importance of integrins in both joint formation and chondrocyte diVerentiation (Bokel and Brown, 2002; Hynes, 2002). The study by Garciadiego‐Cazares et al. (2004) showed that when 51 integrin expression is blocked during development, ectopic joint formation is induced in mouse limb buds. The ectopic joint also induced molecular changes characteristic of joint formation with the expression of specific joint markers such as wnt14 (Hartmann and Tabin, 2001) and gdf5 (Buxton et al., 2001; Francis‐West et al., 1999). In addition, the expression of ECM components characteristic of articular cartilage such as type II collagen was observed. Results also showed that the ectopic joint was new with the dediVerentiation of the chondrocytes associated with changes in ECM components, collagen types I and II and aggrecan, typical of the changing of cellular fate during joint development. The ectopic joint formation was observed between the boundary of proliferating chondrocytes and hypertrophic cartilage, thereby inhibiting prehypertrophic chondrocyte diVerentiation in this region. Misexpression of 51 integrin, however, induced fusion of joints and the diVerentiation of prehypertrophic cells. This study suggests that the location of the ectopic joint may be crucial to our understanding of the process of joint formation, the decision of cell fate among developing chondrocytes, and whether the cells pursue a prehypertrophic program of diVerentiation or participate in joint formation. In conclusion, due to the highly specific nature of the ECM within the joint and its inherent functional requirements, it is of little wonder that the ECM plays a pivotal role in the process of joint diVerentiation and the establishment and maintenance of the structural integrity of the mature phenotype.

V. The Development of Articular Cartilage: Overview There is a traditional view that articular cartilage represents the remnants of the embryonic epiphyses that fails to be replaced by bone through endochondral ossification. While this may prove to be the case, the idea is increasingly being challenged. In particular, there is a notion that articular cartilage arises from a population of cells distinct from that of the embryonic cartilage (Archer et al., 1994; Bland and Ashhurst, 1996b; Ito and Kida, 2000). It is proposed that this subpopulation occupies the very surface cell layer of the newly cavitated joint. As such, the cells may be derived from this region of the embryonic epiphysis or from the cells of the now dispersed interzone

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(Ito and Kida, 2000). In the absence of experimental studies, this issue remains to be resolved. In either case, the cells would be expected to be prespecified. To date, most studies have concentrated on charting the spatiotemporal distributions of matrix components in a number of mammalian species. In general, collagen expression tends to be site‐specific during development, and does not tend to change greatly. Consequently, type II collagen is ubiquitous, type I collagen is restricted to the articular surface, and type III collagen is pericellular. In the basal region, type X collagen is expressed during the establishment of the final tidemark (Bland and Ashhurst, 1996a; Morrison et al., 1996). Interestingly, type V collagen is seen pericellularly in the chondrogenous layers (along with type III collagen) during cavitation, and then within the articular cartilage after cavitation, suggesting that the chondrogenous cells of the interzone give rise to articular cartilage. In contrast to the collagens, the proteoglycans appear to be more dynamic during development. In general, the small leucine‐rich proteoglycans are largely absent from the embryonic epiphyses, but they do label the presumptive articular cartilage. During maturation the pattern becomes more variable, which may depend on age, joint, or species. For example, in the South American opossum (Monodelphis domestica), fibromodulin, decorin, and biglycan become localized to the upper half of the tissue depth. A similar distribution is also observed in the rabbit knee (Archer et al., 1996; Kavanagh and Ashhurst, 1999; Murphy et al., 1999). While the matrilins (1–4) are broadly and dynamically expressed in the embryonic epiphyses, their distribution in articular cartilage is much more restricted to peripheral areas of the joint, or may be absent altogether (Kavanagh and Ashhurst, 1999; Klatt et al., 2002). Finally, it appears that COMP is absent from the presumptive articular cartilage, but after cavitation, it accumulates in the mid‐zone of the tissue during chondrocyte maturation (Murphy et al., 1999). Gepstein et al. (2002) documented the expression of metalloproteinases in articular cartilage. Using the mouse knee as a model, it was found that for MMP2, MMP‐3, MMP‐9, and MMP‐13, a peak of expression was observed in the surface and transitional zones at 2 weeks, compared with similar joints at 1 day and 18 months.

VI. Aspects of Postnatal Joint Biology Once the synovial joint is functional, are the signaling molecules responsible for its formation and growth still responsible for maintaining its integrity throughout the life of the organism? To address this question, Roundtree et al. (2004) created a mouse line that expressed Cre recombinase under the control of a gdf5 promoter that when crossed with a mouse expressing a floxed BMP receptor 1a (BMPr1a) resulted in conditional inactivation of

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this allele in developing joints, adult articular cartilage, and the synovial capsule. BMPr1b is the high‐aYnity receptor for gdf5, so joint specification and formation was not disturbed; however, lack of BMPr1a receptor that binds BMPs2/4 resulted in joint fusions in the wrist and ankles of aVected mice. Most joints in mutant mice, however, were indistinguishable from those in wild‐type mice at least until birth. Shortly after birth, mutant mice showed decreased levels of aggrecan and collagen type II expression, though curiously sox9 levels were unaVected. After 7 weeks, early histological signs of osteoarthritis such as little or no Safranin‐O staining, acellularity and surface fibrillation were observed in articular cartilage of mutant mice. By 9 months large areas of the knee joints of mutant mice were devoid of cartilage and were showing signs of subchondral sclerosis. Loss of responsiveness to transforming growth factor  (TGF‐) through the generation of mice expressing a dominant‐negative allele of TGF‐ type II receptor in articular cartilage also leads to osteoarthritis through replacement of normal articular chondrocytes with cells expressing collagen type X, a marker of terminal chondrocyte diVerentiation (Serra et al., 1997). These two studies highlight that not only maintenance of joint integrity requires continual growth factor stimulation, but also loss of articular cartilage causes significant alterations in underlying bone as observed in increased ihh expression (Serra et al., 1997) and increased collagen type X expression (Roundtree et al., 2004) again reinforcing the view that articular cartilage is a signaling center. Reconstitution or reinforcement of growth factor‐signaling pathways is a technique that is prevalent in tissue engineering; gdf5 has been used to aid healing of ligaments, tendons and in bone augmentation (Rickert et al., 2005; Tashiro et al., 2006; Yoshimoto et al., 2006), and BMPs 2, 4, and 7 in cartilage and bone repair (Reddi, 2003). As with all therapies, caution must be used in activating signaling cascades without a thorough understanding of all the implications. Untimely activation of BMP signaling can contribute to accelerated disease progression as described in a mouse model of spondylarthritis (Lories et al., 2005), and it is well known that exogenous application of BMPs or TGF‐s induces osteophytes and ectopic bone formation in the joint (van Beuningen et al., 2000). The Notch‐signaling protein family plays a significant role in peri‐ and postnatal articular cartilage biology. The Notch family is a highly conserved family of cell surface‐signaling molecules that regulate many developmental cell fate decisions which can be either inductive (e.g., arterial endothelial cell specification) (Lawson et al., 2001) or lateral‐inhibitory (e.g., neuron inhibition) (Artavanis‐Tsakonas et al., 1995). The family consists of four Notch receptors: Notch1, Notch2, Notch3, and Notch4 and several receptors: Jagged1 and Jagged2, Delta1, Delta2, Delta3, and Delta4 and receptor activation by ligand mediates transcription via several transcription factors

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(e.g., CSL, SuH, and Espl) (Weinmaster, 1997). Extensive transgenic mutation studies have shown that mutations in Jagged2 describe the entire contribution of the Notch pathway to early limb development in the mouse (Pan et al., 2005). Conditionally activated limb‐specific Jagged2 mutants, single and compound Notch receptor, and ‐secretase mutations all produce syndactylism, indicating Notch signaling plays a nonautonomous role in digit septation and osseous fusion. Perinatal distribution of Notch family members in mice from e15 to 3 months of age is more dynamic with Notch1 expressed by chondrocytes of the developing articular surface but becoming increasingly restricted to the deeper layers after birth, while Notch2 and Notch4, Delta1 and Jagged2 show a broadly similar distribution being present throughout the articular cartilage prenatally and becoming increasingly restricted to deeper layers with age. Notch3 and Jagged1 are absent from developing articular cartilage but are present in deeper layers at later time points (>1‐month postnatal) (Fig. 7). Earlier work, analyzing cell kinetics by in vivo labeling of articular cartilage, led to the hypothesis that a progenitor/stem cell population resided at the surface of articular cartilage (Fig. 8). In these experiments, the thymidine analogue bromodeoxyuridine (BrDU) was injected into the developing knee joints of 2‐month‐old marsupials (M. domestica) every other day for 2 weeks. Unlike thymidine, where the cells can continue to divide, BrDU often blocks further division after intercalation into the DNA at S‐phase that subsequently can be detected by immunocytochemistry. Chondrocytes of the transitional zone were labeled after 4 days of injections. However, after 10 days a small cohort of cells in the surface zone became immunoreactive. One month after cessation of injections, the transitional zone was markedly depleted of cells (but this was reestablished by 6 months) (Hayes et al., 2001). This putative stem/progenitor cell population was finally isolated from the surface of articular cartilage by diVerential adhesion to fibronectin (Dowthwaite et al., 2004), was capable of forming colonies from an initially low seeding density, and was able to engraft into various tissues of the connective tissue lineage and undergo extensive expansion in vitro without dediVerentiation. In addition, it was demonstrated that isolating Notch1 positive cells from the articular cartilage surface significantly increases their clonality. Chemically blocking the Notch‐signaling pathway using ‐secretase inhibitor DAPT (N‐[N‐(3,5‐diflurophenylacetate)‐L‐alanyl]‐(S)‐ phenylglycine t‐butyl ester) abolished clonality and this abolition could be reversed by constitutively activating Notch1 (Dowthwaite et al., 2004) (Fig. 9). Many studies have concluded that following injury and during disease, chondrocytes undergo reversion to a more immature phenotype, this immature chondrocyte then presages the terminally diVerentiated chondrocyte

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Figure 7 Notch family distribution in neonatal mice knees. Notch1 is distributed in the articular surface (A) and Notch2 is ubiquitous (B) in neonatal cartilage. In contrast, Notch3 (C) is absent from the developing cartilage of the knee joint while Notch4 (D) similar to Notch2 is widespread in the neonatal knee. The Notch ligands Delta and Jagged2 (E and G) are expressed widely in the neonatal knee articular cartilage while Jagged1 (F) is only expressed in the deeper layers of the neonatal cartilage. [Adapted from Hayes et al. (2003).]

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Figure 8 The surface zone of articular cartilage contains a progenitor cell population. Progenitor cells in the articular surface divide to give a new progenitor cell and a transit amplifying cell within the transitional zone. The transit amplifying cell can undergo up to five more cell divisions (of shorter cell cycle time) along the chondrocyte diVerentiation pathway and, thus, contribute to the appositional growth of the tissue. [Adapted from Hayes et al. (2001).]

phenotype that has been shown to be prevalent in the terminal stages of osteoarthritis (von der Mark et al., 1977). Evidence for adoption by articular chondrocytes of a less diVerentiated phenotype in osteoarthritic tissue includes reexpression of procollagen type IIA that is originally expressed at chondrogenesis during early development (Ryan and Sandell, 1990) and increased expression of cell surface markers CD105 and CD166 whose coexpression is thought to define a subset population of mesenchymal stem cells (Alsalameh et al., 2004). Reversion to a chondroprogenitor phenotype during early repair following cartilage injury probably activates normally quiescent chondrocytes to proliferate and repopulate the matrix (Tew et al., 2000), and in latter stages of disease may act to prevent terminal diVerentiation of chondrocytes (Watanabe et al., 2003). To conclude, although a reasonable amount of information concerning the basic mechanisms of joint development is known, we know little of the

28 CFE (mean ± s.e.m)

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*

0.4 0.3 0.2 0.1 0 Surf DAPT

Surf

Deep DAPT

Deep

Figure 9 The surface zone of immature bovine articular cartilage contains Notch1 positive cells. After diVerential adhesion to fibronectin, inhibiting Notch signaling by using the ‐secretase inhibitor DAPT abolishes the colony forming eYciency of surface zone cells (Surf DAPT), suggesting that Notch signaling plays an important role in the clonality of chondroprogenitor cells. [Adapted from Dowthwaite et al. (2004).]

specific mechanisms that regulate the diVering tissue types within the joint and how these might be integrated by various signaling pathways. For example, we know that gdf5, which may be used as a marker of the joint interzone, is important both for the morphogenesis of certain joints and for the development of ligamentous structures as in the knee. It seems certainly the case that wnt14 is central in the specification of the joint interzone and that on its formation the interzone becomes an important signaling center that can modulate the cellular activities in the opposing cartilaginous epiphyses and alter the growth of these elements. An intriguing facet that had yet to be resolved is the actual origin of articular cartilage itself. The notion that articular cartilage is derived from cells within the interzone or ‘‘chondrogenous’’ cells at the periphery of the embryonic epiphyses is attractive and defining the nature of these cells compared with the chondrocytes within the epiphyses will be an important undertaking. The identification of a progenitor population within articular cartilage itself may help in defining the original population of ‘‘articular chondrocytes stem/progenitor cells.’’ Finally, the fact that we can culture and expand these progenitor cells extensively without the loss of cartilage phenotype expression opens the possibility of utilizing them for the biological repair of articular cartilage.

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Morrison, E. H., Ferguson, M. W., Bayliss, M. T., and Archer, C. W. (1996). The development of articular cartilage: I. The spatial and temporal patterns of collagen types. J. Anat. 189(Pt. 1), 9–22. Murphy, J. M., Heinegard, R., McIntosh, A., Sterchi, D., and Barry, F. P. (1999). Distribution of cartilage molecules in the developing mouse joint. Matrix Biol. 18, 487–497. Murray, P. D., and Drachman, D. B. (1969). The role of movement in the development of joints and related structures: The head and neck in the chick embryo. J. Embryol. Exp. Morphol. 22, 349–371. Naiche, L. A., and Papaioannou, V. E. (2003). Loss of Tbx4 blocks hindlimb development and aVects vascularization and fusion of the allantois. Development 130, 2681–2693. Nelson, C. E., Morgan, B. A., Burke, A. C., Laufer, E., DiMambro, E., Murtaugh, L. C., Gonzales, E., Tessarollo, L., Parada, L. F., and Tabin, C. (1996). Analysis of Hox gene expression in the chick limb bud. Development 122, 1449–1466. Niedermaier, M., Schwabe, G. C., Fees, S., Helmrich, A., Brieske, N., Seemann, P., Hecht, J., Seitz, V., Stricker, S., Leschik, G., Schrock, E., Selby, P. B., et al. (2005). An inversion involving the mouse Shh locus results in brachydactyly through dysregulation of Shh expression. J. Clin. Invest. 115, 900–909. Niswander, L. (2002). Interplay between the molecular signals that control vertebrate limb development. Int. J. Dev. Biol. 46, 877–881. O’Shea, K. S., and Dixit, V. M. (1988). Unique distribution of the extracellular matrix component thrombospondin in the developing mouse embryo. J. Cell Biol. 107, 2737–2748. Osborne, A., Lamb, K., Lewthwaite, J., Dowthwaite, G. P., and Pitsillides, A. A. (2002). Short‐ term rigid and flaccid paralyses diminishes growth of embryonic chick limbs and abrogate joint cavity formation but diVerentially preserve pre‐cavitated joints. J. Musculoskel. Neuron Interact. 2, 448–456. Pacifici, M. (1995). Tenascin‐C and the development of articular cartilage. Matrix Biol. 14, 689–698. Pacifici, M., Iwamoto, M., Golden, E. B., Leatherman, J. L., Lee, Y. S., and Chuong, C. M. (1993). Tenascin is associated with articular cartilage development. Dev. Dyn. 198, 123–134. Pacifici, M., Koyama, E., and Iwamoto, M. (2005). Mechanisms of synovial joint and articular cartilage formation: Recent advances, but many lingering mysteries. Birth Defects Res. C Embryo Today 75, 237–248. Pan, Y., Liu, Z., Shen, J., and Kopan, R. (2005). Notch1 and 2 cooperate in limb ectoderm to receive an early Jagged2 signal regulating interdigital apoptosis. Dev. Biol. 286, 472–482. Parr, B. A., and McMahon, A. P. (1995). Dorsalizing signal Wnt‐7a required for normal polarity of D‐V and A‐P axes of mouse limb. Nature 374, 350–353. Pennypacker, J. P., Hassell, J. R., Yamada, K. M., and Pratt, R. M. (1979). The influence of an adhesive cell surface protein on chondrogenic expression in vitro. Exp. Cell Res. 121, 411–415. Pitsillides, A. A. (2003). Identifying and characterizing the joint cavity‐forming cell. Cell Biochem. Funct. 21, 235–240. Pitsillides, A. A., Archer, C. W., Prehm, P., Bayliss, M. T., and Edwards, J. C. (1995). Alterations in hyaluronan synthesis during developing joint cavitation. J. Histochem. Cytochem. 43, 263–273. Price, F. M., Levick, J. R., and Mason, R. M. (1996). Changes in glycosaminoglycan concentration and synovial permeability at raised intra‐articular pressure in rabbit knees. J. Physiol. 495 (Pt. 3), 821–833. RatcliVe, A., and Mow, V. C. (1996). Articular cartilage. In ‘‘Extracellular Matrix: Tissue Function’’ (W. D. Comper, Ed.), Vol. 1, pp. 234–302. Harwood Academic Publishers.

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Recklies, A. D., White, C., Melching, L., and Roughley, P. J. (2001). DiVerential regulation and expression of hyaluronan synthases in human articular chondrocytes, synovial cells and osteosarcoma cells. Biochem. J. 354, 17–24. Reddi, A. H. (2003). Cartilage morphogenetic proteins: Role in joint development, homoeostasis, and regeneration. Ann. Rheum. Dis. 62(Suppl. 2), ii73–ii78. Redman, S. N., Oldfield, S. F., and Archer, C. W. (2005). Current strategies for articular cartilage repair. Eur. Cell. Mater. 9, 23–32; discussion 23–32. Richardson, M. K., JeVery, J. E., and Tabin, C. J. (2004). Proximodistal patterning of the limb: Insights from evolutionary morphology. Evol. Dev. 6, 1–5. Rickert, M., Wang, H., Wieloch, P., Lorenz, H., Steck, E., Sabo, D., and Richter, W. (2005). Adenovirus‐mediated gene transfer of growth and diVerentiation factor‐5 into tenocytes and the healing rat Achilles tendon. Connect. Tissue Res. 46, 175–183. Riddle, R. D., Ensini, M., Nelson, C., Tsuchida, T., Jessell, T. M., and Tabin, C. (1995). Induction of the LIM homeobox gene Lmx1 by WNT7a establishes dorsoventral pattern in the vertebrate limb. Cell 83, 631–640. Roundtree, R. B., Schoor, M., Chen, H., Marks, M. E., Harley, V., Mishina, Y., and Kingsley, D. M. (2004). BMP receptor signalling is required for postnatal maintenance of articular cartilage. PLoS Biol. 2, 1815–1827. Ryan, M. C., and Sandell, L. J. (1990). DiVerential expression of a cysteine‐rich domain in the amino‐terminal propeptide of type II (cartilage) procollagen by alternative splicing of mRNA. J. Biol. Chem. 265, 10334–10339. Salter, D. M. (1993). Tenascin is increased in cartilage and synovium from arthritic knees. Br. J. Rheumatol. 32, 780–786. Sanz‐Ezquerro, J. J., and Tickle, C. (2003). Fgf signaling controls the number of phalanges and tip formation in developing digits. Curr. Biol. 13, 1830–1836. Serra, R., Johnson, M., FilvaroV, E. H., LaBorde, J., Sheehan, D. M., Derynck, R., and Moses, H. L. (1997). Expression of a truncated, kinase‐defective TGF‐beta type II receptor in mouse skeletal tissue promotes terminal chondrocyte diVerentiation and osteoarthritis. J. Cell Biol. 139, 541–552. Settle, S., Marker, P., Gurley, K., Sinha, A., Thacker, A., Wang, Y., Higgins, K., Cunha, G., and Kingsley, D. M. (2001). The BMP family member Gdf7 is required for seminal vesicle growth, branching morphogenesis, and cytodiVerentiation. Dev. Biol. 234, 138–150. Settle, S. H., Jr., Rountree, R. B., Sinha, A., Thacker, A., Higgins, K., and Kingsley, D. M. (2003). Multiple joint and skeletal patterning defects caused by single and double mutations in the mouse Gdf6 and Gdf5 genes. Dev. Biol. 254, 116–130. Shimazu, A., Nah, H. D., Kirsch, T., Koyama, E., Leatherman, J. L., Golden, E. B., Kosher, R. A., and Pacifici, M. (1996). Syndecan‐3 and the control of chondrocyte proliferation during endochondral ossification. Exp. Cell Res. 229, 126–136. Spicer, A. P., and McDonald, J. A. (1998). Characterization and molecular evolution of a vertebrate hyaluronan synthase gene family. J. Biol. Chem. 273, 1923–1932. St‐Jacques, B., Hammerschmidt, M., and McMahon, A. P. (1999). Indian hedgehog signaling regulates proliferation and diVerentiation of chondrocytes and is essential for bone formation. Genes Dev. 13, 2072–2086. Storm, E. E., and Kingsley, D. M. (1996). Joint patterning defects caused by single and double mutations in members of the bone morphogenetic protein (BMP) family. Development 122, 3969–3979. Storm, E. E., and Kingsley, D. M. (1999). GDF5 coordinates bone and joint formation during digit development. Dev. Biol. 209, 11–27. Storm, E. E., Huynh, T. V., Copeland, N. G., Jenkins, N. A., Kingsley, D. M., and Lee, S. J. (1994). Limb alterations in brachypodism mice due to mutations in a new member of the TGF beta‐superfamily. Nature 368, 639–643.

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Development of a Sexually Differentiated Behavior and Its Underlying CNS Arousal Functions Lee‐Ming Kow,*,1 Cristina Florea,*,1 Marlene Schwanzel‐Fukuda,*,1 Nino Devidze,* Hosein Kami Kia,* Anna Lee,* Jin Zhou,* David MacLaughlin,{ Patricia Donahoe,{ and Donald Pfaff * *Laboratory of Neurobiology and Behavior, The Rockefeller University New York, New York 10021 { Department of Pediatric Surgery, Massachusetts General Hospital Harvard Medical School, Boston, Massachusetts 02114

I. Introduction II. Sexual DiVerentiation of Brain Mechanisms Producing Lordosis Behavior A. Mechanisms III. Development of Brain Mechanisms Underlying Arousal IV. Outlook for New Work on Sexually DiVerentiated Behaviors References

This chapter addresses questions regarding lordosis behavior, the most extremely sexually diVerentiated behavior that has been analyzed for its neural and molecular mechanisms. Analysis of this behavior has proved for the first time that specific biochemical reactions in specific nerve cell groups in the brain determine a mammalian behavior. Lordosis is done by the female but not by the male. How did the process of sexual diVerentiation occur? A large literature implicates high levels of testosterone during a critical period during development as being responsible for the defeminization of the brain. A new idea, however, oVers the possibility of direct genetic influences independent of testosterone levels themselves. We propose here that Mullerian Inhibiting Substance (MIS) and its receptors could constitute an example of a nonandrogenic genetic influence. Further, specific sexual behaviors depend on underlying arousal states in the central nervous system (CNS). We have proposed the concept of generalized CNS arousal and provide information as to how generalized arousal forces interact with specifically sexual influences, thus to facilitate sexually diVerentiated mating behaviors. ß 2007, Elsevier Inc.

1

These authors made equal contributions to this chapter.

Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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0070-2153/07 $35.00 DOI: 10.1016/S0070-2153(06)79002-0

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I. Introduction The most prominent, biologically crucial, strongly sexually diVerentiated behavior in mammals is lordosis behavior (Fig. 1), the vertebral dorsiflexion response by which female quadrupeds permit reproduction. Governed by a limbic‐hypothalamic system of estrogen‐binding neurons (PfaV, 1968) (Fig. 2), a system proved to be universal among mammals (Morrell and PfaV, 1978). Lordosis depends on a spino‐midbrain‐spinal neural circuit (Fig. 3) that explains the physiology of the behaviorally adequate sensory stimulation and the production of the motor response of lordosis itself (PfaV, 1999). Later immunocytochemical analyses (Shughrue et al., 1997) separated the contributions of two diVerent gene products, those due to the estrogen receptor (ER)‐ gene from those due to the ER‐ gene. In the ventromedial hypothalamus, a cell group containing neurons crucial for lordosis behavior, the large number of ER‐‐expressing neurons far exceeded those expressing ER‐ . Because ERs are ligand‐activated transcription factors, estrogen actions in the hypothalamic neurons that govern lordosis could be used to generate a list of genes (PfaV, 1999) that have two properties: (1) that their transcript levels are elevated by estrogens and (2) that their gene products foster lordosis behavior (Fig. 4). Looking at Fig. 4, it is easy to see that these genes encode many diVerent kinds of biochemicals: growth‐related products, another transcription factor (the progesterone receptor), neurotransmitter receptors, neuropeptides, and their receptors. Nevertheless, Jessica Mong and I (2004)

Figure 1 Drawing of lordosis behavior (bottom panel) as performed by female rats. In all female quadrupeds, the vertebral dorsiflexion by the female permits fertilization by the male and thus controls reproduction. (Adapted from Becker et al., 1992.)

39

2. Development of a Sexually DiVerentiated Behavior h

cc caud

ic

ac

lot

ot tub lpoa aa lsep

h

m co Amygdala

nst

lh

h

pf cg

ob cc

sc fr

cbllm

ac mt tub

db

mpoa oc aha

vm

pvm dm

ic

scp

mamm vpm arc

Figure 2 Drawing of two sagittal sections through the rat brain looking at it from the left side. Black dots indicate locations of cells expressing ERs as proved by the binding and retention of tritiated estradiol. The limbic‐hypothalamic system revealed in this experiment with rat brain turned out to be universal among vertebrate brains. [Adapted from PfaV and Keiner (1973), with abbreviations therein.]

have argued that the list of genes so far discovered—with more on the way by virtue of microarray studies—encodes products that participate in functional modules assuring the biological adaptiveness of reproductive behaviors. Overall, the lordosis behavior neural circuit—the first worked out for a mammalian behavior—and the corresponding molecular mechanisms proved that specific biochemical reactions in specific parts of the mammalian brain produce a specific, biologically important behavior. Female mammals perform lordosis behavior but males do not (Beach, 1948). The first half of this chapter presents a new approach to the molecular mechanisms by which the sexual diVerentiation of this behavior is produced. Underlying all biologically regulated behaviors is a neural function called generalized central nervous system (CNS) arousal (PfaV, 2006; PfaV et al., 2005).

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Estradiol Midbrain central gray

Medial preoptic Med. ant. hypothal. Ventromedial nucl. hypothal.

Hypothal module

Midbrain reticular form

Midbrain module

Lat. vestib. nucl. Medullary reticular formation

Spinal cord

Lower brainstem module Lateral vestibulosp. and reticulosp. tracts

Dorsal roots LI,L2

Stimuli

L5,L6,S1

Spinal module

Pre Re ssure ce pto rs

Flanks Skin of rump tailbase perineum

Lat. longissimus and transverso spinalis

Lordosis response

Figure 3 Neural circuit for the production of lordosis behavior. A spino‐midbrain‐spinal circuit is regulated by estrogen‐modulated neurons in the ventromedial nucleus of the hypothalamus. [Adapted from PfaV (1980).]

Its operational definition states that a more aroused animal or human being is more responsive to stimuli in all sensory modalities, emits more voluntary motor activity, and is more reactive emotionally. Responsible for the activation of behavior, generalized CNS arousal mechanisms now can be explored with biophysical and molecular techniques. In the second half of this chapter, we review two new sets of studies that address development of arousal mechanisms that drive sexual arousal signaled by medial hypothalamic neurons that in turn control lordosis behavior.

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2. Development of a Sexually DiVerentiated Behavior Gene turned on (in hypothalamus) rRNA and growth

Progesterone receptor nNitric oxide synthase ERa E

Adrenergic a 1 receptor Binds ERb

Muscarinic receptors

Female reproductive behaviors

Enkephalin ⫻ opioid receptors Oxytocin ⫻ oxytocin receptor (in preoptic area) GnRH ⫻ GnRH receptor Prostaglandin-D synthase ( ) Figure 4 Genes that have two properties: Their transcript levels are elevated by estrogens (E) following binding to an ER; and, secondly, that their products foster female reproductive behaviors. The exception is the expression of prostaglandin D synthase in the preoptic area, which works through an estrogen‐triggered disinhibition. [Adapted from PfaV (1999).]

II. Sexual Differentiation of Brain Mechanisms Producing Lordosis Behavior Over what time course does estrogen‐dependent, sexually diVerentiated lordosis behavior develop? We had to answer this question because, although the pioneering work of Christina Williams and her laboratory at Duke University had shown that lordosis can be performed by very young rat pups, but was suspected to be part of a micturition response rather than having a frankly prereproductive character. Complementary to the Williams et al. results, we wanted to see when it became specific to the female and estrogen dependent. Our results (Kow et al., 2005a,b) showed that lordosis behavior can be induced in female, but not in male rat pups with estrogen treatment, with or without progesterone treatment. And it cannot be induced by progesterone alone. In female rats, but not males, lordosis behavior first appeared on Day 15 after birth and quickly reached and stayed on or near the maximal level from

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Day 16 onward. Lordosis was never observed in females without estrogen treatment nor in males with any treatment, indicating that under these conditions, as in adults, lordosis is estrogen‐dependent and that there is a gender diVerence, respectively. These behavioral developments were not accompanied by age or gender diVerences in the expression of ER‐. However, there were large gender differences in the density of progesterone receptor (PR)‐ir cells in the VMN. Females but not males expressed PR. This resonates with the strongly sexually diVerentiated mechanisms illustrated below at several molecular chemical levels. All of these sex diVerences in behavior and PR expression have their onset long after the arrival of the neuroendocrine cells essential for the ovulatory surge of luteinizing hormone—LHRH (GnRH) cells that have migrated from the olfactory placode into the brain (Schwanzel‐Fukuda and PfaV, 1989). The eventual behavioral importance of these cells is that they are required for directing the anterior pituitary gland to command the testes to produce testosterone, thus to support normal sexual arousal in experimental animals (PfaV, 1973; Wu et al., 2006) and in men (Schwanzel‐Fukuda et al., 1989). The dual roles—neuroendocrine and behavioral—of these LHRH (GnRH) neurons serve to synchronize lordosis behavior with ovulation in the female, thus rendering behavior appropriate to the preparations for reproduction in the rest of the female’s body.

A. Mechanisms In light of the strong line of data, mentioned above, historically implicating testosterone administration in the defeminization of behavior, the sexual diVerentiation of the mammalian CNS has tended to be assigned solely to the actions of androgenic hormones and their metabolites (Beach, 1948; Phoenix et al., 1959) (Fig. 5). Administration of testosterone to a female rat during the first day after her birth can completely defeminize her behavior; and correspondingly, castrating the male during first day after his birth, thus depriving him of testicular androgens can completely feminize his behavior (PfaV and Zigmond, 1971). In fact, the emphasis on neonatal testosterone gains strength from the sex diVerences in molecular mechanisms in the brain that support the sexual diVerentiation of sexual behavior. In Fig. 6, I briefly picture some of the steps known to be involved in hypothalamic neurons necessary for the ability of progesterone, acting through its PR, to amplify estrogens’ eVects on lordosis behavior in females but not males. This is an especially interesting train of neurochemical events because PR is in turn a transcription factor. From the early work of Christopher Krebs in my laboratory using the diVerential

43

2. Development of a Sexually DiVerentiated Behavior XX

Normal female

XY

Female neonatal testosterone

Neonatal castration

Normal male

High

Female sex behaviors

Low

High

Ovulatory surge, LH

Low

Low

Anxiety (cage emergence, elev. plus, crowding, chronic stress)

High

Figure 5 A huge literature supports the idea that the presence of high levels of testosterone or testosterone metabolites during a critical period in the development of a mammalian brain will defeminize certain aspects of brain function and masculinize others. Summarized in this figure are the strongest and most obvious examples: the ability of the female brain to produce lordosis behavior and to govern an ovulatory surge of luteinizing hormone. Other behavioral examples would include responses related to anxiety and aggression.

display technique we know that at least a heat‐shock cognate protein 73, a secretory carrier membrane protein SCAMP 4, and a membrane protein DX 25 that can bind progesterone are all three modulated by progesterone treatment. We need to work more, using microarrays, to discern the full menu of changes in the hypothalamic transcriptome induced by progesterone treatment. Likewise, in Fig. 7, the molecular train of events involved in the action of the opioid peptide enkephalin on lordosis behavior is sketched briefly. Some of these are known to be deficient in males. Another neuropeptide system likely involved in gender‐dependent neurotransmission is oxytocin, whose receptor is heavily expressed in the very same neurons as ER‐ and ER‐ , but with a diVerent ratio of  to according to whether the animal is a male or a female (Devidze et al., 2005). For all neuropeptides aVecting lordosis behavior, signal transduction pathways, some of whose elements are listed in Table I are likely estrogen dependent in the female and likely modulated diVerently in the male. For example, patterns of coexpression of genes coding diVerent protein kinase C (PKC) isoforms are significantly diVerent between females and males (Devidze et al., 2005).

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Kow et al. P

Estrogen priming

In

’s not

’s2

Behaviors blocked by PR antisense DNA1 or RU4862

E induces PR mRNA3 In ’s not ’s4 and P binding3a through 5 PR gene promoter by transcription 6 Induces PR knockout blocks8 PR-B mRNA7 P needs new synthesis9 DNA binding10a P induces new stronger in ’s mRNAs10

Binds to PR

Inducing new genes

IN VM Hypothalamus

Has PR11, PRB form mRNA12 E induced13 Correlated with behavior14

E induced in ’s not ’s15

Lordosis behavior16 Figure 6 A large number of molecular functions reveal sex diVerences in the manner in which progesterone (P), having bound to PR, is able to influence behavioral function. Normal molecular chemical steps seen in the female brain are depicted. Some of the strongest sex diVerences are listed on the right. [Adapted from PfaV (1999); and the superscripted references therein.]

1. A New Thought In all of these studies, behavioral, endocrine, and molecular, it has been assumed that sex diVerences are due exclusively to the actions of testosterone during a neonatal critical period. However, several years ago, Arthur Arnold, at the University of California, Los Angeles, put forward a new idea (Arnold, 1996, 1997). He entertained the possibility that gene products deriving from the Y chromosome or from diVerence in the inactivation of the X chromosome could aVect sexual diVerentiation of brain and behavior independent of the eVects of androgenic hormones themselves. If any gene could be shown to do so, Arnold’s idea would lead to a paradigm shift in reproductive biology. The first gene product that caught our attention is in the cascade responsible for diVerentiation of the gonadal ridge. We tested the hypothesis that this gene also participates in CNS development. We asked whether the gene coding for a specific receptor of such a gonadal ridge principle, namely Mullerian Inhibiting Substance (MIS, also called anti‐Mullerian hormone),

45

2. Development of a Sexually DiVerentiated Behavior Enkephalin

Transcribed in VM Hypothalamus

Induced by

E1

In

’s

not

’s2

E ENK mRNA 3 through ENK DNA binding ’s > ’s8 promoter binding4 causing mRNA ’s >> ’s9 5 transcription in VMN specifically6. Opioid d receptor too7.

Occurs rapidly, before lordosis10 TR blocks ENK and lordosis 11,12 Antisense DNA reduces lordosis13

Correlated with lordosis14

Fosters lordosis Figure 7 Certain molecular functions reveal sex diVerences in the manner in which the opioid peptide enkephalin, having bound to  or  opioid receptors, can influence behavioral function. Normal molecular chemical steps seen in the female brain are depicted. Some of the strongest sex diVerences are listed on the right. [Adapted from PfaV (1999); and the superscripted references therein.]

might be expressed in the developing mouse brain. If such receptors were expressed in the brain, it would give concrete evidence for Arnold’s idea of biochemical substances other than testicular androgens aVecting sexual diVerentiation of the brain. MIS is the male gonadal product which mediates a paracrine regression of the embryonic Mullerian duct during a specific period of embryogenesis critical for normal sexual diVerentiation of the internal reproductive structures. It is the ligand of heterodimeric receptors consisting of type I and type II components. For the reasons introduced above, we worked to determine the distribution of mRNA encoding the MIS type II receptor first in the 14‐day‐old mouse embryo by using in situ hybridization and RT‐PCR techniques using oligonucleotide probes and primers designed from the rat MIS type II receptor sequence. We demonstrated the expression of the gene for

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Table I Comparisons of EVects of Various Neurotransmitters and Neuropeptides on Lordosis Behavior, Neuronal Electrical Activity, and Second Messengersa EVects on Agents (Type of Receptor)

Lordosis Behavior

Neuronal Activity

Membrane Enzymes

Lordosis‐facilitating agents Acetylcholine (M3?) Norepinephrine (1) Serotonin (5‐HT2) LHRH PRL Oxytocin Substance P (NK1) GABAA

" " " " " " " "

" " "

" " #

" " " " " " "

on on on on on on on

Lordosis‐inhibiting agents Serotonin (5‐HT1A) Dopamine (D1?D2?) CRF ‐Endorphin Neuropeptide Y Cholecystokinin Glutamate

# # # # # # #

# "or# ? # #? " "

# " " # # "

on AC (D1) or # (D2) on AC on AC on AC on AC on AC, PLC

PLC PLC, PLA2, PLD PLC PLC, PLA2 PLC PLC PLC

a

Adapted from Kow et al. (1994).

MIS type II receptor in the brain and anterior pituitary gland of 14‐day‐old embryonic mouse. We chose this day for the first extensive series of investigations because it is after the onset of expression of MIS, in males, the development of circulation, and the diVerentiation of the gonadal ridge. It is also after the arrival of GnRH neurons in the basal forebrain—GnRH neurons which had migrated into the brain from the olfactory placode (Schwanzel‐ Fukuda and PfaV, 1989), an event essential for normal sexual arousal in males (Schwanzel‐Fukuda et al., 1989). In contrast, Day 14 precedes the closure of the blood–brain barrier. Briefly, we found that MIS type II receptor mRNA was highly expressed in the CNS of the 14‐day‐old embryos, both in the brain and the anterior pituitary gland (Fig. 8). The roof of neopallial cortex of the telencephalon was strongly positive. Additionally, sections of the cranial region showed strong expression in thalamus and hypothalamus, medial septal, and preoptic areas, as well as the roof and the ventral part of the mesencephalon. Furthermore, the presence of MIS type II receptor transcripts was detected in the pons and intraventricular portion of the cerebellar primordium (metencephalon), medulla (myelencephalon), choroid plexus in the roof of the fourth ventricle, and the olfactory epithelium/bulb. Notably, the highest

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Figure 8 Whole‐mount in situ hybridization of mouse 14‐day‐old embryos with DIG‐labeled antisense (A–C, E, H) riboprobes for MIS type II receptor. Sections represented are cortex (A), thalamus (B), hypothalamus (C), anterior pituitary gland (E), and heart (H). No signals above the background level can be seen in the embryos hybridized with the corresponding sense probes (F, negative control), those pretreated with RNAase prior to hybridization with antisense (G, negative control), or those without probes (D, negative control).

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hybridization signal, other than the gonad, was expressed in the anterior pituitary gland. However, no signal was present in spinal and cranial ganglia or in the spinal cord. Analysis of MIS type II mRNA expression in sagittal sections showed strong signals in the lungs, the primordium of the mesonephros/gonads, and the liver. The heart and stomach were negative for MIS type II receptor mRNA. Additional observations were carried out at other developmental stages (Fig. 9). Briefly, in 16‐day embryos much lower levels of expression were seen in the neopallial cortex, diencephalon, cerebellum, and olfactory epithelium/ bulb, with higher levels expressed in the anterior pituitary gland. In 18‐day embryos, very faint signals were persistent only in the frontal cortex, the olfactory mucosa, cerebellum, and diencephalon. Robust and discrete signals were seen in the anterior pituitary gland. On postnatal day 0 (P0), signals were localized in the frontal cortex, cingulate and subcallosal gyri, diencephalon, cerebellum, and very strong ones were seen in the olfactory epithelium and anterior pituitary gland. Surprisingly, no obvious sex diVerences in the patterns or levels of expression were observed throughout gestation or on P0 in these mice. Overall, these results oVer the possibility that MIS is acting to foster sexual diVerentiation or other developmental phenomena in the embryonic mouse brain. The means by which it does so remain to be elucidated. Looking back on this work, the rationale for the result is obvious. A basic tenet of sexual development in mammals is that genetic sex, determined by the presence or absence of a Y chromosome, directs the primordial gonad to diVerentiate into either testis or ovary. Products secreted by the testis (testosterone and MIS) then direct the developmental program leading to male sexual diVerentiation (Josso et al., 2001; Jost, 1953; Teixeira and Donahoe, 1996). In the absence of these hormones, the resulting developmental pathway is female. While it has been held that sexual diVerences of the brain (in a manner similar to secondary sexual diVerentiation of other tissues) are mediated entirely by the action of gonadal steroids during a critical period of embryonic development, which occurs after gonadal diVerentiation (Breedlove et al., 1999), more recent experimental and clinical data of Kolbinger et al. (1991) and Arnold (1997) have cast doubt on the general validity of this classical steroid hormone hypothesis and suggest that other intervening factors and mechanisms may also be involved (Floody and Arnold, 1997; Maxson, 1996). MIS is a member of a large TGF‐ superfamily of growth and diVerentiation factors whose members play important roles in development and feedback control of the hypothalamic‐pituitary‐gonadal hormone axis as well as in the regulation of neural and bone morphogenesis (Cate et al., 1986; Massague, 1998). A growth‐inhibitory gonadal hormone, MIS, is secreted by the Sertoli cells of fetal and adult testis at high levels until puberty when MIS concentration decreases to lower levels which persist throughout male adult

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Figure 9 Whole‐mount in situ hybridization of mouse 12‐ and 13‐day‐old embryos as well as PO, with DIG‐labeled antisense (A, C, E, G) riboprobes for MIS type II receptor. Sections represented are 12‐day cortex (A), 13‐day thalamus (C), PO cortex (E), and PO anterior pituitary gland (G). No signals above the background level can be seen in the embryos hybridized with the corresponding sense probes (F, negative control), those pretreated with RNAase prior to hybridization with antisense (B, H), or those without probes (D).

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life (Baker et al., 1990; Hudson et al., 1990; Josso et al., 1990). MIS is unmeasurable in serum until puberty in females and its expression was originally detected in the antral follicles of the adolescent ovary and falls to undetectable levels at menopause (Bezard et al., 1987; Kuroda et al., 1991; Takahashi et al., 1986; Teng, 1987). A more recent study showed the presence of MIS protein in the ovary at the end of gestation (Rajpert‐De Meyts et al., 1999). MIS is the ligand of a heterodimeric receptor molecule that is analogous to the TGF‐ , activin, and BMP receptors each a serine/threonine kinase (Baarends et al., 1994; Clarke et al., 2001; di Clemente et al., 1994; Teixeira et al., 1996; Visser et al., 2001). On the basis of nomenclature established for other TGF‐ receptors, this heterodimeric pair consists of a type I (50–60 kDA) and type II (70–80 kDA) receptors. Both receptors posses an extracellular ligand‐binding domain, with conserved cysteine residues, a single hydrophobic transmembrane domain, and an intracellular serine/ threonine kinase domain, deletion of which results in an inability to eVect multiple ligand‐mediated signaling responses (Clarke et al., 2001). The extracellular domain of the MIS type II receptor specifically binds the bioactive C‐terminus of MIS, allowing it to dimerize with type I receptor which in turn is responsible for cellular signal transduction (Clarke et al., 2001; Visser et al., 2001). This signaling includes interaction with a family of proteins called Smads described earlier for other TGF‐ family members (Massague, 1998). MIS type I and type II receptors are expressed in the Mullerian ducts of both male and female urogenital ridges (Baarends et al., 1994; Clarke et al., 2001; di Clemente et al., 1994; Teixeira et al., 1996; Visser et al., 2001) and granulosa (Baarends et al., 1994, 1995a; di Clemente et al., 1994), Sertoli (Baarends et al., 1994, 1995b), and Leydig cells (Baarends et al., 1995b; Lee et al., 1999; Racine et al., 1998; Teixeira et al., 1999) of both embryonic and adult gonads. With all of this knowledge in hand, we now know that, in fact, the MIS receptor type II is expressed in both males and females and its expression is widespread and robust. On the basis of receptor expression patterns in the urogenital ridge, we predicted no sexually dimorphic expression of the receptor. The importance of this work for validating the new approach espoused by Arnold (1997) is still not clear, but we are optimistic for the following reasons. MIS (or anti‐Mullerian hormone), a testicular glycoprotein hormone which exhibits a sexually dimorphic pattern of expression, causes regression of the Mullerian ducts (the anlagen of the female internal reproductive structures: uterus, fallopian tubes, and upper third of the vagina) during male embryogenesis (Josso et al., 2001; Teixeira et al., 1996). Originally defined only for its role in the regression of the Mullerian duct, a large number of extra‐Mullerian functions have subsequently been described for MIS, such as control of germ cell maturation and gonadal morphogenesis

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(Hunter, 1996; Ingraham et al., 2000; Lee and Donahoe, 1993), gonadal ridge development (Haqq et al., 1994), inhibition of fetal lung maturation (Catlin et al., 1988, 1990, 1997), and growth suppression of transformed cells (Chin et al., 1991; Masiakos et al., 1999, Segev et al., 2000). It is clear now that the role played by this hormone is not as simple as previously thought and that correct temporal and quantitative expression of MIS and its receptor(s) may be critical for some normal developmental and physiological functions of the body. Molecular and biochemical studies have shown that MIS mediates its signals through type I and type II serine/threonine kinase receptors which are both required to mediate functional signaling responses (Clarke et al., 2001; Visser et al., 2001). MIS binds first to the type II receptor which recruits and trans‐phosphorylates with the type I receptor and which subsequently initiates downstream cellular signaling. Developmental expression of MIS type II receptor in the gonads and urogenital ridges of both sexes has shown high levels at the time of Mullerian duct regression, much lower or even absent levels perinatally and prepubertally, increased levels again (reexpression) with sexual maturation at puberty, and continuance at these high levels of expression in adults (Baarends et al., 1994; di Clemente et al., 1994; Teixeira et al., 1996). Although the expression of the MIS type II receptor is essential for normal gonadal morphogenesis and sexual diVerentiation, the role(s) of MIS signal transduction in the adult gonad has yet to be determined, except for work done in Leydig cells (Josso et al., 2001; Teixeira et al., 1996). Looking back at these results, we comment first on the appropriateness of the methodology and then on possible functional interpretations. Several approaches to in situ hybridization histochemistry were used to determine the distribution of cells that express mRNAs encoding MIS type II receptor in the 14‐day‐old mouse embryonic development. Taking into consideration the high stringency conditions used in the in situ hybridization protocol, confirmed by lower resolution film autoradiography, we believe that the antisense probes employed specifically detected MIS type II receptor transcripts. These were detected not only in the brain, but also in the anterior pituitary gland, suggesting a wider distribution than predicted by the already known functions of MIS. We chose for the initial experiments a 14‐day‐old mouse embryo because the expression of MIS type II receptor had been studied by in situ hybridization (Baarends et al., 1994; di Clemente et al., 1994; Teixeira et al., 1996) during similar times in the gonads and urogenital ridges during the period of active Mullerian duct regression. MIS type II receptor message was localized in the roof of neopallial cortex (future cerebral cortex) where MIS can influence the number, size, and connectivity of neurons in the cortical areas where MIS might help to prevent unregulated expansion of the cortical cell population to achieve a normal

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progression of development. Apoptosis, which is induced by the MIS hormone in the Mullerian duct as well as the lung (Catlin et al., 1997), also plays an essential role during development, causing tissue sculpturing during morphogenesis and the maturation of neuronal circuitry in the CNS. Negative growth regulators like MIS have increasingly been recognized as having critical roles in balancing cellular proliferation and death. MIS type II receptor was also found in the hypothalamus, a key brain structure involved in basic functions such as sexual activity, sleep, food intake, temperature regulation, and motivation, emotion, and autonomic responses. Importantly, the hypothalamus secretes hormones that stimulate or inhibit secretions of the anterior pituitary gland. In septal, hypothalamic, and preoptic areas where sexually dimorphic structures have roles in regulation of reproductive behavior (PfaV, 1999; PfaV et al., 2002), MIS type II receptor signals were abundant. MIS type II receptor message was additionally present in the olfactory epithelium and bulb. Although MIS has not been previously linked to olfaction, it is tempting to speculate that MIS action on the olfactory epithelium/ bulb may be involved in the circuitry between olfaction, copulatory/reproductive behavior, and gonadotropin secretion (McClintock and Newman, 2002). Expression of MIS type II receptor in the anterior pituitary gland (hypophysis cerebri) is of particular interest since this gland aVects the function of several endocrine systems. These results mandate careful study particularly of gonadotropes in the anterior lobe, during and after organogenesis, to explore a potential role for MIS in feedback regulation or other mechanisms (Knobil and Neill, 1994). The anterior pituitary gland displayed the highest level of expression compared to the CNS structures analyzed at these stages of embryological and neonatal development, strongly suggesting a role for this inhibitory hormone in modulating the hypothalamo‐pituitary‐gonadal neuroendocrinological activity, and possibly the expression of the multiple genes that constitute a gonadotrophic phenotype. MIS type II receptor message was also found in the developing hypothalamus, a region known to play a central role in the integration of endocrine, autonomic and behavioral cues essential for reproduction (PfaV et al., 2002), and in the preoptic area where there are sexually dimorphic structures with suspected roles in sexual orientation. MIS type II receptor transcripts were also found in the olfactory epithelium and olfactory bulb. The importance of olfactory cues to copulatory behavior and gonadotropin secretion (PfaV et al., 2002) has been established. These findings tempt us to speculate that MIS could have a role in male sexual diVerentiation of the brain. It remains for future work to delineate potential roles of MIS in brain development and diVerentiation. Does the MIS reach the brain in the genetic male? Certainly MIS is readily measured in serum following secretion from

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the gonads (Baker et al., 1990; Hudson et al., 1990; Josso et al., 1990). Furthermore, MIS administered exogenously is clearly detectable in the serum (Stephen et al., 2001, 2002). Thus, MIS readily distributes in at least some body compartments. The exact timing of blood–brain barrier development in the mouse remains controversial, but it appears not to be closed before 15 days of embryonic development (Bauer and Bauer, 2000; Risau et al., 1986), thus allowing the possibility of circulating MIS to encounter the cerebral MIS receptors reported here and thereby influence basal forebrain diVerentiation. The answers to these questions will help us to define novel mechanisms involved in brain development and sexual diVerentiation (Giedd et al., 1997; Gorski, 1985; Simerly, 1998). It is possible that sexual dimorphism will reside in the presence of the ligand and the MIS type I receptor and Smads, which appear to be diVerentially regulated in the male Mullerian duct and also induced by the ligand and MIS (unpublished data). It remains to compare expression of the type I receptor and Smad 1 and 8 in the brain of males and females to determine if such expression is enhanced by MIS. In any case, our study demonstrates a widespread and enduring pattern of expression for MIS type II receptor in the embryonic and neonatal mouse brain and in the anterior pituitary gland. Certainly, large numbers of genes on the sex chromosomes deserve investigation of the sort we have started with the MIS type II receptor. It seems likely that X chromosome and Y chromosome genes influence brain phenotype in a sex‐specific manner (Arnold and Burgoyne, 2004), but much more work on this subject is needed.

III. Development of Brain Mechanisms Underlying Arousal A much newer subject investigated in the laboratory deals with mechanisms of CNS arousal that underlie all sexually diVerentiated behaviors. Even before they can walk, the laboratory animals we study can respond strongly to olfactory, cutaneous, auditory, and vestibular stimuli. As early as 5 days of age, mice can show a righting reflex and move vigorously by crawling. Thus, it is possible to use this genetically tractable species to study the development of a brain function that underlies sexual arousal and sexually diVerentiated behavior, a function we have named ‘‘generalized arousal’’ of the CNS (PfaV, 2006). Particularly interesting will be how changes in generalized arousal impact specifically sexual arousal that in turn is required for the performance of sexually diVerentiated lordosis behavior. We began with a precisely worded and logically complete operational definition of generalized arousal (PfaV, 2006) mentioned briefly above. This operational definition states that a more aroused animal is more responsive to stimuli in all sensory modalities; emits more voluntary motor activity, and is more reactive emotionally.

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Importantly, we then devised a laboratory assay that measures all three components of this definition and yields quantitative, physical measurements of the behavior of mice (Garey et al., 2003) showing, for example, that female mice are more sensitive to histamine H1 receptor antagonism than males (Easton et al., 2004; Shelley et al., submitted for publication). Biophysical and molecular mechanisms for the development of generalized arousal are under investigation in our laboratory. The manner in which specific arousal forces, such as sexual arousal, combine with generalized arousal to determine the behavior of mice at any age is expressed in the following equation: dA ¼ Fg ðAg Þ þ F1 ðAS1 Þ þ F2 ðAS2 Þ þ    þ Fn ðASn Þ where A is the state of arousal of the CNS at any moment, Ag is generalized arousal, and ASn is a specific form of arousal. How does a specifically sexual influence on arousal, such as a steroid sex hormone, interact with a generalized arousal neurotransmitter to alter the electrical activity of a lordosis‐related neuron. Kow et al. (2005a,b), using electrophysiological recording from the ventromedial nucleus of the hypothalamus in young animals, have been able to show that estrogen administration can potentiate neuronal responses to the arousal‐related transmitter histamine as well as to NMDA (Fig. 10). These results give one example of how a generalized arousal force (histamine) can combine with a specifically sexual (hormonal) influence to determine the activity of a (hypothalamic) neuron that governs a sexually diVerentiated behavior.

IV. Outlook for New Work on Sexually Differentiated Behaviors Analyses of the sexually diVerentiated lordosis behavior showed for the first time that specific biochemical reactions in a specific nerve cell group in a mammalian brain regulate a specific behavior. The classical theory of sexual diVerentiation of the CNS states that testosterone and its metabolites circulating at high levels during a critical developmental period defeminize the brain. Our new microarray results showing male/female diVerences in transcript levels of a large number of genes 24 hours after birth are in line with this classical view, but we have no information yet as to which transcripts are most important or how they exert their behavioral eVects. New work showing the eVects of testosterone on the neonatal transcriptome will help to follow up the astonishing number of genes representing several biochemical systems that appear to be sexually diVerentiated. To date, a reasonable candidate for the new idea of direct nonandrogenic genetic eVects on sexual diVerentiation of the brain is represented by the MIS system and its receptors.

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2. Development of a Sexually DiVerentiated Behavior HA receptors: G-protein−coupled receptors NMDA receptors: Ligand-gated ion channels

Firing rate (spikes/s)

A 8 NMDA HA

n = 4/19

0 12

E2 10 nM B

HA

NMDA

0 12 3

C

n = 5/19

E2 10 nM HA

NMDA n = 2/19

E2 10 nM 5 min The independence of the potentiation of the HA and NMDA excitations suggests that acute E2 employs multiple mechanisms Figure 10 Interactions between generalized arousal transmitters and a specifically sexual arousal‐related hormone as measured by electrophysiological recording. Acute E can potentiate excitatory response caused by NMDA alone (A), by both NMDA and HA (B), or by HA alone (C). The conventions for this figure are the same as in Fig. 2. Excitatory responses to NMDA (thin arrows) and HA (thick arrows) were recorded from three VMN neurons (A–C). In neuron A, NMDA excitation was potentiated to 188% of control by E2, but HA excitation was not. In neuron B, the potentiation of NMDA by E2 was so strong that toward the end of the E2 infusion there was a depolarization block (the excitation was cut short and the firing rate decreased below the pre‐NMDA level) during the second NMDA excitation. Otherwise, the excitation would be much greater. Such a potentiation was not due to sensitization by the preceding HA, because in neuron C, the HA response was potentiated to 274% of control, but the following NMDA excitation was not. Note that the order of transmitter administration was diVerent among A, and B, or C. (Adapted from Kow et al., 2005a,b.)

Again, we do not yet know exactly how MIS would work in the developing brain, if it does. A deep concept underlying all sexual behaviors is that of ‘‘generalized arousal’’ of the CNS. Its existence has been indicated so far by the mathematical statistics of principal components analysis and by the fact that we already know some of its mechanisms. But how does it work, and, in particular, how does it fuel the CNS so that sexually diVerentiated behaviors are performed in a biologically adaptive manner? EVects of arousal‐related transmitters on hypothalamic neuronal electrical activity have been demonstrated, but we still must work out the detailed signal transduction pathways involved, as well as the ion channels that are appropriately opened or closed. Ultimately, the appearance of any sexually diVerentiated behavior depends on this most fundamental brain function.

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Phosphodiesterases Regulate Airway Smooth Muscle Function in Health and Disease Vera P. Krymskaya and Reynold A. Panettieri, Jr. Department of Medicine, Pulmonary, Allergy, and Critical Care Division University of Pennsylvania, Philadelphia, Pennsylvania 19104

I. Introduction II. Developmental and Physiological Responses Regulated by PDEs III. PDEs and Airway Smooth Muscle Function A. PDE and ASM Tone and Contractility B. PDE and ASM Growth and Migration IV. Cytokine and Chemokine Secretion by ASM: A Role for PDEs V. Conclusions Acknowledgments References

On the basis of structure, regulation, and kinetic properties, phosphodiesterases (PDEs) represent a superfamily of enzymes divided into 11 subfamilies that catalyze cytosolic levels of 30 ,50 ‐cyclic adenosine monophosphate (cAMP) or 30 ,50 ‐cyclic guanosine monophosphate (cGMP) to 50 ‐AMP or 50 ‐GMP, respectively. PDE4 represents the major PDE expressed in inflammatory cells as well as airway smooth muscle (ASM), and selective PDE4 inhibitors provide a broad spectrum of anti‐inflammatory eVects such as abrogating cytokine and chemokine release from inflammatory cells and inhibiting inflammatory cell traYcking. Due to cell‐ and tissue‐specific gene expression and regulation, PDEs modulate unique organ‐based functions. New tools or compounds that selectively inhibit PDE subfamilies and genetically engineered mice deficient in selective isoforms have greatly enhanced our understanding of PDE function in airway inflammation and resident cell function. This chapter will focus on recent advances in our understanding of the role of PDE in regulating ASM function. ß 2007, Elsevier Inc.

I. Introduction Phosphodiesterases (PDEs) inhibit cyclic nucleotide‐dependent signaling pathways. The function of PDE as an enzyme degrading 30 ,50 ‐cyclic adenosine monophosphate (cAMP) to 50 ‐AMP was first described in 1958 while studying glycogen metabolism regulated by hormones and neurotransmitters Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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(Sutherland and Rall, 1958). These studies led to five Nobel Prizes and provided the paradigm of second messengers and the signal transduction (Beavo and Brunton, 2002; Houslay, 1998). Stimulation of G‐protein–coupled receptors that activate adenylyl cyclase via the G‐protein subunit, Gs, generates cAMP (Fig. 1). Levels of cAMP with specific cellular components bind to the regulatory subunit of cAMP‐dependent kinase or protein kinase A (PKA) and subsequently activate PKA (Fig. 2). PDEs function as negative regulators of cAMP‐ and 30 ,50 ‐cyclic guanosine monophosphate (cGMP)‐activated signaling pathways. Activated PKA phosphorylates Ser133 of nuclear cAMP response element‐binding protein (CREB), which recruits a coactivator, the CREB‐binding protein (CBP); CBP has intrinsic histone acetyltransferase activity and interacts with RNA polymerase II (Conti et al., 2003; Houslay and Adams, 2003), which enhances transcription of about 105 genes with the cAMP response element (CRE) motif in their promoter regions including among others the glucocorticoid receptor, the cystic fibrosis transmembrane conductance regulator (CFTR) and angiotensinogen (Beavo and Brunton, 2002). In addition to the critical role of cAMP in the activation of PKA and

Albuterol PGE2 Salmeterol Formeterol

TNF-a, IL-1b

Gs Adenylyl cyclase

p38, ERK1/2, JNK NF-kB

cAMP Cilomast rolipram

?

PDE AMP

AP-1

PKA

Nucleus

CRE

Gene

Figure 1 Phosphodiesterases (PDEs) promote degradation of cAMP in ASM cells. Agonist‐ induced stimulation of G‐protein–coupled receptors recruits Gs subunit which activates adenylyl cyclase followed by generation of 30 ,50 ‐cAMP. Increased cAMP levels activate protein kinase A (PKA) which phosphorylates cAMP response element (CRE)‐binding protein and regulates gene transcription. PDE hydrolyzes 30 ,50 ‐cAMP to 50 ‐AMP.

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Adenylyl cyclase PDE4 inhibitor ATP

cAMP[active]

AMP [inactive]

R C

C

R

R

R

Regulatory subunits

PKA[inactive]

Catalytic subunits

C

C

PKA[active]

Proteins

Proteins

Inflammation P

Migration Bronchoconstriction

ATP

ADP Gene expression

Figure 2 Phosphodiesterase (PDE) inhibition abrogates cytokine‐induced eVects in ASM. Increases in cAMP levels promote dissociation of the catalytic subunits of PKA from its regulatory subunits thus rendering PKA active. Activated PKA catalyzes phosphorylation of multiple downstream proteins, which ultimately regulate gene expression, inflammation, migration, and bronchoconstriction. Inhibition of specific PDE isoforms maintains increased levels of cAMP and activation state of cAMP‐dependent cell functions.

transcriptional regulation, there are other nucleotide‐binding proteins whose activities are regulated. cAMP and cGMP bind to the cyclic‐nucleotide‐gated channels including the potassium channel on cardiac pacemaker cells; cAMP can regulate the activities of guanine‐nucleotide exchange factors (GEFs) including Epac (exchange protein directly activated by cAMP) (de Rooij et al., 1998). The activated cAMP‐dependent pathways are inactivated by the hydrolysis of cAMP to 50 ‐AMP eVected by cAMP PDEs, receptor–agonist dissociation, and receptor desensitization. While, at first, investigators focused on determining the precise molecular mechanism regulating cyclic nucleotide production, others recognized that cAMP degradation regulated by PDEs played a more salient role in modulating PKA activity. Interestingly, the maximal rates of cAMP degradation exceed by more than an order of magnitude of the maximal rates of cAMP synthesis (Beavo and Brunton, 2002), suggesting that the degradative pathway may more sensitively regulate cAMP levels than the synthetic pathways (Beavo and Brunton, 2002).

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Classification of PDE superfamily. PDEs represent a superfamily of PDEs including 20 diVerent genes subgrouped into 11 PDE families (Conti et al., 2003; Soderling and Beavo, 2000; Vignola, 2004). The multitude of PDE isoforms, which perform similar biochemical reactions, provide cell and tissue specificity to selectively regulate the amplitude, duration, and localization of cytosolic cAMP. PDEs consist of a modular structure with conserved catalytic domain in the C‐terminal and regulatory domains or motifs in the N‐terminus (Soderling and Beavo, 2000; Vignola, 2004). Each family is characterized by unique enzymatic properties regulated by distinct allosteric activators or inhibitors and possesses specific pharmacological inhibitory profiles. Since comprehensive reviews of PDE isoforms are available (Conti and Jin, 1999; Conti et al., 2003; Essayan, 1999, 2001; Soderling and Beavo, 2000; Vignola, 2004), we will only briefly identify the members of the PDE superfamily and focus specifically on PDE function in ASM. PDE1, or calcium/calmodulin‐dependent PDE, comprises a family of enzymes encoded by three genes (1A, 1B, and 1C), an alternative splicing of which produces multiple PDE1 isoforms with distinct N‐ and C‐terminal regions (Houslay, 1998). PDE1 enzymes are primarily distributed in heart, brain, and lung, and are regulated by a calcium‐binding modulator calmodulin and preferentially hydrolyze cAMP (Essayan, 2001). PDE2, also called cGMP‐stimulated PDE, has a regulatory domain‐binding cGMP with high aYnity, where either membrane‐bound guanylate cyclase is activated by atrial natriuretic factor (ANF), such as kidney cells, or where cytosolic guanylate cyclase is activated by nitric oxide (NO) such as in smooth muscle cells regulated by the endothelium (Houslay, 1998). PDE3, encoded by two genes (3A and 3B), shows a high aYnity to cAMP and is regulated by PKA and insulin (Houslay, 1998). PDE3 remains membrane‐bound and functions in a spatially restricted manner, determining the concentration of local pools of cAMP with primary distribution in heart, lung, liver, platelets, adipose tissue, and immunocytes (Essayan, 2001). PDE4, the largest PDE gene family, is encoded by four genes (4A, 4B, 4C, and 4C) that yield approximately six diVerent splice variants per gene due to alternative mRNA splicing (Houslay, 1998). PDE4 isoforms are cAMP‐ specific, cGMP‐insensitive, and are primarily expressed in the lung, kidney, brain, liver, and immune cells, suggesting a fundamental role in the unique cellular responses on the activation of cAMP signaling cascade where they are expressed (Essayan, 2001). Because of wide distribution of PDE4 isoforms in inflammatory and ASM cells, PDE4 can play a central role in regulating the inflammation associated with asthma and chronic obstructive pulmonary disease (COPD). The cGMP‐specific PDE5 is predominantly expressed in the corpus cavernosum (Burnett, 2005). Because cGMP activates cGMP‐ dependent protein kinase 1, which promotes smooth muscle relaxation permitting intracorporal blood engorgement and pressure increase, PDE5 was

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identified as a therapeutic target to treat erectile dysfunction (Burnett, 2005). PDE6 is also a cGMP‐specific family of enzymes encoded by three genes (6A, 6B, and 6C) with primary distribution in photoreceptors (Essayan, 2001; Vignola, 2004). PDE7, a highly cAMP‐specific isoform encoded by two genes, is expressed primarily in skeletal muscle, heart, kidney, brain, pancreas, and T lymphocytes (Essayan, 2001; Vignola, 2004). PDE8 was first among PDEs identified as insensitive to the nonselective PDE inhibitor isobutyl methyl xanthine (IBMX); PDE8 is also cAMP‐specific and is encoded by two genes (8A and 8B) with predominant distribution in testes, eye, liver, skeletal muscle, heart, kidney, ovary, brain, and T lymphocytes (Soderling and Beavo, 2000). PDE9, like PDE8, is not inhibited by IBMX; however, in contrast to PDE8, PDE9 is cGMP‐specific and predominantly expressed in kidney, ovary, lung, and brain (Soderling and Beavo, 2000). PDE10 is expressed in testes and brain and hydrolyzes both cAMP and cGMP. PDE11 acts as a dual specific PDE with predominant expression in skeletal muscle, prostate, kidney, liver, pituitary and salivary glands, and testes (Essayan, 2001; Soderling and Beavo, 2000; Vignola, 2004). Taken together, PDEs represent a diverse family of enzymes with highly specific enzymatic activity for hydrolyzing cyclic nucleotides, regulating diverse and important cellular functions due to diVerential expression, localization, and mode of regulation.

II. Developmental and Physiological Responses Regulated by PDEs Pharmacological inhibition of PDEs activates cAMP‐dependent pathways, which mimic short‐term eVects of cAMP‐mobilizing hormones and neurotransmitters. In vivo, if long‐term cell homeostasis and diVerentiation are disrupted by increases in cAMP levels, then the inactivation of PDE promotes loss of physiological function (Jin et al., 1999). Studies in Drosophila, zebrafish, and mice with specific PDE knockout support this notion. The inactivating mutation of dunce, a Drosophila analogue of PDE4, induces learning and memory defects in the fly (Davis and Dauwalder, 1991; Dudai et al., 1976). PDE6 was also found as a critical downstream eVector in Wnt/Ca2þ/cGMP signaling during mammalian development mediated by Frizzled‐2 (Ahumada et al., 2002; Wang et al., 2004). Interestingly, in humans, little is known regarding inherited disorders due to loss of PDE function with the exception of PDE6 (Conti and Jin, 1999). PDE6 regulates cGMP levels and is highly expressed in retina. Genetic evidence linked mutational inactivation of PDE6 genes to retinal degeneration and blindness in mammalians (Conti and Jin, 1999). Apparently, loss of PDE6 activity in retina increases cGMP levels that, in part, destroy retinal cells.

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To dissect the function of other PDEs in vivo, transgenic mouse models with disrupted PDE1B (Reed et al., 2002), PDE4B (Ariga et al., 2004), PDE4D (Jin et al., 1999), PDE7A (Yang et al., 2003), PDE10A (Siuciak et al., 2006), and PDE11 (Wayman et al., 2005) were generated. PDE1B‐ deficient mice exhibit exaggerated locomotor hyperactivity and have impaired spatial learning (Reed et al., 2002). Similarly, behavioral testing of PDE10A‐deficient mice revealed that PDE10A inhibits striatal output by reducing medium spiny neuron excitability, suggesting that PDE10A inhibition may serve as a therapeutic target for treatment of behavioral disorders (Siuciak et al., 2006). Mice deficient in PDE4D or PDE11 had profoundly altered reproductive function. PDE4D‐deficient female and male mice showed decreased growth; female mice also demonstrated reduced fertility due to impaired ovulation and diminished gonadotropin sensitivity in granulose cells (Jin et al., 1999). PDE11 knockout mice, while having no gross abnormalities, also showed defects in sperm physiology such as spermatogenesis and fertilization potential (Wayman et al., 2005). In inflammatory cells, activation of cAMP‐dependent signaling pathways inhibits immune and inflammatory responses, including T‐cell activation and proliferation, cytokine release, and recruitment of leukocytes (Essayan, 2001). Since airway inflammation plays a critical role in airway hyperresponsiveness and remodeling in asthma, the inhibition of PDEs was used as a therapeutic strategy to increase cAMP levels and to suppress inflammatory responses (Chung, 2006; Giembycz, 2005; Spina, 2004). The investigation of cellular mechanisms of these eVects revealed the diVerential roles of PDE4 isoforms in airway inflammation. Deficiency of PDE4B and PDE4D, but not PDE4A, markedly aVected neutrophil function in the mouse model of acute lung injury induced by endotoxin inhalation (Ariga et al., 2004). Neutrophil number in bronchoalveolar lavage (BAL) was markedly decreased in LPS‐ exposed PDE4B
/
and PDE4D
/
mice. Decreased neutrophil recruitment occurred due to reduced chemotaxis of the neutrophils, which was associated with decreased CD18 expression in PDE4B‐ and PDE4D‐deficient mice (Ariga et al., 2004). The role of PDE4B and PDE4D appears to be specific for regulating T‐cell (Torphy et al., 1993) and neutrophil functions; in contrast, the deletion of other PDE isoforms in mice (Yang et al., 2003), PDE7A, showed no deficiencies in T‐cell proliferation or cytokine production. While PDE4B‐ and PDE4D‐deficient mice show similar eVects on neutrophil chemotaxis and complement, each controlling neutrophil function, there exist subtle functional diVerences. PDE4B
/
leukocytes produce tumor necrosis factor‐ (TNF‐) in response to LPS; in contrast, stimulation of PDE4D
/
leukocytes with LPS does not generate TNF‐ (Jin and Conti, 2002). Similarly, PDE4B is critical for LPS‐induced TNF‐ production in macrophages (Jin et al., 2005). Collectively, PDE4B and/or PDE4D

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modulate systemic inflammatory responses by regulating expression of cell adhesion molecules, T‐cell proliferation, cytokine production, and neutrophil chemotaxis. Surprisingly, mice deficient in PDE4D also revealed a critical role of this PDE isoform in regulating ASM excitation–contraction coupling and inflammation‐induced remodeling (Hansen et al., 2000). While PDE4 deletion in mice inhibited inflammatory responses, PDE inhibitors may promote heart failure and cardiac arrhythmias, although the molecular pathways mediating these eVects remain unclear. PDE4D‐deficient mice showed progressive cardiomyopathy, accelerated heart failure after myocardial infarction, and cardiac arrhythmias (Lehnart et al., 2005). Other studies found that PDE4D3 complexes with the cardiac ryanodine receptor (RyR2)/calcium‐release channel, whose activation is critical for excitation– contraction coupling in heart muscle. In PDE4D
/
mice, RyR2 channels were hyperphosphorylated by PKA and exhibited a ‘‘leaky’’ phenotype, suggesting that PDE4D deficiency may promote heart failure and arrhythmias by defective regulation of the RyR2 channel (Lehnart et al., 2005). Collectively, studies in Drosophila, zebrafish, and mice demonstrated that diVerent classes of PDEs are critical for development and for physiological functions from flies to mammals.

III. PDEs and Airway Smooth Muscle Function A. PDE and ASM Tone and Contractility Airway smooth muscle (ASM) acts as the primary eVector cell regulating bronchomotor tone. Early studies of ‐adrenergic agonists discovered that short‐term (20 min) incubation of guinea pig tracheas with catecholamines stimulated cAMP production leading to ASM cell relaxation (Douglas et al., 1977). Pharmacological inhibition of PDEs with the nonselective PDE inhibitor, theophylline, further demonstrated that augmentation of cAMP levels evokes ASM relaxation (Newman et al., 1978; Polson et al., 1978; Triner et al., 1977). ASM relaxation is also stimulated by ‐2‐adrenoreceptor agonists such as isoproterenol, albuterol, salmeterol, or formoterol, which are used as therapeutic agents that promote bronchodilation by stimulating receptors on ASM coupled to stimulatory G‐proteins (Fig. 1). Gs protein stimulation activates adenylyl cyclase, increased cAMP formation, and stimulation of PKA. Activated PKA, in part, stimulates calcium‐activated potassium channels leading to ASM relaxation (Kume et al., 1992, 1994). The critical role of cAMP‐PDE signaling in regulating ASM shortening and its relevance to asthma were demonstrated in the study showing that allergen challenge of ASM in vivo promotes PDE activity and desensitizes ASM responsiveness to ‐agonists (Austin et al., 1987; Bai and Sanderson, 2006;

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Hedman and Andersson, 1982). These observations not only characterized the cAMP‐PDE signaling cascade in ASM, but also suggest a role for ‐adrenoreceptor agonists and PDE inhibitors in the management of airways disease (Chung, 2006; Spina, 2004; Vignola, 2004). The recognition of multiple PDE isoforms and their diVerential expression depending on cell or tissue type led to the characterization of PDE isoforms expressed in ASM providing opportunities for development of selective inhibitors. Biochemical studies revealed the presence of PDE1, PDE2, PDE3, PDE4, and PDE5 isozymes in tissue samples prepared from human trachea (Torphy et al., 1993) and bronchi (Rabe et al., 1993). Importantly, while inhibition of both PDE isoforms PDE3 and PDE4 induced relaxation of ASM in trachealis, PDE3 and PDE4 inhibition had diVerential eVects on spontaneous tone and shortening induced by carbachol (Barnes, 1995; Torphy and Cieslinski, 1990). Thus, in contrast to PDE3 inhibition, spontaneous tone was not aVected by PDE4 inhibition, but promoted relaxation of ASM stimulated by carbachol (Barnes, 1995). Clinical studies demonstrated that inhibition of PDE3 promotes bronchodilation, suggesting that PDE3 regulates airway caliber and ASM function (Bardin et al., 1998). The critical role of PDE4 in regulating ASM contractility was supported in studies using PDE4‐deficient mice (Hansen et al., 2000; Mehants et al., 2003). Targeted disruption of PDE4D induced the complete absence of airway responsiveness to methacholine in both allergen‐challenged and naive PDE4D
/
mice (Hansen et al., 2000). However, serotonin‐induced airway obstruction was unaVected by PDE4D deficiency. In contrast, PDE4D deficiency had little eVect on antigen‐induced airway inflammation. These interesting studies suggest that PDE4D inhibition can selectively inhibit airway obstruction to some but not all agonists and that PDE4D inhibition uncouples allergen‐ induced inflammation from allergen‐induced airway hyperresponsiveness. Pharmacological inhibition of PDE4 in humans suggests that PDE4 also plays a role in modulating bronchomotor tone; specifically, PDE4 inhibitors show potential eVectiveness in the treatment of asthma and COPD.

B. PDE and ASM Growth and Migration ASM hyperplasia contributes to the airway remodeling in asthma. Frequent stimulation of ASM by agonists, inflammatory mediators, and growth factors induces, in part, adaptive alterations in the airways that increase the size and numbers of myocytes leading to alterations in ASM mass. Such alterations have important consequences in determining airway caliber and ASM force generation (Gabella, 1979; Malqvist and Arner, 1988; Owens et al., 1988). Elevation of intracellular cAMP levels had been linked to inhibition of

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ASM proliferation (Tomlinson et al., 1995) due to cyclin D1 degradation and cell cycle arrest in G1 phase (Stewart et al., 1999). These studies suggest that increased cAMP levels may abrogate ASM growth due to increased cell proliferation, and inhibition of PDE activity catalyzing cAMP breakdown may serve as a potential therapeutic approach in the prevention of myocyte proliferation (Billington et al., 1999). Proliferation of ASM is associated with cell adhesion to extracellular matrix, which is critical to cell cycle progression (Roovers and Assoian, 2003). Histological findings suggest that proliferating ASM cells adhere to extracellular matrix and migrate along chemotactic gradients, as evidenced by the invasion of smooth muscle‐like cells into the submucosa (Gizycki et al., 1997; JeVery, 2004). Elevated cAMP levels inhibit cell migration in a variety of cell types (Madison, 2003). Accordingly, PDE4B and PDE4D deficiency attenuates chemotaxis of neutrophils due to decreased expression of cell adhesion molecule CD18 (Ariga et al., 2004). Treatment of ASM cell cultures with cAMP‐mobilizing agents such as prostaglandin E2 (PGE2) and salmeterol, or the PDE4 inhibitor cilomilast, significantly inhibited ASM migration (Goncharova et al., 2003). Importantly, the inhibition of PDE4 attenuated PDGF‐induced migration of ASM, suggesting that PDE modulates growth factor‐induced ASM migration. Collectively, evidence supports a critical role of PDEs in regulating ASM contractility, proliferation, and migration.

IV. Cytokine and Chemokine Secretion by ASM: A Role for PDEs Asthma is not only characterized by reversible airway obstruction due to ASM contractility, but also by airway inflammation which is due to the recruitment and activation of inflammatory cells. These cells secrete cytokines and chemokines that can directly regulate ASM function. Therefore, targeted inhibition of PDE4, the predominant isozyme in neutrophils, macrophages, and T cells, potentially could serve as an anti‐inflammatory signal. The eVects of selective and dual specificity PDE inhibitors in the management of airway inflammation associated with asthma and chronic obstructive pulmonary disease were reviewed (Chung, 2006; Giembycz, 2005; Spina, 2004; Vignola, 2004). PDE4 inhibitors decrease airway inflammation, prevent inflammatory cell proliferation, suppress generation of reactive oxygen species in leukocytes, and upregulate expression of anti‐inflammatory cytokines such as interleukin‐10 (IL‐10) (Vignola, 2004). Current evidence shows that ASM not only plays a major role in regulating airway obstruction but also functions as an immunomodulatory cell (Ammit and Panettieri, 2001; Amrani and Panettieri, 2002). ASM secretes

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a number of cytokines and chemoattractants, and bronchial biopsies of ASM in mild asthmatics have revealed constitutive expression of RANTES (regulated on activation, normal T‐cell expressed and secreted) (Berkman et al., 1996), the secretion of which is induced by TNF‐ and interferon‐ (IFN‐ ) in vitro (Ammit et al., 2000). Other chemokines secreted by ASM cells include IL‐8, eotaxin, an eosinophil chemoattractant, and monocyte chemoattractant protein (MCP)‐1, MCP‐2, and MCP‐3 (Lazaar and Panettieri, 2001). ASM cell secretion of proinflammatory mediators and chemokines could serve to recruit inflammatory cells such as neutrophils, T cells (Lazaar et al., 1994), eosinophils, and macrophages that may promote ASM contractility and remodeling. In ASM cells, intracellular levels of cAMP modulate the expression of proinflammatory cytokines and chemokines, which serve as chemoattractants for inflammatory cells and also potentially modulate myocyte function in an autocrine manner. In TNF‐‐stimulated ASM cells, expression of chemokines, eotaxin, and RANTES was inhibited by isoproterenol or PGE2, or by the PDE inhibitors rolipram and cilomilast (Ammit et al., 2000; Pang and Knox, 2001). TNF‐‐induced IL‐8 secretion was also inhibited by the cAMP‐mobilizing agents (Pang and Knox, 2000). Activation of cAMP signaling attenuates both TNF‐‐mediated induction of adhesion molecule ICAM‐1 and VCAM‐1 expression and adhesion of activated T cells to ASM cells (Lazaar et al., 1994). Surprisingly, other cytokines such as IL‐6 are increased by PDE inhibition or by cAMP‐mobilizing agents. Taken together, current evidence suggests that PDEs can play a role in regulating ASM cell function by modulating synthetic function of ASM and recruitment of inflammatory cells in airways.

V. Conclusions PDEs serve as a unique target to decrease bronchoconstriction while acting as an anti‐inflammatory agent in asthma and COPD. While significant progress has been made in characterizing the role for PDEs in regulating ASM function and regulation, further studies are needed to elucidate the cellular and molecular mechanisms by which PDEs are regulated in ASM and how this regulation contributes to ASM contractility and remodeling.

Acknowledgments This work was supported in part by grants from the National Heart, Lung, and Blood Institute to R.A.P. and V.P.K., and from GlaxoSmithKline to R.A.P.

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Lehnart, S. E., Wehrens, X. H. T., Reiken, S., Warrier, S., Belevych, A. E., Harvey, R. D., Ruchter, W., Jin, C., Conti, M., and Marks, A. R. (2005). Phosphodiesterase 4D deficiency in the ryanodine‐receptor complex promotes heart failure and arrhythmias. Cell 123, 25–35. Madison, J. M. (2003). Migration of airway smooth muscle cells. Am. J. Respir. Cell Mol. Biol. 29, 8–11. Malqvist, U., and Arner, A. (1988). Contractile properties during development of hypertrophy of the smooth muscle in the rat portal vein. Acta Physiol. Scand. 133, 49–61. Mehants, C., Jin, S.‐L. C., Wahlstrom, J., Law, E., Umetsu, D. T., and Conti, M. (2003). PDE4D plays a critical role in the control of airway smooth muscle contraction. FASEB J. 17, 1831–1841. Newman, D. J., Colella, D. F., Spainhour, C. B., Jr., Brann, E. G., Zabko‐Potapovich, B., and Wardell, J. R., Jr. (1978). cAMP‐phosphodiesterase inhibitors and tracheal smooth muscle relaxation. Biochem. Pharmacol. 27, 729–732. Owens, G. K., Geisterfer, A. A. T., Wei‐Hwa Yang, Y., and Komoriya, A. (1988). Transforming growth factor‐ ‐induced growth inhibition and cellular hypertrophy in cultured vascular smooth muscle cells. J. Cell Biol. 107, 771–780. Pang, L., and Knox, A. J. (2000). Synergistic inhibition by 2‐agonists and corticosteroids on tumor necrosis factor‐‐induced interleukin‐8 release from cultured human airway smooth‐ muscle cells. Am. J. Respir. Cell Mol. Biol. 23, 79–85. Pang, L., and Knox, A. J. (2001). Regulation of TNF‐‐induced eotaxin release from cultured human airway smooth muscle cells by 2‐agonists and corticosteroids. FASEB J. 15, 261–269. Polson, J. B., Kazanowski, J. J., Goldman, A. L., and Szentivanyi, A. (1978). Inhibition of human pulmonary phosphodiesterase activity by therapeutic levels of theophyline. Clin. Exp. Pharmacol. Physiol. 5, 535–539. Rabe, K. F., Tenor, H., Dent, G., Schudt, C., Liebig, S., and Magnussen, H. (1993). Phosphodiesterase isozymes modulating inherent tone in human airways: Identification and characterization. Am. J. Physiol. 264, L458–L464. Reed, T. M., Repaske, D. R., Snyder, G. L., Greengard, P., and Vorhees, C. V. (2002). Phosphodiesterase 1B knock‐out mice exhibit exaggerated locomotor hyperactivity and DARPP‐32 phosphorylation in response to dopamine agonists and display impaired spatial learning. J. Neurosci. 22, 5188–5197. Roovers, K., and Assoian, R. K. (2003). EVects of Rho kinase and actin stress fibers on sustained extracellular signal‐regulated kinase activity and activation of G1 phase cyclin‐dependent kinases. Mol. Cell. Biol. 23, 4283–4294. Siuciak, J. A., McCarthy, S. A., Chapin, D. S., Fujiwara, R. A., James, L. C., Williams, R. D., Stock, J. L., McNeish, J. D., Strick, C. A., Menniti, F. S., and Schmidt, C. J. (2006). Genetic deletion of the striatum‐enriched phosphodiesterase PDE10A: Evidence for altered striatal function. Neuropharmacology 51, 374–385. Soderling, S. H., and Beavo, J. A. (2000). Regulation of cAMP and cGMP signaling: New phosphodiesterases and new functions. Curr. Opin. Cell Biol. 12, 174–179. Spina, D. (2004). The potential of PDE4 inhibitors in respiratory disease. Curr. Drug Targets Inflamm. Allergy 3, 231–236. Stewart, A. G., Harris, T., Fernandes, D. J., Schachte, L. C., Koutsoubos, V., Guida, E., Ravenhall, C. E., Vadiveloo, P., and Wilson, J. W. (1999). 2‐Adrenergic receptor agonists and cAMP arrest human cultured airway smooth muscle cells in the G1 phase of the cell cycle: Role of proteasome degradation of cyclin D1. Mol. Pharmacol. 56, 1079–1086. Sutherland, E. W., and Rall, T. W. (1958). Fractionation and characterization of cyclic adenine ribonucleotide formed by tissue particles. J. Biol. Chem. 232, 1077–1092. Tomlinson, P. R., Wilson, J. W., and Stewart, A. G. (1995). Salbutamol inhibits the proliferation of human airway smooth muscle cells grown in culture: Relationship to elevated cAMP levels. Biochem. Pharmacol. 49, 1809–1819.

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Role of Astrocytes in Matching Blood Flow to Neuronal Activity Danica Jakovcevic and David R. Harder Department of Physiology, Cardiovascular Research Center Medical College of Wisconsin, Milwaukee, Wisconsin 53226

I. Introduction II. Neurovascular Unit III. Neuron–Astrocyte Interactions and Regulation of Synaptic Transmission in the Brain A. Ca2þ Signaling in Astrocytes B. Astrocyte Release of Glutamate and Modulation of Synaptic Transmission C. Neurometabolic Coupling IV. Neurovascular Coupling: Role of Astrocytes A. Role of Astrocytic Ca2þ Elevations in Coupling Neuronal Activity to the Vasculature B. Vasoactive Mediators of Neurovascular Coupling V. Summary References

The brain is critically dependent on oxygen and glucose supply for normal function. Various neurovascular control mechanisms assure that the blood supply of the brain is adequate to meet the energy needs of its components. Emerging evidence shows that neuronal activity can control microcirculation using astrocytes as a mediator. Astrocytes can sense neuronal activity and are involved in signal transmission. Synaptic activity triggers an increase in the intracellular calcium concentration [Ca2þ]i of adjacent astrocytes, stimulating the release of adenosine triphosphate (ATP) and glutamate. The released ATP mediates the propagation of Ca2þ waves between neighboring astrocytes, thereby recruiting them to mediate adequate cerebrovascular response to neuronal activation. Simultaneously, sodium‐dependent glutamate uptake in astrocytes generates Naþ waves and subsequently increases glucose uptake and metabolism that leads to the formation of lactate, which is then delivered to neurons as an energy substrate. Further, astrocytic Ca2þ elevations can lead to secretion of vasodilatory substances from perivascular endfeet, such as epoxyeicosatrienoic acid (EETs), adenosine, nitric oxide (NO), and cyclooxygenase‐2 (COX‐2) metabolites, resulting in increased local blood flow. Thus, astrocytes by releasing vasoactive molecules mediate the neuron‐astrocyte‐endothelial signaling pathway and play a profound role in coupling blood flow to neuronal activity. ß 2007, Elsevier Inc. Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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I. Introduction Glial cells were originally described by Virchow (1846) as nonneuronal cells constituting the ‘‘glue’’ of the brain. Later studies classified glial cells in the central nervous system (CNS) as astrocytes, oligodendrocytes, and microglia, each of which has diVerent histological characteristics and functions (Ramon y Cayal, 1911). Once thought to be merely supportive elements, maybe due to the fact that they are nonexcitable cells, glial cells have made the big leap toward the ‘‘stars of the show’’ (Hertz and Zielke, 2004). The big part of the change in their status can be attributed to the findings that glia can integrate neuronal inputs and modulate synaptic activity. In addition, astrocytes can also maintain milieu around the active neurons by regulating extracellular Kþ concentration, volume, osmolarity, pH, and concentration of neurotransmitters, particularly glutamate and
‐aminobutyric acid (GABA) at the synaptic cleft. Astrocytes are in close association with neurons, can enwrap synaptic terminals (Ventura and Harris, 1999), and have foot processes ensheathing the capillaries. Because of their anatomical location, they were thought for a long time to have an intermediary role in matching neuronal activity to cerebral blood flow (CBF) and metabolism. When neurons in specific brain regions are highly activated, blood flow increases in a temporally and spatially coordinated manner. This coupling between neuronal activity and blood flow, termed functional hyperemia, was first described by Mosso (1880) and confirmed by Roy and Scherrington (1890). As a consequence of neuronal activation blood flow increases in the active area within seconds and thus ensures adequate supply of oxygen and glucose. This tight coupling between neuronal activity, metabolism, and blood flow has provided the basis for functional brain imaging techniques. Positron emission tomography (PET) uses the coupling between synaptic activity and glucose utility, while functional magnetic resonance imaging (fMRI) is based on the fact that the change in neuronal activity produces variations in the level of hemoglobin oxygenation (Seiyama et al., 2004). The cellular mechanisms of neurovascular coupling have recently been defined and continue to be investigated (Harder et al., 1998, 2000, 2002; Metea and Newman, 2006). Our group has contributed over the last 10 years to the understanding of such cellular and molecular mechanisms (see Section IV.B.5).

II. Neurovascular Unit In 1885, Golgi first described the contact of astrocytic foot processes with arterioles and capillaries (Golgi, 1885). By sending specialized processes to both the vasculature and synaptic contacts, astrocytes have a unique

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anatomical position between neurons and arterioles to couple neuronal activity to blood supply. Astrocytes are small cells characterized by small somata (<10 mm) and numerous highly branched processes. The astrocytic process is in close contact with both pre‐ and postsynaptic terminals (Ventura and Harris, 1999). It is estimated that one astrocyte makes more than 100,000 synaptic contacts, which suggests their important role in synaptic regulation. Astrocytes sense synaptic activity and accordingly release a variety of vasoactive molecules such as glutamate, adenosine triphosphate (ATP), nitric oxide (NO), prostaglandin, epoxyeicosatrienoic acid (EETs), and D‐serine. Their role as an active element of the synapse together with pre‐ and postsynaptic terminals has led to the term ‘‘tripartite synapse’’ (Araque et al., 1999a). It is now clear that groups of neurons and their associated astrocytes are functionally coupled to smooth muscle cells (SMC) and endothelial cells on the microvessels, thereby regulating their blood supply. These functional units are called neurovascular units (Harder et al., 2002; Iadecola, 2004; Lo et al., 2003). Defining the close interactions among neurons, astrocytes, and blood vessels is the key for understanding the mechanisms of coupling between neuronal activity and blood flow. Blood flow to the brain is provided by extracerebral and intracerebral arteries and arterioles. Large arteries supply branching pial arterioles that dive deeper into the brain (Jones, 1970). In general, extracerebral vessels are innervated by peripheral nerves, while intracerebral microvessels are regulated by local interneurons and central neuronal terminals (Hamel, 2006). Arteries and arterioles consist of an endothelial cell layer, SMC, and adventitia (Peters et al., 1991). The penetrating arteriole is surrounded by an invagination of the pia mater, which forms a perivascular space (Virchow–Robin space). When the basement membranes of the pia and the penetrating arteriole coalesce, the space disappears and the vessel becomes a capillary. Capillaries consist of endothelial cells, pericytes, and capillary basal lamina on which astrocytic endfeet are attached. Cerebral endothelial cells are not fenestrated and are connected by tight junctions, which prevent large molecules from passing freely into the brain tissue, and thus present the basis of the blood–brain barrier. Moreover, by exerting its influence on SMC through myoendothelial gap junctions or by the release of vasoactive factors, endothelial cells have an important role in the control of CBF. Endothelial cells produce vasodilator agents, such as NO, prostacyclin, carbon monoxide, endothelium‐derived hyperpolarizing factor (EDHF), and vasoconstrictor agents such as endothelin, tromboxan A2 (TXA2), and prostaglandin F2 (PGF2) (Andresen et al., 2006; Faraci and Heistad, 1998). NO produced by endothelial NO synthase (eNOS) contributes to the basal tone of cerebral arteries (Atochin et al., 2003; Prado et al., 1992). In addition, NO derived from neuronal NO synthase (nNOS), located in

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interneurons with perivascular projections, can also dilate cerebral arteries (Gotoh et al., 2001; Iadecola et al., 1993; Ignacio et al., 1997). Thus, physical and biochemical stimuli induce release of chemical signals from endothelial cells, astrocytes, and neurons that produce changes in arteriolar smooth muscle tone by causing intracellular Ca2þ elevations and alternations in the phosphorylation state of myosin that are presented as a change in vascular diameter. Moreover, pial arteries oVer the greatest resistance to flow. Therefore during functional hyperemia, vasodilation of the downstream arterioles in the area of activation is not enough to increase the CBF eVectively unless upstream vessels also dilate. Indeed, it has been found that upstream dilations in cerebral arteries occur following neuronal activation (Cox et al., 1993). The mechanism for the propagation of vasodilation from the initial site of activation to upstream arteries is still not clear. Communication between endothelial cells through gap junctions (Collins et al., 1998) as well as myoendothelial gap junctions that link neighboring endothelial cells and SMC (Little et al., 1995; Sandow and Hill, 2000) could be involved in these coordinated vascular responses. Also, vasodilation of downstream branches increases blood flow velocity in upstream vessels and thus increases shear stress on endothelial cells, which produces further vasodilation by the release of vasoactive metabolites from these cells (Busse and Fleming, 2003; Koller et al., 1993). Taken together, neurovascular signaling is a complex event regulated at diVerent levels of the neuron‐astrocyte‐endothelial cell pathway and requires coordinated vasoactive responses within the vascular network. The purpose of this chapter is to define the role of astrocytes in these highly regulated neurovascular units, particularly related to functional hyperemia.

III. Neuron–Astrocyte Interactions and Regulation of Synaptic Transmission in the Brain A. Ca2þ Signaling in Astrocytes Neurons form neuronal networks via synapses while astrocytes form syncytium‐like organizations via gap junctions. Gap junctions are hemichannels that allow the passage of small molecules between cytoplasm of adjacent cells. The major constituent of these hemichannels is connexin 43. These hemichannels have low open permeability in physiological states, but some conditions can change permeability such as an increase in extracellular Kþ and low extracellular Ca2þ (Giaume and McCarthy, 1996). Astrocytes have ionotropic and metobotropic glutamate receptors (mGluR). Glutamate released in the synaptic space binds to mGluR on astrocytes and

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elicits an increase in intracellular Ca (Bezzi et al., 2001; Haydon, 2001). However, the mechanism responsible for the mGluR‐stimulated Ca2þ increase in astrocytes is not completely known. But, it appears now that the activation of G‐protein–coupled receptors leads to the activation of phospholipase C (PLC), resulting in the production of inositol‐1,4,5‐trisphosphate (IP3) and diacylglycerol (DAG). IP3 causes the release of Ca2þ from IP3‐sensitive Ca2þ stores (endoplasmatic reticulum). This increase in intracellular Ca2þ triggers signaling within the astrocytic network (Verkhratsky and Kettenmann, 1996). First studies performed in cultured cells described that synaptically released glutamate induces transient increases of Ca2þ in astrocytes (Cornell‐Bell et al., 1990; Smithson et al., 1992). These elevations spread between adjacent astrocytes indicating a form of long‐range signaling. More recent studies performed in brain slices have shown that glutamate‐dependent Ca2þ signaling is present in nervous tissue and not just in culture cells (Dani et al., 1992; Porter, 1996). One study showed that in acute hippocampal slices axonal stimulation induces glutamate‐dependent Ca2þ increases in astrocytes, which might be mediated by prostaglandin production (Bezzi et al., 1998). Astrocytes have a variety of receptors, and thus, they can respond to several other synaptically released transmitters, such as noradrenaline, histamine, acethylcholine, ATP, and GABA, with an increase in intracellular Ca2þ. In 1990, Cornell Bell showed in cell culture that synaptically released glutamate induces Ca2þ elevation in astrocytes, and that these Ca2þ signals could be spread from one astrocyte to another (Cornell‐Bell et al., 1990). Two hypotheses have emerged that attempt to explain the mechanism by which the Ca2þ waves can spread in neighboring astrocytes. It was thought that Ca2þ waves are mediated by IP3 propagating through gap junction channels (Sanderson et al., 1994; Sneyd et al., 1995). Thus, IP3 produced in astrocyte can pass via gap junctions and induce release of Ca2þ from internal stores of neighboring cells. However, some studies have shown that IP3 diVusion by itself is not suYcient for maintenance of Ca2þ waves and that an extracellular signal might also be involved. This extracellular pathway is confirmed by the studies of Enkvist and Hassinger (Enkvist and McCarthy, 1992; Hassinger et al., 1996). The latter study has shown that Ca2þ waves can pass between disconnected cells as long as the distance between them does not exceed approximately 120 mm. It seems that the release of ATP by astrocytes is the main signaling mechanism involved. ATP may be released to the extracellular space by gap junctions and act via purinergic receptors (P2Y type) of neighboring astrocytes inducing IP3‐mediated mobilization of Ca2þ from internal stores (Cotrina et al., 1998a; Guthrie et al., 1999). In conclusion, ATP is responsible for IP3 regeneration and long‐range Ca2þ signaling, while cells that are closer together likely utilize diVusion of IP3 to propagate calcium signals.

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B. Astrocyte Release of Glutamate and Modulation of Synaptic Transmission Calcium elevation in astrocytes triggers release of glutamate (Parpura et al., 1994), most likely by exocytosis. Glutamate release from astrocytes is Ca2þ dependent and injection of clostridial toxins into astrocytes can block the calcium‐dependent glutamate release (Araque et al., 2000; Bezzi et al., 1998). However, other types of nonexocytotic transmitter release have been described. For example, volume‐regulated anion channels are involved in transport of anionic amino acids (glutamate, taurin) when astrocytes swell, together with reverse operation of glutamate transporters (Szatkowski et al., 1990). The purinergic P2X receptors and gap junction hemichannels seem to be involved in glutamate release in conditions of low extracellular Ca2þ concentrations, which increase their opening probability. Nevertheless, glutamate could also be released by cystine/glutamate exchangers (Melendez et al., 2005), which are involved in cystine accumulation in astrocytes. Cystine is necessary for the production of glutathione, an important antioxidant in the brain (Dringen, 2000). As mentioned before, glial cells are now considered a part of the ‘‘tripartite synapse.’’ They integrate the signals from the synapses, respond via Ca2þ elevations and release of gliotransmitters that can modulate synaptic transmission in a feedback manner. Using a variety of stimuli that elevate astrocytic Ca2þ (bradykinin, photo‐, and electrical stimulation), previous cell culture studies showed that the Ca2þ elevation in astrocytes led to Ca2þ elevations in adjacent neurons (Araque et al., 1999b; Hassinger et al., 1996; Nedergaard, 1994; Parpura et al., 1994). In 1997, Pasti described in her study, for the first time in brain slices, that astrocytes can signal to neurons by releasing glutamate (Pasti et al., 1997). She also showed that glutamate released from astrocytes induced neuronal Ca2þ elevations. Nedergaard (1994) reported that astrocyte‐neuron signaling was attenuated by gap junction blockers, suggesting the role of gap junctions in mediating this signal. In 1998, Araque first showed that the glutamate released from astrocytes can modulate synaptic transmission in hippocampal neurons (Araque et al., 1998a,b). Further, Robitaille (1998) has demonstrated that neurotransmitters released by the motor nerves activate the perisynaptic Schwann cell, which in turn feeds back to the nerve modulating neurotransmitter release. In addition to glutamate and ATP, the Ca2þ signal in astrocytes also triggers release of D‐serine, homocysteic acid, and arginine which can aVect glutamatergic neurotransmission (Fellin and Carmignoto, 2004). D‐Serine acts at the glycine site of the N‐methyl‐D‐aspartate (NMDA) receptor and facilitates Ca2þ influx through this channel (Baranano et al., 2001). The detailed description of the modulatory eVect of gliotransmitter on synaptic transmission is out of the scope of this chapter, but the reader is referred

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to recent reviews (Araque et al., 1999b; Haydon, 2001; Haydon and Carmignoto, 2006). Bidirectional neuron–astrocyte interactions are critical for normal brain function, and our understanding of the mechanisms involved may help us discover the right treatment for a variety of neurological diseases.

C. Neurometabolic Coupling The brain represents only 2% of our body mass, but its energy requirements are very high. It uses 20% of the whole oxygen and glucose consumption of the body. The location of astrocytic foot processes at synapses and number of astrocytes in the brain (outnumbering neurons 10:1) appear as if astrocytes are ideal to sense synaptic activity and couple it to energy metabolism (Derouiche and Frotscher, 1991). Glutamate released during neuronal activity in the synaptic cleft is rapidly removed by neighboring astrocytes. Astrocytes contain excitatory amino acid transporters (EAAT) that uptake glutamate, thus preventing its accumulation in the synaptic cleft and excitotoxicity. Glutamate is cotransported with three Naþ ions that elicit intracellular sodium elevations. This Naþ elevation results in an activation of the Naþ/Kþ‐ATPase, causing an increased energy demand in astrocytes, which in turn enhances glycolysis. Simultaneously, glutamate‐activated Ca2þ increase in astrocytes stimulates glutamate and ATP release. Released glutamate is taken up by Na/Glu transporters and triggers an intracellular Naþ wave (Bernardinelli et al., 2004). Sodium also has the ability to diVuse through gap junctions and participate in Naþ wave extensions (Rose and Ransom, 1997). It appears that glutamate release and reuptake help sustain the regenerative propagation of the Naþ signal. Therefore, Naþ waves could recruit surrounding astrocytes to increase glucose uptake and metabolism, coordinating metabolic substrates to local neuronal activity. Briefly, glucose is transported via glucose transporter (GLUT)‐1 from endothelial cells to astrocytic foot processes. Glucose can also enter neurons directly via GLUT‐3. In the astrocyte, glucose is used for the production of glycogen and glycolysis. The main product of glycolysis in astrocytes is lactate. Lactate is transported out of the astrocyte and is taken up by monocarboxylate transporters on the neuron where lactate is converted into pyruvate, which is used as a substrate for oxidative metabolism (Benarroch, 2005; Charles, 2005; Magistretti and Pellerin, 1999; Magistretti et al., 1999). This astrocyte‐neuron lactate shunt is assumed to be critical for metabolic support of active neurons, although it is argued that glucose may be a direct source of energy during neuronal activity. Parras has reported that the exposure of neuron to glutamate inhibits glucose uptake by neurons, providing

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additional evidence that neurons rely on astrocytes for energy metabolism (Parras et al., 2004). Although Ca2þ signals mediate astrocytic control of brain microcirculation, their physiological role is still unclear. Ca2þ waves through glutamate release/reuptake and resulting Naþ waves can send a metabolic signal and provide energy substrates (lactate) to neurons.

IV. Neurovascular Coupling: Role of Astrocytes A. Role of Astrocytic Ca2þ Elevations in Coupling Neuronal Activity to the Vasculature Because of the anatomical position and the vicinity of foot processes to contractile elements in blood vessels, astrocytes have long been thought to marginally contribute to the regulation of CBF. Studies over the past few years show that synaptic activity, and not the energy deficit due to glucose or oxygen use, is the trigger for neuronal activity‐dependent vasodilation (Sandor, 1999). Significant evidence supporting a role of astrocytes in regulation of CBF comes from the brain slice preparations. Several studies demonstrated that astrocytes are more likely compartmentalized because Ca2þ oscillations were noticed in small portions of the astrocytic foot processes, the so‐called ‘‘microdomains’’ (Grosche et al., 1999; Pasti et al., 1997). Pasti described in her study that glutamate‐dependent Ca2þ oscillations in astrocytes increased in frequency according to the increased level of synaptic activity. This observation and the reports from other studies that Ca2þ oscillations propagate to perivascular endfeet (Zonta et al., 2003a) indicate that astrocytes act as sensors of synaptic activity and then send messages to blood vessels about its intensity. However, some studies in brain slices show that the range of Ca2þ signaling is less extensive than those observed in cultures (Sul et al., 2004). It is likely that ectonucleotidases, enzymes that hydrolyze ATP to AMP, are more eVective in extracellular spaces of the brain slice. In that case, rapid degradation of ATP could explain, at least in part, the much smaller range of Ca2þ signaling. Further, Filosa et al. (2004) have shown the first evidence of rapid neurovascular coupling in brain slices that is consistent with functional hyperemia in vivo. The authors measured simultaneously Ca2þ oscillations in astrocytes and parenchymal arterioles. It is well known that arteriole at rest exhibits periodic contractions and relaxations that accompany Ca2þ oscillations in SMC, and this phenomenon is called vasomotion. By using electrical field stimulation, they induced elevations in astrocytic Ca2þ, while arteriolar Ca2þ oscillations and vasomotion were suppressed. This indicates that inhibition of vasomotion contributes to functional

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hyperemia. Moreover, another study in brain slices demonstrates that addition of miconazole, the epoxygenase inhibitor, increases the frequency of vasomotion, thereby indicating that EETs can contribute to inhibition of rhythmic contractions of vessels during neuronal activity (Lovick et al., 2005). The brain slice preparation has several limitations. First, a relevant part of synaptic connections is lost and second, absence of intraluminal flow and shear stress in slices decreases myogenic tone. This could explain significant delay (
30 s) between neuronal stimulation and vasodilation in brain slices (Zonta et al., 2003a) compared to in vivo experiments (1–2 s). In brain slice experiments, Zonta reported that Ca2þ elevations in astrocytic endfeet trigger dilation of cerebral arteries and that antagonist of mGluR significantly reduced hyperemic response to forepaw stimulation in vivo (Zonta et al., 2003a). However, Mulligan and MacVicar reported diVerent vascular response to astrocytic Ca2þ elevations in brain slice preparations (Mulligan and MacVicar, 2004). They utilized flash photolysis of caged Ca2þ to induce Ca2þ waves in astrocytes, which propagated to astrocytic endfeet causing vasoconstriction. The application of noradrenaline had the same eVect, and 0 0 ‐tetraacetic acid Ca2þ chelator 1,2‐bis(o‐aminophenoxy) ethane‐N,N,N,N (BAPTA) diminished Ca2þ elevations to noradrenaline. Moreover, the constrictor response to Ca2þ elevations was blocked by a phospholipase A2 (PLA2) inhibitor and by an inhibitor of 20‐hydroxyeicosatetraenoic acid (20‐HETE) synthesis. Thus, the authors proposed that astrocytic Ca2þ elevations activate PLA2 and mobilize arachidonic acid (AA) from membrane phospholipid pools. AA is then converted to 20‐HETE by P450 !‐hydroxylase in SMC. 20‐HETE inhibits calcium‐activated potassium channels (KCa) inducing cell depolarization, Ca2þ influx through voltage‐dependent Ca2þ channels and vasoconstriction (Gebremedhin et al., 2000). The contrast between findings of Zonta and Mulligan can be explained, at least in part, by diVerent experimental conditions. Zonta preincubated slices in N‐nitro‐L‐ arginine methyl ester (L‐NAME) to block NO formation and preconstrict blood vessels mimicking myogenic tone, while Mulligan examined only nonconstricted arterioles. Accordingly, when myogenic tone is lost, vessels are in a dilated state and the eVect of dilating agents can be lost or diminished. A recent in vivo study provided further support for the role of astrocytes in functional hyperemia (Takano et al., 2006). Photolysis of caged Ca2þ in astrocytic endfeet causes rapid vasodilation (1–2 s), and cyclooxygenase (COX) inhibitors blocked the photolysis‐induced vasodilation. In conclusion, evidence accumulated so far indicates that astrocytes can sense diVerent levels of neuronal activity and integrate them into defined Ca2þ oscillations that propagate to perivascular endfeet and cause the release of dilating agents, such as EETs, prostaglandin E2 (PGE2), adenosine, as well as the formation of constrictive agents such as 20‐HETE.

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B. Vasoactive Mediators of Neurovascular Coupling It has been shown that neurovascular coupling is a complex process and that astrocytes can produce a plethora of vasoactive agents that can contribute to regulation of CBF. Candidate vasodilator signals triggered by neuronal activity include Kþ ions, adenosine, NO, EETs, and PGE2. 1. Potassium Ions Because of the high Kþ conductance in astrocytic endfeet and the observation in Mu¨ller cells in the retina (Newman, 1986; Newman et al., 1984) that astrocytes can uptake Kþ ions released during neuronal activity and redistribute them through foot processes to the perivascular space, Kþ was considered as a mechanism to couple neuronal activity to blood flow (Trachtenberg and Pollen, 1970). Potassium would be ‘‘buVered’’ from the synaptic cleft via inward rectifier Kþ (Kir) channels located on astrocytic processes (Ransom and Sontheimer, 1995). However, it was shown that inhibition of Kþ uptake into astrocyte by barium, a Kþ channel blocker, did not decrease CBF response to electrical stimulation of the cortex (Kraig, 1989). 2. Nitric Oxide NO is synthesized from L‐arginine by the enzyme NO synthase (NOS) (Palmer et al., 1988). There are three isoforms of NOS: endothelial (eNOS), neuronal (nNOS), and inducible (iNOS). nNOS and eNOS exist under normal conditions and iNOS is expressed after exposure of the brain to cytokines or endotoxins (Brian et al., 1995; Faraci and Brian, 1994; Iadecola et al., 1994). NO has been implicated as a mediator of vasodilation during hypercapnia, coupling of blood flow to neuronal activation, and cortical spreading depression (Phillis, 1989; Wahl et al., 1994). The cerebral vasodilation induced by CO2 is attenuated by the inhibition of NOS (Iadecola, 1992), and this attenuation can be completely reversed by NO donors. nNOS is present in a small population of interneurons (Wang et al., 2005). Because NO is a highly diVusible molecule, the presence of NOS in neurons that lie near microvessels suggests that NO can diVuse from these neurons during activation and influence local vascular tone (Iadecola et al., 1993; Lovick et al., 1999). Activation of NMDA receptors of these neurons results in the rise in intracellular Ca2þ, which leads to NOS activation and NO production (Connor and Muller, 1991). The released NO then diVuses out of the cell and exerts its action by stimulating soluble guanylyl cyclase (sGC) in neighboring arterial SMC (Moncada et al., 1991). The accumulation of cyclic GMP (cGMP) leads to reduction in intracellular Ca2þ, myosin dephosphorylation, and SMC relaxation (Ahlner et al., 1991). Also, NO produced by nNOS participates in the maintenance of resting CBF (Cholet et al., 1997;

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Yang et al., 1999) and has a role in functional hyperemia. Inhibition of nNOS by 7‐nitroindazole (7‐NI) reduces the increase in CBF to whisker stimulation by 50–60% (Cholet et al., 1996; Lindauer et al., 1999). Studies in knockout mice show that L‐NNA, a nonselective inhibitor of NOS, attenuated functional hyperemia in eNOS knockout mice (Ayata et al., 1996), but not in nNOS knockout mice (Ma et al., 1996). Taken together, these data indicate that neuronal NO is one of the mediators involved in the vasodilatory response associated with somatosensory stimulation. Other studies, however, have reported no eVect of NOS inhibitors on the cerebrovasodilation to neuronal stimulation in somatosensory cortex (Wang et al., 1993). The reason for the discrepancies is not clear, although diVerences in the degree of NOS inhibition may play a role (Iadecola et al., 1995). Moreover, Lindauer demonstrated that NOS inhibitor as well as inhibitor of sGC attenuates functional hyperemia in the somatosensory cortex and can be restored by NO donors that elevate levels of cGMP. These data suggest that basal level of NO and cGMP is required to provide adequate CBF response to stimulation. It seems that there are regional diVerences in the role of NO in certain parts of the brain. NOS activity in rat cerebellum is tenfold higher than in the somatosensory cortex (Cholet et al., 1996), and glutamate‐induced vasodilation can be attenuated by NOS inhibitors but cannot be restored by NO donors or cGMP. Thus, in cerebellum NO acts more as a mediator rather than modulator because NOS activity is required for intact response. 3. Adenosine Evidence implicating adenosine in CBF regulation derives from experiments using adenosine antagonists and potentiators. Data from studies using models of cerebellar parallel fiber stimulation (Li and Iadecola, 1994) and whisker stimulation‐induced hyperemia (Dirnagl et al., 1994) demonstrate that theophylline, a nonselective adenosine receptor antagonist, attenuated the increase in CBF during stimulation and that coapplication of NOS inhibitor (L‐NNA) attenuates the vasodilation further. These data suggest that NO and adenosine contribute to increase in CBF during neuronal activation. However, some studies demonstrate the lack of eVect of caVeine, an adenosine receptor antagonist, on whiskers stimulation induced vasodilation and argue against the role of adenosine in functional hyperemia (Gotoh et al., 2001). It has been shown that adenosine causes vasodilation of pia vessels via A2A receptors (Morii et al., 1986; Ngai et al., 1998). It seems that the eVect of adenosine on intraparenchimal arterioles is mediated by diVerent adenosine receptors. Shi et al. (2004) have shown that activation of A2B receptors contributes to functional hyperemia during whisker stimulation.

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Adenosine is derived from ATP. There are a variety of ectonucleotidases that are responsible for the hydrolysis of ATP to AMP. AMP can then be converted to adenosine via a 50 ‐nucleotidase (Zimmermann, 1996). Adenosine has been proposed as a regulator of CBF because of the rapid rise of adenosine in the brain during hypoxia, hypotension, ischemia, and seizures (Phillis, 1989, 2004; Winn et al., 1979, 1980, 1981). Conditions in which the phosphorylation potential of cells is decreased, such as hypoxia, lead to increased level of 50 ‐AMP and subsequently adenosine accumulation. Thus, in response to hypoxia, suYcient concentrations of adenosine are achieved in the extracellular space to dilate pial vasculature and increase blood supply (Zimmermann, 1996). Moreover, ATP can be released from neurons and glial cells (Cotrina et al., 1998b; Wang et al., 2005). ATP is released as a coneurotransmitter via synapses, where it activates purinergic receptors (Zimmermann, 1996). Released ATP can also be metabolized to adenosine, which is a potent inhibitory neuromodulator (Cunha, 2001; Trussell and Jackson, 1985; Zimmermann, 1996). It is well known that glia can modulate neuronal activity by the release of glutamate (Araque et al., 2001; Haydon, 2001). Some studies suggest that ATP released from glia can also contribute to neuronal modulation (Cotrina et al., 2000; Newman, 2001). ATP released from glia binds neuronal adenosine receptors and inhibits neurons in the retina (Newman, 2003). Further, Mendoza‐Fernandez demonstrated in brain slice preparation that ATP inhibits the glutamate synaptic release via P2Y receptors in pyramidal hippocampal neurons (Mendoza‐Fernandez et al., 2000). As mentioned before, ATP is also released from astrocytes through gap junctions into extracellular space, where it binds P2Y receptors on adjacent astrocytes. P2Y receptors act via G‐protein to activate PLC, leading to the formation of IP3 with mobilization of intracellular Ca2þ. Thus, ATP acts as a mediator of a long‐range Ca2þ signaling between astrocytes and contributes in the coupling of neuronal activity and CBF. 4. Prostaglandins It has been shown that astrocytes in culture release PGE2, an important vasodilator, in a ‘‘pulsatile’’ manner, as a reaction to glutamate‐mediated Ca2þ elevations (Zonta et al., 2003b). Further, Zonta et al. (2003a) have shown in rat cortical slices that stimulation of mGluR triggers Ca2þ elevations in astrocyte endfeet that are in contact with arterioles and that this produces vasodilation. Antagonists of mGluR inhibited these Ca2þ elevations in astrocytes and impaired the vasodilative response. In this study, the authors proposed that astrocytic Ca2þ elevations elicited release of vasoactive agents, most likely a COX product that leads to an increase in CBF. This hypothesis is based on the evidence from cell cultures (Bezzi et al., 1998) and

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in vivo experiments, which reported that COX inhibitors attenuated the CBF response during neuronal activation (Bakalova et al., 2002; Golanov and Reis, 1994). Further, activation of perivascular astrocytes by patch pipette triggered vessel dilation by prostaglandins derived from AA. In brief, glutamate‐mediated Ca2þ increase in astrocytes activates PLA2, leading to mobilization of AA and production of prostaglandins by COX pathway. There are several isoforms of COX, but COX‐2 seems to be the major isoform contributing to functional hyperemia (Niwa et al., 2000). COX‐2 is present in dendritic and terminal processes of neurons close to perivascular astrocytes (Wang et al., 2005), and its expression in astrocytes is very low. Moreover, in vivo experiments show that the CBF increase to whisker stimulation is attenuated (40–50%) by COX‐2 inhibitors or in COX‐2 null mice (Niwa et al., 2000), whereas COX‐1 inhibitors did not alter CBF response to whisker stimulation (Niwa et al., 2001). Altogether, these data indicate the important role of astrocyte‐derived COX‐2 metabolites in functional hyperemic response of the brain. 5. Epoxyeicosatrienoic Acid Ellis has demonstrated that EETs are potent dilators of cat and rabbit cerebral arterioles in vivo (Ellis et al., 1990). EETs are products of the enzyme P450 epoxygenase that metabolizes AA to EETs (Amruthesh et al., 1993; Murphy et al., 1988). Astrocytes express P450 2C11 cDNA that is a homologue of liver epoxygenase (Alkayed et al., 1996b). Previous studies reported that glutamate can induce release of AA from astrocytes (Stella et al., 1994a,b). Data from our laboratory show that glutamate stimulation of astrocyte cultures produced a threefold increase in EETs production and that increase was inhibited by miconazole, an inhibitior of P450 epoxygenase activity. Also, P450 2C11 protein expression was increased in astrocyte culture after 12 hours of treatment of glutamate (Alkayed et al., 1997). Further, inhibition of P450 epoxygenase by miconazole attenuated the increase of CBF elicited by glutamate (Bhardwaj et al., 2000). Miconazole also decreased baseline CBF by 30% (Alkayed et al., 1996a), but the same result could not be reproduced in another study (Peng et al., 2002). However, Lovick demonstrated in brain slices that EETs could be involved in the vasodilating eVect produced by astrocytes and regulation of basal CBF (Lovick et al., 2005). Addition of miconazole produced a decrease in diameter of the vessel and an increase in the frequency of vasomotion, suggesting that EETs contribute to the maintenance of cerebral vascular dilator tone and functional hyperemia. Briefly, glutamate released into the synaptic cleft activates PLC to release DAG, which by the action of DAG lipase releases AA in astrocytes (Chuang et al., 1993). The released AA can be metabolized by COX to prostaglandins and by P450 epoxygenase to EETs. There are four isomers of EETs:

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5,6‐, 8,9‐, 11,12‐, and 14,15‐EETs. 11,12‐ and 14,15‐EETs are the most stable and the most potent. EETs released from astrocytes can activate Kþ channels in vascular SMC (Fig. 1), thereby enhancing an outward Kþ current and hyperpolarizing the cell, which leads to vasodilation and an increase in CBF

Hyperpolarization = Dilation Depolarization = Constriction

Presynaptic

Postsynaptic

Capillary Glutamate

PIP2

Flow

Neuron

PLC

Astrocyte

DAG DAG lipase

IP3

AA Arteriole

Ca2+

P450 2C11

PLA2 AA

? Hyperpolarization

EETs COX K+ PGF2 TXA2

PGI2 PGE2

VSMC Hyperpolarization

Depolarization Ca2+

K+ VSMC

K+

Figure 1 Glutamate released from synaptic terminals of neurons during increased neuronal activity can bind glutamate receptors on astrocytes, thus increasing intracellular calcium and AA release from astrocytic membranes by activation of PLC, PLA2, and DAG lipases. The free AA thus released can be metabolized by COX to prostaglandin (PG) F2, thromboxane A2 (TXA2), PGI2, and PGE2 and by P450 epoxygenases to EETs. DiVusion of these eicosanoids from astrocytic foot processes onto cerebral arterioles leads to modulation of cerebral vascular smooth muscle cell (VSMC) membrane potential and thus contractile state. Depolarization by PGF2 and TXA2 leads to constriction, whereas hyperpolarization by PGI2, PGE2, and EETs leads to dilation and increases in local CBF. This anatomic‐metabolic arrangement of neurons, astrocytes, and cerebral arterioles may be the basis for coupling of neuronal activity to increases in blood flow. PIP2 indicates phosphatidylinositol 4,5‐biphosphate; IP3, inositol triphosphate.

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(Alkayed et al., 1996b; Amruthesh et al., 1992; Gebremedhin et al., 1992; LeZer and Fedinec, 1997). The precise mechanism how EETs act on Kþ channels in vascular SMC is still unknown. Some studies indicate that small‐ molecular‐weight G‐proteins are involved (Li et al., 1999). It is also important to note that EETs may be stored in astrocytic membranes and released by activation of phospholipases to glutamate stimuli (Shivachar et al., 1995). Apart from the action of EETs on Kþ channels, it has been shown that astrocytes participate in angiogenesis both in vivo (Suarez et al., 1994) and in vitro (Laterra et al., 1990). However, EETs mitogenic eVect is also well known in renal epithelial and glomerular mesanglial cells (Chen et al., 1998, 1999; Harris et al., 1990; Homma et al., 1993). In 1993, Laterra described that cerebral endothelial cells form capillary‐like formations only when cocultured with astrocytes (Laterra et al., 1990). Therefore, it appears that astrocytes by release of some soluble factor may induce capillary endothelial cell tube formation. Indeed, data from our laboratory show that astrocyte‐ conditioned culture media stimulated proliferation of cerebral microvascular endothelial cells in culture. Also, formation of capillary tubes in the cultured astrocytes and endothelial cells was blocked by 17‐ODYA, inhibitor of P450 enzymes (Harder et al., 2002; Munzenmaier and Harder, 2000). 17‐ODYA is a potent inhibitor of P450 epoxygenase and !‐hydroxylase in higher dose. These data together suggest that EETs produced from astrocytes are involved in the process of tube formation. The finding that tube formations were only formed in cocultures where there is physical contact of astrocytes and endothelial cells, while astrocyte‐conditioned media only had a mitogenic eVect on endothelial cells indicate that the other factors besides EETs are also involved in angiogenesis. It has been demonstrated that the chronic neuronal activation in one region (e.g., during exercise) increases microvessel density in that area (Isaacs et al., 1992; Tuor et al., 1994). Thus, considering these data, it would be expected that chronic release of EETs in one region due to increased neuronal activity would induce capillary formation (Harder et al., 2002). In that case, functional hyperemia would be the regulator of capillary density. Accordingly, the areas with higher capillary density would receive more blood per unit time and, therefore, more eYciently couple blood flow to increase a metabolic need of activated neurons.

V. Summary Astrocytes are no longer regarded as just supportive structures among the brain cells, but play multiple roles in neuronal energy metabolism, synaptic function, pH, water homeostasis, and production of antioxidants in the brain. Despite a growing number of studies confirming the role of astrocytes in neurovascular coupling, the mechanism is still not fully understood. It is

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now clear that neuronal activity induced Ca elevations and Ca2þ waves are spreading between neighboring astrocytes through extracellular ATP signaling and gap junctions, causing release of gliotransmitters, such as glutamate and ATP, from astrocytes that can in turn modulate neuronal activity and synaptic transmission. In addition, glutamate release/reuptake from astrocytes elicits Naþ waves and subsequently glycolysis, thereby connecting neuronal activity to energy metabolism. Simultaneously, astrocytic Ca2þ elevations propagate to endfeet processes and suppress Ca2þ oscillations and vasomotion in nearby arterioles. Further, the G‐protein–coupled receptor‐mediated Ca2þ elevations in astrocytes induce release of diVerent vasoactive substances that mediate CBF responses to neuronal activation. It is also possible that additional signaling molecules from neurons (e.g., Kþ or NO) can activate astrocytes or vascular SMC directly. It has been shown that astrocytes release a number of vasoactive substances such as NO, prostaglandins, EETs, and adenosine, making them potential candidates for mediating neurovascular coupling. In conclusion, multiple factors and signaling pathways contribute to functional hyperemia and show that astrocytes have a central role in this process. Developing a detailed insight into mechanisms that underlie these interactions in the neurovascular unit may help us understand the pathophysiology and find treatments for diseases associated with brain microcirculation such as stroke, ischemia, migraine, and Alzheimer’s disease.

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Elastin‐Elastases and Inflamm‐Aging Frank Antonicelli,* Georges Bellon,* Laurent Debelle,{ and William Hornebeck* *Faculty of Medicine Extracellular Matrix and Cell Signaling—Reims University UMR 6198 CNRS 51095 Reims Cedex, France { Faculty of Science Extracellular Matrix and Cell Signaling—Reims University UMR 6198 CNRS 51095 Reims Cedex, France

I. Introduction A. The MMPs: Cytokines Interplay in Inflammation B. MMPs as Matrix ‘‘Decryptases’’ C. Elastic Fibers Fragmentation and Aging II. Elastic Fibers: Formation and Degradation A. Synthesis and Association (Coacervation) of Tropoelastin Molecules B. Development of Elastic Fibers C. Elastases in Human D. Mechanism of Elastin Degradation by Elastases III. Biological Activities of Elastin Peptides (EPs): The ‘‘Elastin Receptor System’’ A. EPs as Modifiers of Cell Behavior B. The S‐Gal Complex C. Cell Signaling Pathways Mediated by S‐Gal Occupancy IV. Elastolysis, Aging, and AAAs A. Elastase(s)‐Driven Elastin Fragmentation in Arterial Diseases: General Considerations B. Proangiogenic EVect of EPs C. Influence of EPs on T‐Lymphocytes Polarization V. The Elastin Connection in Melanoma A. Aging, Fibroblasts Senescence, and Cancer Progression B. EPs as Modifiers of the Dermal Stroma C. EPs and Melanoma Progression D. EPs‐Mediated Angiogenic Phenotype E. EPs and Inflamm‐Aging in Melanoma VI. Concluding Remarks References

Degradation of elastin, the main amorphous component of elastic fibers, by elastases belonging to the serine, metallo, or cysteine families leads to the generation of elastin fragments, designated as elastokines in keeping with their cytokine‐like properties. Generation of elastokines from one of the Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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longest lived protein in human might represent a strong tissue repair signal. Indeed, they (1) exhibit potent chemotactic activity for leukocytes, (2) stimulate fibroblast and smooth muscle cell proliferation, and (3) display proangiogenic activity as potent as VEGF. However, continuous exposure of cells to these matrikines, through increased elastase(s) expression with age, can contribute to the formation of a chronic inflammatory state, that is, inflamm‐aging. Importantly, binding of elastokines to S‐Gal, their cognate receptor, proved to stimulate matrix metalloproteinase expression in normal and cancer cells. Besides, these elastin fragments can polarize lymphocytes toward a Th‐1 response or induce an osteogenic response in smooth muscle cells, and arterial wall calcification. In this chapter, emphasis will be made on the contribution of elastokines on the genesis of age‐related arterial wall diseases, particularly abdominal aortic aneurysms (AAAs). An elastokine theory of AAAs progression will be proposed. Age is one main risk factor of cancer incidence and development. The myriad of biological eVects exerted by elastokines on stromal and inflammatory cells led us to hypothesize that they might be main actors in elaborating a favorable cancerization field in melanoma; for instance these peptides could catalyze the vertical growth phase transition in melanoma through increased expression of gelatinase A and membrane‐type 1 matrix metalloproteinase. ß 2007, Elsevier Inc.

I. Introduction The progressive and irreversible processes that decrease the capacity of the organism to adapt to the changing conditions of its environment were considered as one hallmark of human aging (Robert, 1999). Such definition might be extrapolated to a tissular context where modification of cell microenvironment with aging can perturb tissue homeostasis and organ function. Inflammation, a condition that highly modifies such environment, has essentially a protective function for organism (Larsen and Henson, 1983), but its chronicity is associated with most age‐associated diseases as athero‐ arteriosclerosis, myocardial infarction, or type II diabetes for instance (McGeer and McGeer, 2004). Chronic inflammation is characterized by the local infiltration of monocytes–macrophages and lymphocytes which release a myriad of mediators, that is arachidonic metabolites, cytokines, chemokines, vasoactive polyamines, reactive oxygen species (ROS), and reactive nitrogen species (RNS) (Sarkar and Fisher, 2006). These mediators display multiple biological functions, among them a capacity to influence the expression of matrix metalloproteinases (MMPs), key modifiers of the cell microenvironment. Indeed, matrix proteolysis is a hallmark of inflammation that

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can ultimately progress to chronic inflammatory disease (Nelson and Melendez, 2004).

A. The MMPs: Cytokines Interplay in Inflammation MMPs belong to the subfamily M10—clan MB of the zinc endopeptidase family in the MEROPS database (http://www.merops.sanger.ac.uk/). This family comprises at least 23 members which display structural homologies. Most MMPs contain a propeptide domain (average length: 80 amino acids), a zinc catalytic domain (average length: 170 amino acids), a ‘‘hinge’’ region, and a hemopexin‐like domain (average length: 200 amino acids); some MMPs as matrilysin (MMP‐7) lack hinge region and hemopexin domain, while gelatinases (MMP‐2 and MMP‐9) possess three fibronectin‐ type II modules adjacent to the catalytic domain. Within the active site, Zn2þ coordinates with three histidines present in the HEXXHXXGXXH consensus sequence. All MMPs are synthesized as zymogens and latency of enzymes is governed through Zn2þ coordination with a ‘‘cysteine switch’’ motif PRCGXPD in the propeptide domain (Ala‐aho and Kahari, 2005; Nagase and Woessner, 1999; Nagase et al., 2006). Most MMP genes, except MMP‐2, are inducible and in many instances ROS were described to enhance MMP expression. Ras, MAPKs, and PI3K, which have been involved in signaling pathway leading to MMP upregulation, are ROS targets (Fisher and Voorhees, 1998); alternatively, ROS might inhibit tyrosine phosphatases and also modulate the activity of several transcription factors such as AP‐1 and Ets involved in MMP regulation (Westermarck et al., 2001). In addition, oxidants such as HOCL and ONOO-, produced during chronic inflammation, can react with the critical cysteine in MMP prodomain and switch‐oV enzyme latency (Peppin and Weiss, 1986). MMP expression is regulated by several cytokines in vitro and in vivo (Mauviel, 1993). Particularly, interleukin‐1
(IL‐1
) and tumor necrosis factor‐ (TNF‐), which both activate the NF‐B pathway, were described to enhance MMP expression in many cell types (Deschamps and Spinale, 2006; Leppert et al., 1995a; Vaday and Lider, 2000). It needs to be emphasized that up‐ or downregulation of one MMP by a given cytokine might depend on the cell type; also, ‘‘proinflammatory’’ cytokines or ‘‘anti‐ inflammatory’’ can be associated with either an increase or a decrease of MMP expression (Shimizu et al., 2005). The cytokine‐ and chemokine‐mediated influence on MMP production may be counterbalanced or exacerbated by a direct eVect of these proteases on the degradation or processing of these mediators, respectively (McCawley and Matrisian, 2001; Somerville et al., 2003). Proteolytic processing of proIL‐1
to an active form has been reported to be induced by MMP‐2, MMP‐3, and MMP‐9 (McCawley and Matrisian, 2001); similarly, proTNF‐ can be

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converted to active cytokine by MMP‐9 and proteinase‐3, a neutrophil protease (Robache‐Gallea et al., 1995). On the contrary, neutrophil elastase can degrade and inactivate TNF‐ and all three neutrophil elastases inactivate IL‐6, another proinflammatory cytokine (Pham, 2006). Chemokines equally mediate chronic inflammation. MMPs were shown to cleave an N‐terminal tetrapeptide from MCP‐1, MCP‐2, MCP‐3, and MCP‐4, leading to molecules with reduced agonistic activity and antagonistic eVect. However, among the MMP family, matrilysin (MMP‐7) and gelatinase B (MMP‐9) were totally devoid of any degrading activity toward MCPs (McQuibban et al., 2000, 2002). Besides, N‐terminal truncation of CXC‐chemokine ligand (IL‐8) by MMP‐9 generates a variant which displays 10‐ to 27‐fold higher neutrophil chemoattracting activity (Opdenakker et al., 2001).

B. MMPs as Matrix ‘‘Decryptases’’ Matrix has not to be considered as a passive template during inflammation but, instead, together with cytokines and MMPs, acts as an active partner which may dictate the duration of the injury insult (Mott and Werb, 2004). First, matrix serves as a specialized ‘‘reservoir’’ for cytokines and growth factors. TNF‐ and IL‐2 interact avidly with fibronectin and laminin. IL‐4 as well as IFN‐, MIP‐1
, and RANTES bind to heparan side chain of proteoglycans (Vaday and Lider, 2000). Either neutral endopeptidases as MMPs or heparanase may release these matrix‐bound cytokines, which locally can modulate the phenotype of adjacent cells, or can amplify the localized inflammation by further directing neutrophils attraction (McCawley and Matrisian, 2001). Moreover, MMPs can reveal cryptic sites in these matrix macromolecules (Schenk and Quaranta, 2003). Such ‘‘decryptase’’ activity may ultimately result to the liberation of matrix fragments displaying biological activity which can be distinct from their unproteolyzed parent macromolecule (Hornebeck et al., 2002). It is now recognized that proteolysis of any matrix constituent may give rise to such fragments, we designated as ‘‘matrikines’’ in keeping with their cytokine‐like properties (Hornebeck and Maquart, 2003). Matrikines possess specific properties and cytokines, growth factors, integrins, and G‐protein‐linked receptors have been described to act as matrikine receptors. In some instances, as for tenascin‐C which contains 14 tandem EGF‐like repeats, or elastin (see below), matrikines are presented as multiple motifs to their receptor. Also, the mode of presentation of the matrikine may dictate its influence in modifying cell behavior. As one example, endostatin in its immobilized form accelerates endothelial cells migration, whereas it exhibits an inhibitory eVect when used in a soluble form (Quaranta, 2002; Tran et al., 2005). A series of matrikines have been identified as antiangiogenic molecules; they are generated from proteolysis of type IV collagen (1 chain: arresten;

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2 chain: canstatin; 3 chain: tumstatin), type XVIII collagen (endostatin), perlecan (endorepellin), fibronectin (anastellin) (Nyberg et al., 2005). These compounds proved to interfere with tumor progression in various animal models and some of them entered clinical trials (Clamp and Jayson, 2005). Other matrikines, in turn, as those originating from SPARC or fibronectin proteolysis have proangiogenic activity (Bellon et al., 2004). Although most studies focused on matrikines interaction with either endothelial cells or cancer cells, it has to be delineated that matrix fragments, from fibronectin, laminin, or collagen, have neutrophil chemotactic activity (Proctor, 1987; Weinberger et al., 2005). By contrast, 185–203 peptide from 3 chain of type IV collagen, for instance, with potent antiangiogenic activity, can also inhibit O2˙
and degranulation of leucocytes treated with forskolin or dibutyryl‐cAMP (Monboisse et al., 1994). Modification of cell environment by this interplay between MMPs, cytokines, and matrikines would probably influence the duration of a chronic inflammatory state and aging. C. Elastic Fibers Fragmentation and Aging Physiological functions decline at highly variable rates with age but, strikingly, elastic functions decline rapidly (Pasquali‐Ronchetti and Baccarani‐ Contri, 1997). Most importantly, several age‐related diseases aVect organs with high elastin content as vessels, lungs, and skin (Robert, 1999). The main amorphous component of elastic fibers, that is elastin, is a remarkable long‐ lived protein; measurements of aspartic acid racemization or prevalence of nuclear weapons‐related 14C indicated that human lung elastin longevity averaged human lifespan (Shapiro et al., 1991). Although elastin turnover may diVer among elastic tissues, it is now widely accepted that elastolysis plays a dominant function in observed alteration of elastic fibers with age. Indeed, fragmentation of elastic fibers is a hallmark of skin and arterial wall aging, and is concomitant with increased elastinolytic activity in the corresponding tissues. We will here document that generated elastin fragments are potent modulators of chronic inflammation, thus influencing arterial wall diseases particularly abdominal aortic aneurysms (AAAs). Action of these matrikines on stromal cells may equally create a favorable ‘‘cancerization field’’ for melanoma progression.

II. Elastic Fibers: Formation and Degradation A. Synthesis and Association (Coacervation) of Tropoelastin Molecules Phylogenetic studies revealed that elastin was present in all vertebrate species. It is distributed unequally within elastic tissue, representing 2–4% of dry weight in skin to up to 40% in large arteries (Rosenbloom et al., 1993).

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The 45‐kb human elastin gene, as a simple copy, is localized to chromosome 7q11.1–21.1 and consists of small exons interspersed between large introns (coding ratio 1:20) (Fazio et al., 1991). Hydrophobic and hydrophilic (cross‐ linking) domains of tropoelastin are encoded by separate exons. Elastin is synthesized as tropoelastin, a 70‐kDa polypeptide encoded by a 3‐ to 5‐kb mRNA; important alternative splicing of the primary transcripts, occurring in a cassette‐like fashion, is one characteristic of elastin gene aVecting either cross‐linking or hydrophobic domains (Indik et al., 1987; Yeh et al., 1989). Exon 26A, which corresponds to an atypical hydrophilic sequence in tropoelastin, is always spliced out in human skin fibroblasts while being always included in terminally diVerentiated keratinocytes (Hirano et al., 2001). No canonical TATA box could be evidenced in the human elastin gene and diVerent sites of transcription initiation were observed. Analysis of the 50 ‐flanking region of the human elastic gene has shown the presence of several putative cis‐regulatory elements as multiple SP‐1‐ and AP‐2‐binding sites as well as AP‐1 and cAMP repressive elements (Kahari et al., 1990). Recent data indicated that the 50 end of the first exon of the tropoelastin gene could regulate important transcriptional activity in fetal rat lung fibroblasts (Pierce et al., 2006). Implication of these cis‐elements in gene expression is dependent on cellular stimuli. For instance, IL‐1
and TNF‐ were found to upregulate and downregulate, respectively, elastin gene expression at the transcriptional level (Kahari et al., 1992a; Mauviel et al., 1993). Posttranscriptional mechanisms might be equally invoked in the regulation of elastin expression as documented for 1,25‐dihydroxyvitamin D3, phorbol esters, or tumor growth factor‐
(Kahari et al., 1992b; Parks et al., 1992). It is now recognized that elastogenic cells as human skin fibroblasts loose their ability to synthesize elastin when they are serially passaged in culture and reach senescence (Sephel and Davidson, 1986; Shelton et al., 1999). However, it was demonstrated that, contrary to the steady state level of elastin mRNAs which dropped down with dermal fibroblast passage, elastin pre‐mRNAs amounts remained invariant (Swee et al., 1995). Tropoelastin is unique among matrix macromolecules, as it can undergo coacervation, a self‐association process, which appears to be an inverse temperature transition. Contrary to heat denaturation, coacervation of tropoelastin molecules by increasing the temperature results in the formation of organized filamentous structures (Miao et al., 2003; Toonkool et al., 2001). An investigation demonstrated that tropoelastin coacervation gives rise to the formation of droplets of average size equal to 5 mm (Clarke et al., 2006). In the presence of lysyl oxidase, the enzyme catalyzing the formation of elastin cross‐links, that is desmosimes, these droplets could be fused further forming elastin remnants. Coacervation of elastin cannot be considered as a random phenomenon and arises from specific interactions between hydrophobic domains of tropoelastin (Clarke et al., 2006). Domain 26 of tropoelastin

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consisting of four nonapeptide repeats of the sequence XGXGXGVPG was described to play a pivotal function in coacervation process (Toonkool et al., 2001). Also, domain 20 appears to confer greater propensity for coacervation of elastin‐derived polypeptides (Miao et al., 2003).

B. Development of Elastic Fibers Coacervation of tropoelastin is dependent on several factors including pH, ionic strength, and also concentration of individual monomers. As demonstrated for other matrix constituents, such as collagen, laminin, or fibronectin (Kozel et al., 2006), an active participation of cell in directing elastic fiber formation was evidenced using a cDNA construct encoding bovine tropoelastin in‐frame with a timer reporter. Elastin globules, reminiscent of coacervation droplets, are formed on the cell surface and further transferred and incorporated into preexisting elastic fibers. The participation of elastin‐ binding proteins, at specialized assembly sites at the cell plasma membrane, as integrin v
3, elastonectin, elastin‐binding protein, that is S‐Gal (or EBP) as well as its Neu‐1‐associated partner, or cell surface proteoglycans, are probably required to achieve the proper orientation and concentration of tropoelastin for coacervation and further cross‐linking by lysyl oxidase (Hinek, 1996; Hinek et al., 2006; Hornebeck et al., 1986; Rodgers and Weiss, 2004). For decades, it was recognized that elastic fiber consisted of an elastin core surrounded by a mantle of microfibrils which originally served as a template for fiber formation (Frances and Robert, 1984). More than 30 distinct molecules were reported to be part of the elastic fiber architecture and identified as microfibrillar molecules, present either at the elastic fiber interface or participating to its formation (Kielty et al., 2002). Several data argue for the pivotal importance of fibrillin‐1, MAGP‐1, and fibulin‐5 in the development of elastic fibers (Kozel et al., 2003; Yanagisawa et al., 2002). Fibrillin‐1 mutations are linked to Marfan syndrome, a heritable disease characterized notably by main aortic defects associated with disorganized elastic fibers. Recently, a transglutaminase cross‐linked between tropoelastin and a central fibrillin fragment was evidenced (Rock et al., 2004). However, somewhat paradoxically, inactivation of the gene encoding fibrillin‐1 had only minimal eVect on elastic fiber formation but were involved in tissue homeostasis (Pereira et al., 1997). On the contrary, fibulin‐5 KO‐mice develop important elastinopathy with intense disorganization of elastic fibers (Yanagisawa et al., 2002). Probably, the distinct pattern of elastic fibers in arterial wall, where they consist in sheet‐like lamella, and skin in which elastic fibers consist of a continuous array of oxytalan, elaunin, and mature elactic fibers implies qualitative and quantitative diVerences in molecules directing their vectorial formation.

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C. Elastases in Human The term elastases refers to endopeptidases able to degrade insoluble elastin and thus to release soluble elastin fragments (Bieth, 1986). The first elastase was isolated from porcine pancreas which contains two distinct elastinolytic proteases (Balo and Banga, 1950). Human pancreas only expresses one elastase named pancreatic elastase II, a serine protease of the chymotrypsin family (Bieth, 1986). Silencing of the human elastase I (EC.3.4.21.36) gene during evolution in pancreatic acinar cells was attributed to mutation that inactivates enhancer and promoter elements (Rose and MacDonald, 1997). Recently, however, elastase‐1 expression at the mRNA level was evidenced in human cultured primary keratinocytes (Talas et al., 2000) and some 30 years ago, we isolated a serine elastase from aorta smooth cells which exhibited immunologic cross‐reactivity with pancreatic elastase‐1 (Hornebeck et al., 1975). In response to injury, polynuclear leukocytes and macrophages can mobilize an army of elastases which can belong to serine (clan SA, family S1), metallo (clan MB, family M10), and cysteine (clan CA, family C1) families of proteases (Table I). Azurophilic granules of polymorphonuclear leukocytes store three serine elastases named leukocyte elastase, cathepsin G, and myeloblastin (also known as proteinase‐3), whose physiological function lies on the participation in the killing and digestion of bacteria (Bangalore et al., 1990; Bieth, 1986; Hoidal et al., 1994). Besides elastin, PMN serine elastases degrade several matrix and nonmatrix substrates. Particularly, the potentiality of myeloblastin to activate TNF‐ and latent form of TGF‐
as well as its ability to promote IL‐8 expression by endothelial cells is highly suggestive of a putative proinflammatory function (Berger et al., 1996; Robache‐Gallea et al., 1995). Their concentration within leukocytes may be as high as 3 mg of enzyme per 106 cells as found for elastase and in response to several stimuli, azurophilic granules are mobilized to the cell plasma membrane (Bieth, 1986). Monocytes contain only low level of elastase and diVerentiation into macrophages leads to downregulation of serine elastases and expression of metallo and cysteine elastases (Filippov et al., 2003; Reddy et al., 1995). Four endopeptidases, among the matrixin family, are elastases and strikingly, all can be produced by human macrophages. The 13‐kb human macrophage elastase (MMP‐12) contains 10 exons and 9 introns (Belaaouaj et al., 1995; Shapiro et al., 1993). It encodes for a 54‐kDa zymogen enzyme which can be processed to a 22‐kDa active form. Similar to all elastases from this family, MMP‐12 has a broad specificity; importantly, it can inactivate 1‐proteinase inhibitor and also generate angiostatin, a potent antiangiogenic peptide, from plasminogen (Gronski et al., 1997). MMP‐12 expression is upregulated by IL‐1
, TNF‐, vascular endothelial growth factor, as well as by statins which, in contrast, display anti‐inflammatory and immunomodulatory functions (Arikan et al., 2005). Human matrilysin (MMP‐7) gene comprises

Table I

Elastases in Inflammatory Cells

Chromosomal Localization

Average Molecular Weight (Active Form)

Serine elastases: Clan SA, family S1 Leukocyte elastase E.C.3.4.21.37

19 p ter

30 kDa

Polymorphonuclear leukocyte (macrophages, lymphocytes)

Cathepsin G

E.C.3.4.21.20

14q11.2

28.5 kDa

Polymorphonuclear leukocyte

Myeloblastin (proteinase‐3)

E.C.3.4.21.76

19p13.3

29 kDa

Polymorphonuclear leukocyte

11q22.2–22.3

22 kDa

Macrophage

Belaaouaj et al., 1995

11q21–q22

19 kDa

Macrophage

E.C.3.4.24.24

16q21–q13

62 kDa

Monocyte

Gaire et al., 1994; Wilson and Matrisian, 1996 Collier et al., 1991

E.C.3.4.24.35

20q12–q13

82 kDa

Macrophages (polymorphonuclear leukocytes—lymphocytes)

Collier et al., 1991; Fridman et al., 1995; Opdenakker et al., 2001

Trivial Name(s)

Enzyme Classification

Metalloelastases: Clan MB, family M 10 Macrophage elastase E.C.3.4.24.65. (MMP‐12) Matrilysin‐pump‐1 (MMP‐7) E.C.3.4.24.23 Gelatinase A ; type IV collagenase (MMP‐2) Gelatinase B ; type V collagenase (MMP‐9)

Main Producing Cell (Others)

References

Bieth, 1986; Takahashi et al., 1988; Zimmer et al., 1992 Bangalore et al., 1990; Hohn et al., 1989 Hoidal et al., 1994; Sturrock et al., 1992

(Continued )

Table I

Continued

Trivial Name(s)

Enzyme Classification

Chromosomal Localization

Cysteine elastases: Clan CA, family C1 Cathepsin L E.C.3.4.22.15

9q21.22

Cathepsin S

E.C.3.4.22.27

Cathepsin K (cathepsin 02; cathepsin X) Cathepsin V (cathepsin L2)

Average Molecular Weight (Active Form)

Main Producing Cell (Others)

References

Macrophage

Kirschke et al., 1995

1q21

28 kDa (heavy chain: 24 kDa, light chain: 4 kDa) 24 kDa

Macrophage

E.C.3.4.22.38

1q21

28 kDa

Macrophage

E.C.3.4.22–43

9q21.22

29 kDa

Macrophage

Bromme et al., 1991; Kirschke and Wiederanders, 1994 Gelb et al., 1995; Li et al., 1995 Itoh et al., 1999; Yasuda et al., 2004

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6 exons and encodes a proenzyme with the smallest Mr  28,000 among MMPs. ProMMP‐7 is then activated by various proteases to a 19,000 Mr active form. Lack of hemopexin (PEX) domain in MMP‐7 confers to the protease a weaker aYnity for TIMPs (Gaire et al., 1994; Wilson and Matrisian, 1996; Woessner, 1995). Similar to MMP‐12, MMP‐7 can cleave 1‐proteinase inhibitor (Sires et al., 1994) and its expression is mainly upregulated in lipid‐laden macrophages (Halpert et al., 1996). Contrary to other MMPs, gelatinase A is produced in a constitutive fashion by many cell types in culture. Gelatinase A gene has 13 exons and encodes a proenzyme of Mr  72,000. Activation of gelatinase A zymogen is also unique since it requires the formation of a ternary complex between progelatinase A, one molecule of its natural inhibitor, that is TIMP‐2, and one molecule of its activator, that is membrane‐type matrix metalloprotease‐1 (MT1‐MMP). Also, either heparin or elastin can serve as templates to increase local concentration of enzyme further leading to an autoactivation process (Hornebeck et al., 2002). Although classification of gelatinase A as an elastase is undeniable when insoluble elastin is used as an in vitro substrate (Emonard and Hornebeck, 1997), degrading activity of this enzyme toward skin elastic fibers is much less important (Berton et al., 2000). On the contrary, gelatinase B (MMP‐9), which may be liberated from any inflammatory cell type, was found to hydrolyze human skin elastic fibers more potently than human leukocyte elastase (Berton et al., 2000). This enzyme occupies a central role in inflammation, as being a potent regulator of cytokine and chemokine function and may be considered as one main proinflammatory mediator. Importantly, gelatinase B knockout mice are resistant to induced bullous pemphigoid, cancer cell invasion, postischemic cardiac rupture, AAAs, and experimental autoimmune encephalomyelitis (Opdenakker et al., 2001). The contribution of cysteine proteases in elastolysis was first neglected, considering that these enzymes exert their activity optimally at lower pH (5.5–6.5) than serine or metalloelastases; furthermore they are stable only for a limited period of time under physiological conditions. However, activation of monocyte‐derived macrophages proved to stimulate the expression of vacuolar‐type Hþ‐ATPase components, leading to acidification of the pericellular space (Reddy et al., 1995). Macrophages express four distinct cysteine elastases. Cathepsin L, as the first described elastolytic protease, is synthesized as a preproprotease (M  37,500) and its activation may involve autocatalysis or limited proteolysis by cathepsin D or metalloendopeptidases. As other members of this family (Kirschke et al., 1995), cathepsin S, contrary to other cysteine elastases, displays proteolytic activity at neutral pH. The preproenzyme (M  37,479), following removal of the signal peptide in the endoplasmic reticulum, is targeted to lysosomes via the mannose‐6 phosphate receptor and transformed to mature enzyme by autocatalysis

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(Kirschke and Wiederanders, 1994). Cathepsin K is a potent elastinolytic enzyme. It is produced by activated macrophages (Punturieri et al., 2000) but is mainly expressed by osteoclasts where it actively mediates bone resorption (Gelb et al., 1995; Li et al., 1995). Cathepsin V, formerly named cathepsin L2, was shown to exhibit the highest elastolytic activity yet described among mammalian elastases (Itoh et al., 1999; Yasuda et al., 2004). Of note, an investigation indicated that caspases 2, 3, and 7 present on the plasma membrane of apoptotic smooth muscle cells could degrade elastin (Cowan et al., 2005).

D. Mechanism of Elastin Degradation by Elastases Matrix degradation, as occurring during inflammatory processes, is now attributed mainly to pericellular proteolysis where cells can use plasma membrane‐associated proteases or endopeptidases in sequestered microenvironments (Hornebeck et al., 2002). In vivo, elastin remnants with mottled appearance have been frequently observed at macrophage, smooth muscle cells, fibroblast, or cancer cells vicinity (Ntayi et al., 2004). In vitro, contact between monocyte‐derived macrophages and elastin was found to be a prerequisite for elastinolytic activity (Reddy et al., 1995). Such requirement for elastolysis to occur might have several origins: (1) Cell–elastin interface is rapidly acidified by a vacuolar‐type Hþ‐ATPase, creating conditions of activity of cysteine elastases in specialized microenvironmental compartments (Reddy et al., 1995). (2) Cosegregation of cysteine, metalloelastases, but also components of the plasmin (ogen) system may be favorable for induction of protein cascades (Woessner and Nagase, 2000). (3) Activated proteases might display synergistic eVect against elastin as observed for leukocyte elastase and cathepsin G (Boudier et al., 1991). (4) Elastases from one family can locally inactivate physiological inhibitors of elastases from other families, thus increasing delay time of inhibition and quantum elastolytic events (Campbell et al., 2000; Johnson et al., 1986; Liu et al., 2000; Okada et al., 1988; Overall and Dean, 2006). Furthermore, elastase bound to elastin might be partially refractory to inhibition by its naturally occurring inhibitor (Hornebeck and Schnebli, 1982). (5) Finally, locally produced elastin fragments act as amplifiers of elastolysis (see below). The first step in elastase‐mediated degradation of insoluble elastin by serine elastases, that is pancreatic and leukocyte elastases or metalloelastases as gelatinases, consists in a rapid adsorption of enzyme onto its substrate (Hornebeck et al., 1984). For leukocyte elastase, interaction between carboxylate groups of elastin and clusters of positive charges in enzyme structure are involved in binding. In such adsorbed state, elastase(s) cannot freely diVuse from its substrate and elastin is preferentially hydrolyzed by preadsorbed

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enzyme (Hornebeck et al., 1984). Gelatinase(s) also bind to elastin, although much less eYciently as leukocyte elastase (Emonard and Hornebeck, 1997). The presence of fibronectin type II‐like repeats inserted in tandem in the catalytic domain of both enzymes are necessary for interaction and elastin‐ degrading ability (Emonard and Hornebeck, 1997). Activation energies of elastin degradation by human leukocyte elastase, matrilysin, and gelatinase B, deduced from Arrhenius plots, are equal to 13.6, 8.4, and 11 kcal/mol, respectively (Mecham et al., 1997). Activation energy of elastolysis by porcine pancreatic elastase lies in the same range: 8.2 kcal/mol, close to values obtained using hydroxyl anions (0.1‐N NaOH) as elastin solubilizing agent: 11.96 kcal/mol (Hornebeck et al., 1972). It indicates that energy requirements for elastolysis by elastases are similar to most enzyme‐catalyzed reaction, suggesting that scissible peptide bonds in elastin are accessible to these elastases.

III. Biological Activities of Elastin Peptides (EPs): The ‘‘Elastin Receptor System’’ A. EPs as Modifiers of Cell Behavior The transforming activity of plasmin‐generated fibronectin peptides on chicken embryo fibroblasts, which was not observed with intact fibronectin, probably represents one of the first examples of a bioactivity developed by a matricryptic fragment (De Petro et al., 1981). Parallely, in the early 1980s, it was evidenced that tropoelastin and proteolytic fragments of insoluble elastin displayed potent chemotactic activity for human monocytes and fibroblasts (Senior et al., 1984). It needs to be emphasized that the putative in vivo activity of elastin peptides (EPs) was earlier suspected from experiments showing their proarteriosclerotic eVect when injected to rabbits (Robert et al., 1971). Since, EPs obtained from alkaline (‐elastin), acid (‐elastin), or enzymatic hydrolysis of bovine or human elastins were described to modify several functions of normal cells in vitro, which are listed in Table IIA. As other matrikines, the biological eVect exerted by EPs might depend on their mode of presentation to cells. For instance, substratum‐bound EPs inhibit aortic smooth muscle cells migration, while they caused their migration when used in solution in Boyden’s chamber (Ooyama et al., 1987). Also, the site responsible of the biological eVect might not be cryptic in the elastin precursor since tropoelastin and elastin degradation fragments were found to promote the migration or proliferation of normal or transformed cells to similar extent (Jung et al., 1998). Possibly, however, such site might be hidden in vivo in elastic fibers due to the association of elastin with microfibrils (Kielty et al., 2002).

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Table IIA Biological Activities of Elastin Peptides: Derived from Organoalkaline (E), Acid (E), or Enzymatic Hydrolysates of Human or Bovine Insoluble Elastins Cell Monocytes

Lymphocytes Fibroblasts

Smooth muscle cells

Endothelial cells

EVect

References

Chemotaxis

Modification of ion fluxes ROS liberation Inhibition of cholesterol synthesis Proliferation Induction of apoptosis Chemotaxis Proliferation Chemotaxis Proliferation Modification of ion fluxes Induction of angiogenesis Vasorelaxing eVect and NO liberation

Chen et al., 1997; Jacob et al., 1987; Kamisato et al., 1997; Senior et al., 1980; Uemura and Okamoto, 1997 Jacob et al., 1987 Fulop et al., 1986 Varga et al., 1997 Poggi and Mingari, 1995 Peterszegi et al., 1999 Mecham et al., 1989 Ghuysen‐Itard et al., 1992; Kamoun et al., 1995; Tajima et al., 1997 Ooyama et al., 1987 Mochizuki et al., 2002 Jacob et al., 1987 Robinet et al., 2005 Faury et al., 1998a,b

Table IIB Biological EVects of Elastin Peptides: Derived from Defined Peptide Sequences of Human Tropoelastin Peptide Sequence

Cell

EVect

Monocyte

Chemotaxis

Fibroblast

Chemotaxis Proliferation

Smooth muscle cells Lymphocyte Endothelial cell

Proliferation Osteogenesis Th‐1 polarization Chemotaxis Induction of angiogenesis

PGAIPG LREGDPSS

Fibroblast Monocyte

Chemotaxis Chemotaxis

GFGVGAGVP

Fibroblast Endothelial cell Endothelial cell Smooth muscle cell

Chemotaxis Chemotaxis Chemotaxis Proliferation

VGVAPG

GLGVGAGVP VPGVG

References Bisaccia et al., 1998; Hance et al., 2002; Senior et al., 1984 Senior et al., 1984 Kamoun et al., 1995; Tajima et al., 1997 Mochizuki et al., 2002 Simionescu et al., 2005 Debret et al., 2005 Long et al., 1989 Robinet et al., 2005 Grosso and Scott, 1993a,b Bisaccia et al., 1998; Long et al., 1989 Long et al., 1989 Long et al., 1988 Long et al., 1988 Wachi et al., 1995

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Consequently, numerous studies were undertaken at aims to define the structure of bioactive fragments, as well as to unravel the nature of an elusive elastin receptor. A 67‐kDa elastin‐binding protein (EBP) was first isolated and described as a membrane‐associated protein with lectin‐like property (Hinek et al., 1988). Tropoelastin‐like VGVAPG sequence, which is found as several repeats in the tropoelastin sequence corresponding to exon 24 of the gene (Table III), binds avidly to EBP with KD in the nanomolar range (Mecham et al., 1989). Indeed, VGVAPG recapitulates most properties of elastin hydrolysates on cell chemotaxis or proliferation (Table IIB). Similar eVects were also observed using PGAIPG, a peptide from tropoelastin covalent structure, and LGTIPG, a peptide from laminin. EBP was then named as the elastin/ laminin receptor (Mecham et al., 1989), a designation which somewhat created moment’s confusion. Later, EBP was further identified as a spliced variant of
‐galactosidase (S‐Gal) whose matrix binding site (Privitera et al., 1998) can accommodate peptide with GXXPG consensus sequence, adopting a preferential type VIII
‐turn conformation (Brassart et al., 2001). Such motif (GXXPG) is highly represented in tropoelastin, that is 28 times, but fibrillins, molecules at the elastin–microfibrils interface or associated with elastic fibers, also contain repeats of such sequence that might display similar biological activity as VGVAPG (Booms et al., 2006; Hornebeck et al., 2005) (Table III). However, elastase‐driven proteolysis of these matrix constituents might generate or degrade these biological fragments. In recent investigations, the VGVAPG motif could be obtained following hydrolysis of a human elastin exon 24‐encoded product by myeloblastin or leukocyte Table III

GXXPG‐Containing Matrix Macromoleculesa

Tropoelastin Exon 24: GLVPG VGVAPG VGVAPG VGVAPG VGLAPG VGVAPG VGVAPG VGVAPGI Molecule Tropoelastin Fibrillin‐1 Fibrillin‐2 Fibrillin‐3 Emilin‐1 Emilin‐2 Collagen VIII Fibronectin Collagen III 1 a

Location Within Elastic Fibers

Number of GXXPG Sequences

Core Microfibrils Microfibrils Microfibrils Elastin–microfibril interface Elastin–microfibril interface Some elastic fibers At vicinity At vicinity

28 3 7 4 3 3 3 7 3

Adapted from Booms et al. (2006), Hornebeck et al. (2005), and Kielty et al. (2002).

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elastase (Lombard et al., 2005, 2006), but other elastases with distinct specificity as MMP‐12 might have an opposite eVect (Barroso et al., 2006). As mentioned in Table IIB, other elastin sequences could promote cell chemotaxis, suggesting that elastin might possess several cell receptors; particularly, the ability of the hydrophilic LREGDPSS from exon 26A to induce monocyte chemotaxis supports this contention (Bisaccia et al., 1998).

B. The S‐Gal Complex The elastin receptor complex, or S‐Gal complex, is derived from the lysosomal
‐galactosidase complex. It is expressed at the surface of numerous cell types (Hinek et al., 1988) and comprises a peripheral 67‐kDa subunit which actually binds elastin, that is S‐Gal (formerly EBP), a 55‐kDa protective protein/cathepsin A (PPCA), and a 61‐kDa membrane‐bound neuraminidase, Neu‐1 (Mecham et al., 1989). The S‐Gal complex is both an actor of elastin synthesis (Hinek et al., 2006) and a receptor for its fragments. The S‐Gal subunit is devoid of enzymatic activity but retains the ability to bind to
‐galactosugars such as galactose or lactose. The alternative splicing of
‐galactosidase primary transcripts consists of two deletions. The first one deletes exons 3 and 4, and introduces a frameshift; the second one occurs in exon 6 and restores the initial frame. As a consequence, S‐Gal possesses a unique 32‐residue sequence. This segment is thought to contain the elastin‐ binding site of the molecule, that is, VVGSPSAQDEASPL as antibodies directed against this region substantially block EP eVects (Mochizuki et al., 2002). The second subunit of the S‐Gal complex, PPCA, is synthesized as a 54‐kDa zymogen, and is processed to 32‐kDa and 20‐kDa subunits linked by interchain disulfide bridges (Hiraiwa, 1999). It possesses a cathepsin A activity in the lysosome. The precise role of PPCA in the S‐Gal complex is still unknown but, by analogy to its function in the lysosome, it could protect the two other subunits and thereby contribute to the structural integrity of the whole receptor complex. The last subunit of the complex, the lysosomal neuraminidase, Neu‐1, which is synthesized as a 45.5‐kDa preproenzyme, and further glycosylated and processed to a mature enzyme, cleaves the terminal N‐glycosidic sialic acid from various sialoconjugates (Hinek et al., 2006). Recent results indicate that Neu‐1 is one of the key actors of elastogenesis (Hinek et al., 2006). When Neu‐1 is phosphorylated at its C‐terminus, it is addressed to the cell surface together with PPCA so that the PPCA/Neu‐1 complex found at the cell surface is more abundant (Lukong et al., 2001). On ligand binding, S‐Gal is recruited to the cell surface where it forms the S‐Gal complex together with the membrane‐residing PPCA and Neu‐1 subunits. The occupancy of S‐Gal lectin site by galactosugars dramatically

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reduces the aYnity of its elastin site for EPs or tropoelastin and sheds S‐Gal from the complex. Consequently, lactose is commonly used to antagonize EPs eVects mediated by S‐Gal.

C. Cell Signaling Pathways Mediated by S‐Gal Occupancy An examination of the literature reveals that the signaling pathways triggered by EPs are highly dependent on (1) the considered cell type and (2) the nature of the agonist, that is, EPs, elastin hydrolysates, tropoelastin, or even elastin, as well as (3) its presentation to cells. For instance, the chemotactic eVect of EPs for monocytes does not involve a protein kinase C (PKC) (Kamisato et al., 1997; Uemura and Okamoto, 1997), whereas a similar response of M27 cell relies on the activation of such a kinase (Blood and Zetter, 1989). Likewise, EPs have been reported to promote the proliferation of smooth muscle cells (Mochizuki et al., 2002), whereas elastin has been reported to have the opposite eVect (Karnik et al., 2003). To summarize, EPs were initially shown to influence the mobilization of intracellular ions, intracellular levels of cAMP and cGMP, and their usual signaling targets, PKC, PKA, and PKG (Blood and Zetter, 1989; Fulop et al., 1986; Kamisato et al., 1997; Uemura and Okamoto, 1997). Owing to developments in this field, the general picture of EPs signaling is now slightly modified. Indeed, several studies (Debret et al., 2005; Duca et al., 2002; Mochizuki et al., 2002) highlighted that extracellular signal‐ regulated kinase‐signaling activation is a common feature of EPs signaling in normal cells (Fig. 1). Of note, analysis of increased smooth muscle cells proliferation following EPs treatment point out that phosphorylation of Erk1/2 is a critical event in the EPs‐dependent mitogenic ‘‘signaling’’ (Mochizuki et al., 2002). In these cells, the Erk pathway is the only MAP kinase cascade involved. Its activation requires a functional S‐Gal receptor, activation of G proteins, opening of L‐type channels and Ca2þ influx, activation of phosphotyrosine kinases, notably c‐Src, followed by Ras phosphorylation, Raf activation, and the initiation of the MAP/Erk Kinase. Interestingly, the authors also show that inhibition of the PDGF receptor kinase activity significantly decreased Erk1/2 phosphorylation levels, suggesting that the PDGF receptor could be transactivated via c‐Src and could trigger the Ras‐Raf‐MEK1/2‐Erk1/2 phosphorylation cascade. The activation of the MEK/Erk pathway has also been reported in dermal fibroblasts treated with EPs (Duca et al., 2005). In these cells, a strong and sustained activation of the MEK/Erk module is achieved. This, in turn, leads to the activation of AP‐1 family transcription factors and the initiation of MMP‐1 gene transcription. In contrast to what holds for smooth muscle cells, MEK activation is not Ras‐dependent. In this cell model, MEK is

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EP (VGVAPG)n Ca2+

EBP PP ?

ER

b Cai

Ga

?

Nase

g Multiple pathways (PI3K, PKA, Transactivation of RTKs,…)

Cyclins A, E, D CDK-2 CD-4

Cell proliferation

Erk1/2

NF-k B, AP-1,…

MMPs expression

Figure 1 A general scheme of signaling pathway triggered by elastin peptides (Duca et al., 2002, 2005; Mochizuki et al., 2002).

activated simultaneously by two distinct signaling cascades. On one hand, a pertussis toxin‐sensitive G‐protein dissociates, leading to the activation of p110 phosphatidylinositol‐3 kinase (PI3K). This kinase activates Raf‐1 which phosphorylates MEK. On the other hand, an increase in the intracellular cAMP level activates PKA, which in turn, contributes to the activation of another isoform of Raf, that is B‐Raf, which phosphorylates MEK. S‐Gal occupancy by EPs also leads to activation of MEK/Erk pathway in endothelial cells. However, contrary to fibroblasts, activation only requires one ‘‘linear’’ signaling cascade involving chronological activation of PI3K, AKT, eNOs, and nitric oxide (NO) release; NO, in turn, through cGMP and PKG1‐ activation will ultimately activate the MEK/Erk module (Robinet et al., unpublished data). Whether EPs‐mediated eVects involve, as documented for smooth muscle cells, transactivation of a tyrosine kinase receptor such as VEGF Receptor, that is Flk‐1/KDR, remains to be determined. Erk activation results in a series of cellular eVect; however, whatever the cell type, its activation following EPs binding to S‐Gal upregulates MMP expression. Here again, the overexpressed MMP(s)‐type depends on the cell type considered (1) MMP‐1 and MMP‐3 in fibroblasts, (2) MT1‐MMP and MMP‐2 in endothelial cells and melanoma cells, and (3) MMP‐9 in lymphocytes.

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IV. Elastolysis, Aging, and AAAs A. Elastase(s)‐Driven Elastin Fragmentation in Arterial Diseases: General Considerations The conspicuous fragmentation of human aortic elastic fibers with age was correlated, in our initial studies, with an increase of elastase activity as determined using ‐elastin as substrate (Hornebeck et al., 1976). Since enzyme levels were determined in specimens devoid of atherosclerotic lesions, we hypothesized that elastolysis of elastic fibers in the media of aged individuals could be attributed to smooth muscle cell serine elastase, whose amount increased with cell passages (Hornebeck et al., 1975). Other elastases from distinct families can equally participate in age‐associated elastic fibers breakdown, and arterial aging was associated with increased MT1‐MMP‐ mediated proMMP‐2 activation in human aorta (McNulty et al., 2005). These endopeptidases display low elastinolytic activities as compared to leukocyte elastases, a property which is consistent with the progressive degradation of arterial elastin over decades. Locally generated EPs are catalysts of major changes within the arterial wall. First, as we and other investigators demonstrated, EPs can mediate amplification of elastolysis through upregulation of smooth muscle cell elastase (Cohen et al., 1992; Hornebeck and Robert, 1977). Similarly, S‐Gal activation by elastin fragments stimulates MMP‐2 expression by aortic smooth muscle cells (Cohen et al., 1992). Fragmentation of elastic fibers and calcification of vascular elastin are two main features of arteriosclerosis. Treatment of aortic smooth muscle cells with EPs was found to enhance alkaline phosphatase activity, osteocalcin levels, and core‐binding factor ‐1 expression, an osteoclast‐ specific transcription factor (Simionescu et al., 2005). The induction of an osteogenic cellular response following elastin degradation was further confirmed in vivo using elastin implanted subdermally in female rats (Lee et al., 2006). These data suggest that elastase inhibition might be one strategy to impede aorta elastin calcification; indeed, administration of a synthetic MMP inhibitor to rats treated with vitamin D3 substantially attenuated aortic calcification (Qin et al., 2006). Binding of calcium ions to neutral sites in elastin results in progressive loss of elasticity of the polymer and consequently to rigidification of arteries; besides, it induces transconformation of elastin, which favors lipids deposition and potentiates elastolysis (Hornebeck and Partridge, 1975; Robert et al., 1984). Such auto‐sustained mechanism of elastolysis is probably a main actor in further development of chronic inflammation through monocytes recruitment by EPs. The pivotal role of inflammation in atherosclerosis (Libby, 2002) and particularly in AAAs is now well appreciated (Shimizu et al., 2004, 2006).

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AAAs were first designated as ‘‘inflammatory aneurysms’’ where inflammatory cells as macrophages, polymorphonuclear neutrophils (PMN), and T cells gain access to aortic media and intima from vasa vasorum. Although several factors can attract monocytes, Hance et al. (2002) demonstrated, using VGVAPG antibodies (BA4), that EPs were mainly responsible of monocytes chemotactic activity in human AAAs extracts (Hance et al., 2002). The presence of such S‐Gal‐binding motif peptide has been demonstrated in vascular lesions leading to the segmental destruction of aortic media (Juvonen et al., 1994), and its increased level in the blood circulation is considered as a useful indication of a high rupture risk in AAAs (Lindholt et al., 2001; Petersen et al., 2002). Besides, VGVAPG peptide induces characteristic feature of AAAs in the Anidjar/Dobrin rat model (Nackman et al., 1997). It has to be emphasized that in AAAs, the burst of elastolysis is triggered by elastases from inflammatory cells, that is neutrophil elastase, MMPs, or cathepsins, which either can act individually or in concert to degrade elastin (Longo et al., 2002). Thus, all these previous data highly suggested that elastase(s)‐driven elastin degradation is one main underlying mechanism leading to chronic inflammation within arteries and progression of AAAs. This contention is further supported by two additive biological properties exerted by these matrix fragments, namely their proangiogenic activity and their influence on T‐cell polarization. B. Proangiogenic Effect of EPs 1. Elastolysis, Angiogenesis, and Aneurysms The pathophysiology of AAAs is a complex and multifactorial process characterized by aortic wall remodeling and proteolysis of matrix proteins, immune and inflammatory responses, stress within arterial wall, genetic traits, and neovascularization (Ailawadi et al., 2003; Wassef et al., 2001). Biochemical and immunological studies on AAAs showed fragmentation of collagen and elastin fibers in both the media and adventitia, thinning of the medial wall associated with smooth muscle cell loss, abundant infiltration of macrophages and lymphocytes, and increased expression of proteinases. Proteinase upregulation was temporally and spatially associated with disruption of the aortic media and angiogenesis (Ailawadi et al., 2003; Lopez‐Candales et al., 1997; Rossignol et al., 2002; Wassef et al., 2001). AAAs are also characterized by inflammation and neovascularization. The crucial role of MMP‐2 and MMP‐9 in the pathogenesis of AAAs and elastin fragmentation has been pinpointed by Baxter’s group (Longo et al., 2002). These authors demonstrated that mice deficient in MMP‐9 or MMP‐2 expression underwent AAAs induction, indicating that macrophage‐derived MMP‐9 and mesenchymal cell‐derived MMP‐2 might work in concert to

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produce aortic aneurysms. Thus, MMP‐2 and MMP‐9 can be considered as potential therapeutic targets to impair elastin fragmentation in aneurysm development. Other elastinolytic enzymes, as for instance MMP‐12 or leukocyte elastase, may also contribute to aneurysmal progression (Boudghene et al., 1993; Busuttil et al., 1982; Cannon and Read, 1982; Nackman et al., 1997; Saito et al., 2002). Angiogenesis, the formation of new blood vessels from preexisting vasculature, consists in a series of temporally and tightly regulated events that include capillary vasodilation and increase of vessel permeability facilitating endothelial cell activation, degradation of basement membrane, and migration, proliferation, and maturation of new capillary tubes. These events are under the control of a number of factors among which are growth factors, cytokines, proteinases, mainly MMPs, extracellular matrix proteins and/or matrix‐derived proteolytic fragments as matrikines, and cell adhesion proteins (Bellon et al., 2004; Carmeliet and Jain, 2000; Folkman, 1995; Risau, 1997). Disruption of the local balance among these factors can favor the emergency of new blood vessels or can determine their quiescence or regression within a tissue (Hanahan and Folkman, 1996). Neovascularization aVecting all layers of the arterial wall in human AAAs is associated to the development and growth rate of aneurysms (Herron et al., 1991). Its intensity is closely related to the extension of inflammatory infiltrate, particularly macrophage‐rich (Holmes et al., 1995; Satta et al., 1998). It was also reported that neovascularization is an ongoing process in the mature aortic aneurysm characterized by the presence of thick‐walled microvessels in the media and adventitia, and thin‐walled capillaries in the subintima (Paik et al., 2004). For the sake of comparison, the extent of medial neovascularization is threefold higher in AAAs than in atherosclerosis (Holmes et al., 1995). A direct link between elastin fragmentation and neoangiogenesis in AAAs was demonstrated in an aneurysm model in which perfusion of pancreatic elastase or VGVAPG in rat infrarenal aorta induced adventitial angiogenesis (Nackman et al., 1997). Overall, it was therefore tempting to speculate that ongoing angiogenesis in blood vessels of AAAs can be the consequence of progressive elastin fragmentation. However, we cannot exclude the possible involvement of ongoing elastogenesis in AAAs neovascularization since human AAAs were reported to contain fourfold more macrophage‐derived tropoelastin protein than normal arteries (Krettek et al., 2003). 2. Elastin Fragments as Proangiogenic Matrikines EPs are present in the blood circulation at increased concentrations during vascular pathologies such as arteriosclerosis, acute aortic dissection, and aneurysmal and ulcerative manifestations of atherosclerosis. However, their

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level is unchanged in distal aortic occlusive disease and acute myocardial infarction, though a moderate increase was reported in patients with ischemic heart disease (BaydanoV et al., 1987; Fulop et al., 1990; Petersen et al., 2002; Shinohara et al., 2003). On the contrary, the age‐related concentrations of circulating EPs in human healthy subjects revealed discrepancies. Thus, no change, decreased or increased EPs levels were reported (BaydanoV et al., 1987; Bizbiz et al., 1997; Fulop et al., 1990). Factors such as circulating elastin autoantobodies, elastin‐binding proteins, elastin‐binding receptor, or antibody‐dependent elastin reactivity have been proposed to influence the levels of circulating EPs and to account for such discrepancies when determined by immunoassays (Sivaprasad et al., 2005; Wei et al., 1993). Earlier reports suggested that EPs can positively regulate endothelial cells and angiogenesis (Faury et al., 1995, 1998a,b; Robinet et al., 2005). Indeed, these investigations indicated that VGV‐containing EPs mediated a dose‐ and NO‐dependent vasorelaxation in rat aorta rings (Faury et al., 1998a,b, 1995) associated with an increase in cytoplasmic‐ and nuclear‐free calcium in human vascular endothelial cells (Edelberg and Reed, 2003; Faury et al., 1998a,b; Weinsaft and Edelberg, 2001). Moreover, administration of either VGVAPG or pancreatic elastase used as aneurysm models in rats proved to reproduce the main characteristics of the disease (Nackman et al., 1997). Our data indicated that elastin peptides exhibited proangiogenic activity (Robinet et al., 2005). ‐Elastin as well as VGVAPG motif‐containing peptide was found to accelerate angiogenesis in the chick chorioallantoic membrane model, as well as to stimulate both macrovascular and microvascular endothelial cells migration and tubulogenesis in the matrigel and collagen ex vivo models. These eVects are S‐Gal‐dependent and mediated through upregulation and activation of MT1‐MMP, since inhibitors and siRNAs directed against this MMP totally suppressed elastin‐mediated angiogenesis (Fig. 2). Unpublished results from our laboratory further evidenced that binding of elastin peptides to S‐Gal results in the activation of the PI3K/ AKT/eNOS/MEK/ Erk1/2 axis. Besides, upregulation of MT1‐MMP in microvascular endothelial cells was found to be triggered by elastin peptide‐ mediated NO and cGMP production. Therefore, NO may be the main conductor of elastin‐mediated angiogenesis. Besides, EPs were found to impede endothelial cell death; their function as survival factors has been recently emphasized by data from recent investigations showing their cardioprotective and regenerative functions in a model of heart ischemia‐reperfusion in rats. For instance, a decrease of heart necrosis area and creatine kinase release were observed by administration of EPs under conditions of pre‐ and postconditioning. These eVects were NO‐dependent and involved the RISK (reperfusion injury salvage kinase) pathway consisting in activation of PI3K/AKT/eNOS/Erk1/2 signaling (Robinet et al., 2007).

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Collagen model KE Control

Matrigel model KE Control

MTI-MMP protein expression KE (µg/ml) 0 0.1 10 100

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NO S-Gal PP

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sGC cGMP

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PI3−K

craf-1 MEK1/2 Erk1/2

Activation of MT1-MMP expression Figure 2 Signaling pathway mediated by the binding of VGVAPG motif on S‐Gal and angiogenesis connection. Elastin peptides as ‐elastin (KE) and VGVAPG favor in vivo and in vitro angiogenesis as shown in the chick chorioallantoic membrane (CAM), collagen, and matrigel models through upregulation of NO‐mediated MT1‐MMP. Elastin‐mediated angiogenesis involves the PI3K /AKT/eNOS / Erk1/2 axis (Robinet et al., 2005, 2007).

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3. Neovascularization and Aging in AAAs: A Paradox Numerous studies reported that angiogenesis is impaired in aged individuals, a condition that contributes to delayed wound repair, neurodegeneration, and renal disease; it is particularly detrimental during the revascularization of the ischemic heart or collateral vessel development in cardiovascular diseases (Edelberg and Reed, 2003; Weinsaft and Edelberg, 2001). Age‐related alterations in the neovascular process is the result of cellular dysfunction, dysregulation of growth factor expression and their cognate receptors, changes in MMPs and matrix proteins expression, and defective inflammatory process. Such alterations contribute to delayed and impaired neovascularization in aged tissues. Most studies, aiming at relating angiogenesis and aging, focused on in vitro and animal models under normal or pathological conditions (Reed and Edelberg, 2004). Many components and multiple steps of the neovascular response were demonstrated to be altered with age, some of them are summarized in Table IV. Besides, studies showed that the bioavailability of NO, a crucial factor for the integrity and function of endothelium, is reduced in aged endothelial cells (Haendeler, 2006). NO downregulation in aged tissue has been reported to be associated with deficiencies in the synthesis or activity of eNOS and iNOS with, loss of AKT activity, and enhanced production of ROS, especially O2
leading to peroxynitrite formation (Bach et al., 2005; van der Loo et al., 2000). Most of in vitro and in vivo studies showed that the eNOS enzyme system is defective at both the expression and activity levels (Bach et al., 2005; Matsushita et al., 2001; Rivard et al., 1999; Smith and Hagen, 2003; Tschudi et al., 1996), though age‐related increase of eNOS and iNOS expression or activity were reported in some studies (Cernadas et al., 1998; van der Loo et al., 2000). Among actors controlling angiogenesis, AKT, a kinase playing a central role in modulating endothelial cell survival through phosphorylation of both eNOS and telomerase reverse transcriptase (hTERT) subunit (Chang et al., 2002; Kim and Chung, 2002), increases with endothelial cellular senescence. Consequently, inhibition of AKT extends the life span of these cells (Minamino et al., 2004). It is now well documented that the decrease of TERT activity during the process of endothelial cell aging precedes the onset of replicative senescence that can be delayed by NO (Vasa et al., 2000). In the context of angiogenesis and development of vascular diseases, proangiogenic or proatherogenic factors as TNF‐ and oxLDL have been reported to induce TERT inactivation in endothelial cells through reduction of AKT phosphorylation (Breitschopf et al., 2001). Such an AKT‐dependent mechanism can be correlated to age‐ related loss in active plasma membrane‐bound eNOS protein which binds less eYciently to AKT (Smith et al., 2006). Overall, eNOS, AKT, and telomerase activities appear to be crucial actors in endothelial cell aging and apoptosis, and consequently in angiogenesis.

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Neovascularization and Aging References

(1) Endothelial cells dysfunction Decreased migration Reduced proliferation Defective response to VEGF Defective response to the antiproliferative eVects of TGF‐
1 Increased sensitivity to apoptotic stimuli (2) Reduced expression of growth factors VEGF

b‐FGF IGF‐1 TGF‐


Arthur et al., 1998; Mogford et al., 2002; Reed et al., 2000, 2001 Nissen et al., 1998; Rivard et al., 1999; Wang et al., 2004 Rivard et al., 1999 McCaVrey and Falcone, 1993 Chavakis and Dimmeler, 2002; HoVmann et al., 2001; Vasa et al., 2000 Arthur et al., 1998; Ferrara and Gerber, 2001; Ito et al., 2005; Nissen et al., 1998; Rivard et al., 1999; Sadoun and Reed, 2003; Wang et al., 2004 Augustin‐Voss et al., 1993; Sartippour et al., 2002; Swift et al., 1999 Trejo et al., 2004 Reed et al., 1998

(3) Elastin, MMPs, and inhibitors in aged endothelial cells Decreased sensitivity of endothelial cells to Faury, 1998; Fulop et al., 2001 elastin peptides Downregulation of MMP‐1 expression Koike et al., 2003; Reed et al., 2001 Downregulation of MMP‐2 activity Reed et al., 2000 Increased expression of TIMP‐1 and TIMP‐2 Koike et al., 2003

Thus, the observed intense neovascularization in AAAs, an age‐regulated disease, appears somewhat paradoxical. Such paradox also applies to elastin peptides influence on endothelial cells. Indeed, coupling of S‐Gal to its signaling pathway was described to decline with age. For instance, the endothelium‐mediated relaxation of aorta rings by EPs is lost in aged rats; also hearts from 24‐month‐old rats failed to respond to EPs in an ex vivo model of ischemia/reperfusion injury (unpublished data). Such ineYciency of EPs on endothelium functions with aging can be attributed to AKT/eNOs axis as described above, to S‐Gal, and particularly to Neu‐1, dysfunction, or both. Although elastin peptides are not the only proangiogenic factors that might trigger angiogenesis in AAAs, it is also conceivable that, among cells able to participate to neovascularization, circulating endothelial cells or progenitor and mesenchymal stem cells may also play a crucial role

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(Ghajar et al., 2006; Sata, 2006). Such cells can be recruited from the bone marrow after tissue injury (Goon et al., 2006). VEGF, angiopoietin‐1, FGF, SDF‐1, PIGF, and MCSF were reported to be involved in the recruitment of these cells. Indeed, endothelial cell progenitors might restore aging‐impaired cardiac angiogenic function (Edelberg et al., 2002). However, it is not known whether locally produced elastin peptides may also be involved in this process. Mesenchymal stem cells and endothelial cell progenitors possess the ability to diVerentiate into endothelial lineage cells and to contribute to vascular homeostasis. We can hypothesized that these de novo endothelial cells could possess a functional elastin receptor and might respond to elastin peptides independently of age.

C. Influence of EPs on T‐Lymphocytes Polarization 1. The Th‐1/Th‐2 Balance in Aging and Arterial Wall Diseases The cytokines network produced by T cells (CD4, CD8, NK) plays a key role in cell–cell communication and matrix regulation. Likewise, the balance regulating pro‐ and anti‐inflammatory cytokines, that is the Th‐1/Th‐2 balance, is of equally importance in host protection against infections. However, besides its involvement in developing a chronic inflammation, aging is also associated with a shift toward Th‐2 cytokines at the expense of Th‐1 cytokines in T cells (Alberti et al., 2006; Rink et al., 1998; Shearer, 1997). Accordingly, the level of the most potent inducer of cell immunity, IFN‐, declines with aging (Gardner and Murasko, 2002), whereas increase of TNF‐ is used as a marker of inflammatory diseases in aged individuals (Di Iorio et al., 2003). Within atherosclerotic lesions, the immune response is characterized by Th‐1 lymphocytic subpopulation predominance with an increase in expression of IL‐2, IL‐12, IL‐15, and IL‐18 and the absence of IL‐4, IL‐5, and IL‐10 (Frostegard et al., 1999). Accumulation of immune cells in AAAs lesions shows a large predominance of CD4þ T cells as compared to CD8þ cells (Hansson et al., 1989; Koch et al., 1990; Watanabe et al., 1995). If everyone agrees on the importance of the inflammatory reaction on AAAs formation, delineation of specific cytokines involved in AAAs progression is still controversial. Indeed, studies using animal model showed the key role of CD4þ cells and IFN‐, a Th‐1 cytokine, in the development of AAAs (Xiong et al., 2004), whereas other investigators pointed out that a Th‐2 response prevailed in aneurysms progression characterized by upregulation of IL‐4, IL‐5, and IL‐10 and low levels of the Th‐1‐associated INF‐ (Schonbeck et al., 2002). Th‐2 predominant immune response in AAAs was confirmed in

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a murine model of aortic allograft (Shimizu et al., 2004). Such Th‐2 polarization of T lymphocytes is in keeping with the decreased IFN‐ expression observed in elderly people. The diVerent regulatory roles attributed to the Th‐1 and Th‐2 cytokines in matrix remodeling are certainly of importance in AAAs. Th‐1 cytokines release messengers such as IFN‐ and TNF‐, which will prompt macrophages activation, and then contribute to proteases and proinflammatory cytokines secretion at site of injury. 2. Elastin Peptides Polarize T Lymphocytes Toward a Th‐1 Response Elastin peptides concentration in the circulation might interfere diVerently with time on the underlying mechanisms that drive AAAs development. Previous studies described the presence of the S‐Gal‐elastin receptor on isolated T lymphocytes from human tonsils or atherosclerotic plaques (Peterszegi et al., 1996, 1997a,b) as well as on the surface of nonactivated and activated human circulating lymphocytes (Debret et al., 2005). In this setting, we demonstrated that interaction between EPs and its receptor induced a shift of the systemic T‐cell response toward Th‐1 phenotype (Debret et al., 2005) (Fig. 3). Production of Th‐1 cytokines is positively controlled by IL‐12 and negatively by IL‐4, a Th‐2 cytokine which favors Th‐2 cytokines production. According to Libby’s group, IL‐4 is associated with an increased AAAs formation, increased elastic tissue fragmentation, as well as MMP‐9 and MMP‐12 expression (Shimizu et al., 2004). A fine control of Th‐1/Th‐2 balance is consequently of great importance in AAAs progression. Following EPs stimulation, we found that the levels of the prominent Th‐2 cytokines IL‐5 and IL‐10 secreted by lymphocytes are reduced, whereas the amounts of Th‐1 cytokines IFN‐ and IL‐2 are enhanced. In addition, the ability of EPs to direct the T‐lymphocyte population toward a Th‐1 profile is still eVective when T cells have already been Th‐1 preorientated by an IL‐12 treatment. Strikingly, EPs were also shown to reverse the Th‐2 profile induced by IL‐4 into a Th‐1 profile. Such an eVect is more relevant of atherosclerosis, in which a Th‐1 immune response predominates, rather than AAAs where Th‐2 inflammatory responses seem to predominate. It is conceivable that initiation of an early inflammatory response within the damaged aortic wall will interfere with the biological eVects of elastin peptides by regulating the Th‐1/Th‐2‐mediated immune response, thus either preventing aneurysmal development by reversing Th‐2 profile or amplifying the Th‐1 profile associated to atherosclerosis. More recently, the presence of NK and NKT cells which possess a Th‐0 cytokine profile producing type 2 as well as type 1 (IL‐2 and IFN‐)

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LT

EP Elastin

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IL-5 IL-10

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proMMP-9

proIL-8 → IL-8 Figure 3 Th‐1 polarization mediated by elastin peptides binding on S‐Gal. EPs binding directs T‐lymphocyte population toward a Th‐1 profile from nonorientated lymphocyte (LT), Th‐1‐ preorientated lymphocyte by IL‐12 treatment (Th‐1), and Th‐2‐preorientated lymphocyte following IL‐4 treatment (Th‐2). proMMP‐9 secretion was increased following lymphocytes (LT, Th‐1, Th‐2) stimulation with EPs.

cytokines in AAAs was reported (Chan et al., 2003, 2005a,b). Such an observation suggests that these cells may also play important role in AAAs disease progression particularly through the production of IL‐4 which increases with age (Plackett et al., 2004). Parallely, EPs also markedly increase levels of proMMP‐9 following S‐Gal activation (Debret et al., 2005). Indeed, S‐gal occupancy by EPs led to Erk1/2 and AP‐1 DNA‐binding activation, key actors in coordinating MMPs (Duca et al., 2002) and IFN‐ or IL‐2 transcription (Schafer et al., 2003). Consequently, activation of this transduction pathway might contribute to generate a positive feedback mechanism through exacerbation of MMP‐9 production since IFN‐ and IL‐2 have been reported to stimulate MMP‐9 release by various cell types (Leppert et al., 1995b; Wang et al., 2000; Xiong et al., 2004). Thus, considering that (1) MMP‐9, as a zymogen form, can hydrolyze macromolecular substrates such as type IV collagen (Bannikov et al., 2002) and (2) MMP‐9 displays a wide specificity, an upregulation of active MMP‐9 production by EPs‐activated lymphocyte would in turn facilitate AAAs development as described previously (Galis et al., 1994; Leppert et al., 1995b; Pyo et al., 2000).

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V. The Elastin Connection in Melanoma A. Aging, Fibroblasts Senescence, and Cancer Progression Age is the most important risk factor for the development of cancer in human (Syrigos et al., 2005). Both sexes are aVected with an incidence and a mortality rate superior for men than for women (Desai et al., 2006). Such rising of cancer incidence with age mainly applies to carcinomas, which tend, however, to be less aggressive in oldest individuals (Swetter et al., 2004). Family history of melanoma as well as the presence of numerous atypical nevi depict main risk factors for melanoma but immunosuppression, sun exposure, and age have to be considered as equally important in the progression of this cancer (Miller and Mihm, 2006). Indeed, melanoma incidence and mortality rise continuously in older individuals (>65 years) and elderly patients have a worse prognosis. The pivotal function of the tumor microenvironment in suppressing or promoting the malignant phenotype is now well documented (Kalluri and Zeisberg, 2006) and accumulation of senescent cells in tissues may promote cancer progression in aged individuals (Martens et al., 2003). Among stressful conditions, exposure to hypoxia, UVB, and H2O2 can trigger premature senescence displaying features of replicative senescence such as irreversible growth arrest and SA
‐Gal expression (Finkel and Holbrook, 2000; Toussaint et al., 2000, 2002). Recently, a series of studies revealed that senescent fibroblasts, to a much higher extent as compared to presenescent fibroblasts stimulated the growth of preneoplastic as well as neoplastic epithelial cells. Importantly, it was observed that such growth rate enhancement of tumor cells could be attributed to substances secreted by senescent fibroblasts; as a matter of fact, deposited matrix from senescent fibroblasts markedly increased the growth of preneoplastic epithelial cells (Krtolica and Campisi, 2002). A number of investigations, including microarray analysis, demonstrated that senescence fibroblasts developed a strong inflammatory‐type response with upregulation of MCP‐1, GRO‐, IL‐1
, and IL‐15 (Shelton et al., 1999). Parallely, a set of matrix‐degrading proteases are overexpressed. Among them are the partners of the proteolytic cascade leading to collagenolysis namely uPA, tPA, stromelysin(s) (MMP‐3), and collagenase (MMP‐1). It has to be considered that enzymes displaying elastolytic activity, such as cathepsin K, MMP‐2, and a 94‐kDa metalloendopeptidase, are also upregulated in senescent fibroblasts (Homsy et al., 1988; Zeng and Millis, 1994). Such potent proteolytic phenotype is not counterbalanced by either increased expression of MMP inhibitor or upregulation of matrix synthesis but instead TIMP‐1, elastin, as well as type I and III collagens expression are downregulated with fibroblasts senescence (Bizot‐Foulon et al., 1995).

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Such proteolytic pattern can create a favorable cancerization field through disruption of matrix barriers promoting cancer cell invasiveness. Additionally, MMPs as MMP‐3 and MMP‐1 can influence cancer development. MMP‐3 was found to cleave E‐cadherin and catenins from cell–cell contacts, thus inducing an epithelium–mesenchymal transition; also, exposure of mammary epithelial cells to MMP‐3 induces the expression of an alternatively spliced form of Rac‐1, which further results in ROS overproduction, DNA damage, and genomic instability (Radisky et al., 2005). Alternatively, cleavage of protease‐activated receptor‐1, a G‐protein–coupled receptor, by MMP‐1 at the appropriate site leads to receptor activation, Ca2þ signaling, and breast carcinoma cell migration (Boire et al., 2005). Additionally, investigation highlighted unexpected localizations of MMP in association with cell mitochondria and nuclei, where they may be involved in apoptosis as MMP‐3 (Si‐Tayeb et al., 2006), or on the contrary can confer resistance to lamin A/C degradation and apoptosis as MMP‐1 (Limb et al., 2005). Altogether, these data indicated that fibroblasts senescence, whose pattern of gene expression (proteases, cytokines) is somewhat similar to that of cancer associated fibroblasts, might play a significant role in the growth and spread of skin cancer including melanoma.

B. EPs as Modifiers of the Dermal Stroma As observed in arteries, human skin intrinsic aging is characterized by important alteration of elastic fibers with nearly total disappearance of oxytalan fibers as well as fragmentation and, in some areas, scarcity of elaunin and dermal elastic fibers. The situation is somewhat diVerent in extrinsic skin aging, that is photoaging. Although, UV‐mediated activation of fibroblasts and inflammatory cells can contribute to the liberation of MMP‐1, MMP‐3, and MMP‐9 (Fisher and Voorhees, 1998), a massive accumulation of ‘‘elastotic’’ material is observed in the upper and mid‐dermis (Uitto and Bernstein, 1998). Collagen fibers meshwork, at the vicinity of elastic fibers, appears greatly degenerated (Uitto and Bernstein, 1998). Increased elastases production by senescent fibroblasts (Cat K, fibroblast metalloelastase, MMP‐2) is probably a key factor of dermal elastolysis and production of EPs. Importantly, exposure of human skin fibroblasts to UV, one main risk factor in melanoma initiation and fibroblast senescence in cancer, was shown to induce the upregulation of MMP‐1, MMP‐3, and also MMP‐9 through activation of cytokine and growth factors receptors leading to increased expression of the transcription factor AP‐1 (Fisher and Voorhees, 1998). We evidenced that EPs could upregulate proMMP‐1 and proMMP‐3 expression in skin fibroblasts maintained in culture, thus potentiating collagenolysis (Brassart et al., 2001). MMP production is regulated both at the mRNA

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and protein levels via a signaling cascade in which MEK/Erk module activates AP‐1 transcription factors (Duca et al., 2005). However, under our experimental conditions, MMP‐1 and MMP‐3 were secreted as zymogens. We initially hypothesized that generation of active enzyme would necessitate the participation of the plamin(ogen) system from inflammatory or cancer cells since early‐passage skin fibroblasts only produced low levels of either tPA or uPA. However, these serine proteases are highly expressed in senescent fibroblasts, suggesting that the uPA (tPA)‐plamin(ogen)‐(pro)MMP‐3‐ (pro)MMP‐1 proteolytic cascade could be triggered by aged fibroblasts in presence of EPs. As a matter of fact, MMP‐1 is considered as one main target enzyme in cancer and its expression is most often negatively correlated with survival (Overall and Kleifeld, 2006). It has been shown that cellular senescence can be provoked in normal cells by continuous exposure to mitogenic signals and overexpression of signal transducing proteins such as Ras, Raf, or AKT (Minamino et al., 2004; Serrano et al., 1997; Sewing et al., 1997). As mentioned previously, EPs increases fibroblasts proliferation (Kamoun et al., 1995) and S‐Gal occupancy trigger Ras or Raf activation depending on cell type (Duca et al., 2005; Mochizuki et al., 2002). It raises the possibility that the prolonged action of these matrikines, as occurring during aging, might somewhat accelerate a senescent phenotype in fibroblasts.

C. EPs and Melanoma Progression Melanoma can progress in several stages from dysplastic nevus to radial growth phase (RGP) nontumorigenic primary melanoma, which is characterized by increased expression of MMP‐9. RGP is then followed by a vertical growth phase (VGP) where melanoma cells infiltrate and invade the dermis as large cluster of cells and further develop metastases. Transition from RGP to VGP is associated with a proteolytic switch, where increased expression of MMP‐1, MT1‐MMP, and MMP‐2 is mainly evidenced (Hornebeck et al., 2005; Labrousse et al., 2004). Implicitly, it might be hypothesized that one or several factors from underlying dermis could be responsible for RGP–VGP transition. Alteration of elastic fibers and liberation of elastin peptides, as part of an age‐associated cancerization field, might belong to these promoting factors. Our earliest investigation demonstrated that EPs exhibited strong chemotactic activity for melanoma cells expressing S‐Gal (Timar et al., 1991). More recently, we observed that S‐Gal expression was intensively expressed at the melanoma invasion site and colocalized with intense expression of MMP‐2 and MT1‐MMP (Ntayi et al., 2004). In vitro experiments demonstrated that S‐Gal occupancy by EPs upregulated MT1‐MMP and MMP‐2 expression and activation, which

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together with MMP‐1 participate in type I collagen invasion in a modified Boyden chamber assay. It has to be underlined that MMP‐2 upregulation was reported to be correlated with low survival rate, independently of Clark and Breslow microstage (Hofmann et al., 2000). Such EPs‐mediated MMP‐2 overproduction is in keeping with the intense fragmentation of elastic fibers at the front of melanoma invasion; to that respect, disappearance of elastic fibers in the melanoma depth was linked to adverse prognosis (Hornebeck et al., 2005) (Fig. 4). As observed in normal cell types, EPs binding to S‐Gal at the surface of highly invasive melanoma cells, triggered MAPK pathway known to be recruited by matrikines for MMP and cytokine(s) such as IL‐1
, IL‐8, and Gro‐ expression (Debret et al., 2006). Both Erk1/2 and p38 were activated and gel shift analysis indicated that AP‐1 DNA‐binding was increased 1 hour following EPs stimulation. Unexpectedly, a series of genetic and pharmacological approaches indicated that MAPK pathway was not involved in EPs‐mediated IL‐1
upregulation. Instead, supplementation of culture medium with SN50, a specific NF‐B inhibitor, or/and IB overexpression totally abolished overexpression of IL‐1
mRNA by EPs. In parallel, we demonstrated that EPs stimulation led to NF‐B translocation and DNA

RGP

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Dermis Normal elastic fibers in the reticular dermis at distance of melanoma tumor

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MAPK P38 AP-1

NF-k B IL-1b IL-8 GROa

Figure 4 EPs binding to S‐Gal on melanoma cell surface favors tumor progression. Fragmented appearance of elastic fibers in melanoma lesions is associated with RGP–VGP transition. Locally generated EPs will increase proteolytic enzymes secretion and activation (Ntayi et al., 2004), and will induce MAPK‐P38 transduction pathways, AP‐1‐NF‐B transcription factors, and cytokines (IL‐1
, IL‐8, Gro) expression (Debret et al., 2006), thus generating a dual feedback loop.

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binding on IL‐1
promoter (Debret et al., 2006) (Fig. 4). Therefore, signaling pathways such as NIK, PI3K/AKT, PKC, or MAP3Kinase, but not MEK/ Erk1/2, might be requested for NF‐B‐mediated eVect on IL‐1
gene expression on EPs stimulation in melanoma cells (Amiri and Richmond, 2005). IL‐ 1
released by EPs‐activated melanoma can potentiate MMP‐3 and MMP‐1 expression by fibroblasts at vicinity and favor melanoma cell invasion through increased collagenolysis. Most often, NF‐B is constitutively activated in many carcinoma and melanoma. Our data supported the contention that EPs could participate in sustained NF‐B activation at the invasive front of melanoma, thus contributing to apoptosis resistance, proliferation, and invasion of melanoma cells. Interestingly, NF‐B also mediates transcription of IL‐8, which was described to markedly increase the tumorigenicity of melanoma cells through MMP‐2 upregulation (Luca et al., 1997).

D. EPs‐Mediated Angiogenic Phenotype As documented in Section IV, EPs, at concentration as low as 10 ng/ml, could accelerate the rate of pseudotubes formation from micro‐ or macrovascular endothelial cells in vitro and in vivo (Robinet et al., 2005). Angiogenesis is one hallmark of melanoma progression since horizontal growth phase, characterized by slow progression, is avascular, whereas VGP, associated with metastasis and death, appears highly vascularized (Sosman and Puzanov, 2006; Srivastava et al., 2003). However, important heterogeneity in the extent of angiogenesis among individuals has been reported and the prognostic significance of angiogenesis in melanoma progression is still matter of debate. For some investigators, tumor vascularity was found to be the most important factor determining overall survival (Kashani‐Sabet et al., 2002), whereas others claimed that tumor thickness, but not tumor vascularization, was the only independent variable associated with disease‐ free survival (Ilmonen et al., 1999). As we reported (Robinet et al., 2005), EPs‐mediated influence on pseudotubes formation was attributed to MT1‐MMP upregulation, which besides could activate proMMP‐2, thus contributing to elastolysis and an amplification loop. Like in melanoma cells, we also noticed (Bellon, G., Fahem, A., and Hornebeck, W., unpublished observations) that EPs, through S‐Gal occupancy, could induce IL‐8 expression by HUVEC and a dermal vascular endothelial cell line (HMEC‐1). IL‐8 is overexpressed in the majority of cutaneous melanoma and its high serum level was correlated with poor overall survival (Nurnberg et al., 1999). IL‐8 treatment of melanoma cell line with low metastatic potential significantly enhances their proliferation and metastatic potential (Varney et al., 2003). Accordingly, IL‐8 and CXCR2 (IL‐8R2) expressions are lower in melanoma with Clark level I and II as

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compared with specimens with Clark level IV and V (Varney et al., 2006). In this setting, binding of IL‐8 secreted from endothelial cells onto CXCR1 (IL‐8R1) was found to induce chemotaxis and transendothelial migration of melanoma cells (Ramjeesingh et al., 2003). Of interest for melanoma progression, MMP‐2 activity is also increased in presence of EPs and IL‐8 processing induced by MMP‐9 potentiates more than 10‐fold of its activity (Van den Steen et al., 2000). All eVects of EPs on endothelial cells appeared NO‐mediated, which is rapidly released from cells following supplementation of culture medium with peptides concentration as low as 10 ng/ml. NO eVects on melanoma progression are pleiotropic, being either beneficial or detrimental (Ekmekcioglu et al., 2005). For instance, NO release by endothelium at the vicinity of B16 melanoma cells in the mouse pulmonary circulation induces tumor cell apoptosis (Qiu et al., 2003); also, NO were reported to increase the eYciency of cytostatic therapy against melanoma metastasis (Konovalova et al., 2003; Matthews et al., 2001). In turn, exposure to NO could induce MMP‐1 expression in melanoma cells through triggering Erk1/2 and p38 signaling pathways, thus contributing positively to tumor progression (Ishii et al., 2003). However, somewhat in contradistinction with the increased incidence of melanoma with aging, eNOS expression and activity and NO production are impaired in senescent human endothelial cells (Haendeler, 2006; Matsushita et al., 2001). Accordingly delayed angiogenesis, associated with reduced levels of eNOS, p‐eNOS, and iNOS, was observed in aged rats using a polyvinyl alcohol sponge implant model (Bach et al., 2005). Although aging eVect on angiogenesis is quite disturbing with regard to cancer progression, it needs to be delineated that melanoma cells could elaborate two other biological mechanisms to supply nutriments in order to restore tumor progression. First, melanoma cells can disseminate by lymphatic vessels to lymph nodes, and lymphangiogenesis is now recognized as one main prognostic factor for cutaneous melanomas (Dadras and Detmar, 2004; Streit and Detmar, 2003). Nevertheless, to our knowledge, the influence of aging on lymphatic network has still received little attention. Besides, recent investigations revealed that aggressive melanoma cells reverted to an embryonic‐like phenotype and formed vascular‐like channel networks, a phenomenon designated as vasculogenic mimicry (Maniotis et al., 1999). Formation of such channels could be complementary to angiogenesis in tumor dissemination. Among factors that can promote vascular mimicry are MMPs‐ driven matricryptins (or matrikines) formation together with their corresponding melanoma cells receptor. Cleavage of laminin‐5 2 chain by MT1‐MMP and/or MMP‐2 results in fragments with migratory and vasculogenic properties for melanoma cells (Hendrix et al., 2003). Therefore, EPs might indirectly participate in formation of such melanoma channels as being potent inducers of MT1‐MMP and MMP‐2 for this cancer cell type. In keeping with

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recent data from J. Timar’s group, who showed that parallel expression of IIb
3 and V
3 integrins promoted the angiogenic phenotype of melanoma cells (Dome et al., 2005), matrix fragments such as the hydrophilic C‐terminal part of tropoelastin that binds to
3 integrin (Rodgers and Weiss, 2004) could, contrary to VGVAPG eVect, somewhat interfere with vascular mimicry. Indeed, there is no direct relationship between pro‐/antiangiogenic and pro‐/antivascular mimicry. For instance, endostatin, a potent antiangiogenic matrikine, proved to be ineYcient in interfering notably on vascular mimicry, probably due to the lower level of expression of receptor as 5
1 on melanoma cells as compared to endothelial cells (Hendrix et al., 2003).

E. EPs and Inflamm‐Aging in Melanoma In the mid‐nineteenth century, Virchow already suggested that cancer developed mainly at sites of chronic inflammation. Nowadays, the causal link between inflammation, innate immunity, and cancer progression becomes more widely accepted (Balkwill and Mantovani, 2001; Coussens and Werb, 2002). Inflammatory cells and cytokines expressed at site of tissue injury can play a dual role, either by favoring tumor growth and invasion or by mounting an immune response. To that respect, inflammatory cells at tumor site can be considered either as important tumor promoter or, instead as tumor suppressor as part of a host response to suppress tumor progression. This diVerential regulation might be dependent on a balance controlled by a threshold beyond which inflammation could become aggressive, and which could be a mirror of genetic background, stress history, and most importantly aging. As we mentioned previously, EPs produced by the combined action of elastases from melanoma cells and fibroblasts have potent chemotactic activity for monocytes. Tumor‐associated macrophages are main components of the infiltrate of most tumors and macrophage infiltration was found to be correlated with Breslow index in melanoma (Hussein, 2006). Although, macrophages have potent stimulatory eVect on tumor cell proliferation and angiogenesis, they also display killing activity for cancer cells when appropriately activated (Fulop et al., 1997; Varga et al., 1988, 1997). Accordingly, low‐grade melanoma is characterized by the presence of consequent lymphocyte brisk infiltrate at tumor vicinity, but its intensity decreases substantially in high‐grade tumor (Clemente et al., 1996). Interestingly, EPs were found to be chemoattractant for T lymphocytes (Antonicelli, unpublished observation) and studies by Robert’s group previously evidenced that S‐Gal occupancy by EPs could stimulate their proliferation (Peterszegi and Robert, 1998). To that respect, EPs may be considered to exhibit a beneficial influence against melanoma progression. Indeed, a topical escape from immunologic

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surveillance is observed in human cancer progression. Among the various mechanisms involved, inhibition of tumor‐infiltrating lymphocyte (TIL) cytolytic potential can be held responsible for increased tumor progression. Both CD4þ and CD8þ TIL are requested to develop an eYcient tumor response. This cooperation involves lymphokines expressed by CD4þ TIL in the activation of the cytolytic activity of CD8þ TIL. As a matter of fact, decreased proportion of the CD4þ TIL is correlated with SCC tumor progression (Sheu et al., 1999). This prognostic significance of low CD4þ/CD8þ ratio in TIL has also been demonstrated for metastatic melanoma (Hernberg et al., 1997) and lung carcinoma (Yoshino et al., 1993). In a previous study, we demonstrated the presence of lymphocyte at the vicinity of the melanoma tumor (Ntayi et al., 2004) that might have been recruited by EPs. To obtain an eYcient host immune response against most cancer, including melanoma, the Th‐1/Th‐2 balance is equally of importance. As already depicted, EPs favor a Th‐1 profile in vitro. Thus, release of IFN‐ and IL‐2 from EPs‐activated CD4þ lymphocytes could reactivate CD8þ lymphocytes and therefore promote a protective antitumor immune response. This EPs‐ mediated putative beneficial influence may be counterbalanced by their inducing eVect on protease expression by lymphocytes (Fig. 5). Strikingly, EPs were described to stimulate the expression of leukocyte elastase and cathepsin G by lymphocytes (Peterszegi and Robert, 1998), and we

Keratinocytes

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Cell growth invasion Elastin EP EP

CTL +

CD8 TIL

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Th-1 orientation IL-2 IFN-g

EP

Th-1 LT CD4+

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Figure 5 Hypothetical scheme of EPs eVect on Th‐1/Th‐2 orientation in melanoma progression. Generation of elastin peptides at front of tumor invasion by proteolytic enzymes favors lymphocyte recruitment and orientation (Th‐1). If the lymphocyte infiltrate (TIL) is consistent enough, Th‐1 cytokines (IFN‐, IL‐2) secretion will activate cytolytic T lymphocyte (CTL) and favor tumor regression. Change in cell targeted by EPs, melanoma instead of lymphocyte, on aging or immunodepression increases MMPs secretion and melanoma progression.

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demonstrated that EPs‐induced Th‐1 cytokine upregulation is associated with activation of Erk1/2, increase of AP‐1 DNA binding, and enhance proMMP‐9 secretion (Debret et al., 2005) that, in turn, may potentiate immunodepression through IL‐2RA cleavage (Sheu et al., 2001). Thus, EP might initiate a local elastolytic amplification loop with combined action of leukocyte elastase, cathepsin G, and MMP‐9, increasing the concentration of elastin fragments at lymphocyte vicinity, therefore favoring Th‐1 orientation. However, exacerbation of this feedback loop could reverse the beneficial eVect into a detrimental eVect. Indeed, it has been shown that excess EPs (mg/ml) induce necrosis and apoptosis of lymphocytes expressing S‐Gal (Peterszegi and Robert, 1998) (Fig. 5). The decrease of immune response during aging, leading to higher susceptibility to infections and cancer, is well documented. Fulop and coworkers pinpointed that alteration in the composition of lipid rafts (ganglioside M1, cholesterol) with aging could modify the functions of neutrophils and CD4þ and CD8þ T cells (Fulop et al., 2006; Larbi et al., 2006). Such changes deeply alter signaling pathway triggered by several receptors. It might be of interest to evaluate whether S‐Gal uncoupling, as observed in other cell types, can be similarly evidenced in aged lymphocytes modifying EPs chemotactic eVects and/or polarization influence. Of note, EPs‐mediated eVect on elastase production by neutrophils and lymphocytes were found unaVected by age, raising the intriguing possibility that only deleterious functions of these matrikines are kept invariant in oldest individuals (Robert, 1999).

VI. Concluding Remarks Intuitively, one may hypothesize that degradation of one of the longest lived protein in human can give rise to potent signals for cell survival and tissue repair (Senior et al., 1980). Indeed, elastin peptides are among the most potent chemotactic peptides for leukocytes and display proangiogenic activity close to the one exerted by VEGF (Robinet et al., 2005). At an average 10‐fold higher concentration, these elastin fragments can promote lymphocyte proliferation (Peterszegi and Robert, 1998; Peterszegi et al., 1996). The array of elastases expressed by macrophages within aVected elastic tissue will further amplify elastolysis and elastin fragments, we designated as elastokines, will direct the migration and induce a remodeling program by fibroblasts or smooth muscle cells. All events are mediated, mostly by interaction between S‐Gal and peptides containing a GXXPG consensus sequence; it is worthwhile to specify that such sequence is also present as multiple motifs in microfibrils and elastin‐associated proteins (Booms et al., 2006; Kielty et al., 2002), thus suggesting that proteolytic attack of these molecules might represent the primary repair signal for a given elastic tissue. Aging is characterized by increased expression of tissue elastases, leading to progressive disruption of elastic fibers and release of elastokines, which

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Matrix proteolysis

MMP1,3

Elastin SMC

Fibroblast Elastases: metallo, serine, cysteine classes

PMN (Chemotaxis)

AGING

T-lymphocyte (Chemotaxis, Th-1 response)

«Elastokines » Elastase Cat. G

MMP9 elastase Cat. G

Aging Osteoblast-like cell (Elastin fiber calcification) Monocyte (Inhibition of cholesterol synthesis chemotaxis?)

MMP2, 14 MT1-MMP, NO

Endothelial cell (angiogenesis vasodilation)

Arterial wall diseases

Cancer cell (melanoma) (migration) Cancer

Figure 6 Biological properties of elastin peptides.

can further (1) stimulate MMP expression; (2) induce the diVerentiation of smooth muscle cells to osteoblasts and therefore participate to elastic fiber calcification and lipid deposition; and (3) polarize lymphocytes to a Th‐1 cytokine profile, creating favorable conditions for chronic inflammation and further development of arterial wall diseases (Fig. 6). Similarly, these elastin peptides increase MMP expression by fibroblasts; furthermore, continuous exposure of fibroblasts to elastokines leading to Ras and/or RAF as well as AKT activation could accelerate a fibroblast senescent phenotype, a contention that needs to be confirmed by transcriptomic and proteomic analyses. With age, as in the arterial wall, a chronic inflammatory state could be elaborated within dermis favoring here a cancerization field. Besides these above listed functions, elastokines display potent chemotactic activity for melanoma cells and trigger MT1‐MMP and MMP‐2 expression by these cells, properties that may catalyze the RGP–VGP transition in the progression of this cancer (Fig. 5). The property of genetic traits to display beneficial and deleterious eVect for an organism has been named antagonistic pleiotropy. To that respect, a posttranslational mechanism as elastolysis might share similar property, being quite essential in directing elastic tissue repair in young individuals but participating to the formation of a chronic inflammatory state during aging

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linked to the development of degenerative diseases and cancer. Of note, increased level of elastokines into the blood circulation is considered to have prognostic value in AAAs; to our knowledge, variation in the amount of elastin peptides within the circulation, at aims to be related to disease progression, has not been evaluated in skin, breast, or lung cancers. Elastase‐type proteases, mainly from the MMP family, are conductors of this chronic inflammation state through their ability to generate matrikines and/or to modulate the activity of several cytokines, chemokines, and growth factors. Recently, MMPs have been classified as either target or antitarget in cancer progression (Overall and Kleifeld, 2006). Among targets in carcinomas and melanomas are collagenase, that is MMP‐1, and gelatinase A, that is MMP‐2. Importantly, gelatinase B, that is MMP‐9, is mainly involved in the earlier RGP of melanoma progression. Although several elastases from diVerent families can collaborate in elastic fibers destruction during aortic abdominal aneurysms, combined invalidation of gelatinases genes render mice totally resistant to aneurysm formation. For several years, we attempted to design specific gelatinase inhibitors, taking galardin as a template for structural modification of primed or unprimed subsites. Our recent data indicated that introducing an isobutylenic group and long alkyl chain at the S’1 subsite of galardin confered specificity for MMP‐2 inhibition with Ki’s orders of magnitude threetimes lower than those determined for other MMPs (Marcq et al., 2003). Modification of the hydroxamate group with a hydrazide, which allowed synthesis of galardin derivatives occupying unprimed subsites, led instead to compounds which inhibited preferentially MMP‐9 (Auge et al., 2004). Elastases as pancreatic or leukocyte elastase, MMP‐2, and MMP‐9 bind to elastin using domains remote from their active site (Emonard and Hornebeck, 1997). Fibronectin domains, mainly FnII (Kashani‐Sabet et al., 2002) for gelatinase A, were shown to direct gelatinase interaction to elastin and elastolysis (Berton et al., 2001). The exosite domain in leukocyte elastase responsible for elastin binding has not been characterized; strikingly, N‐terminal domain of porcine pancreatic elastase and human leukocyte elastase were described to present structural homologies with the V14‐specific sequence of S‐Gal (Hinek, 1996; Hinek and Rabinovitch, 1994; Hinek et al., 1988). The design of peptide, peptide mimetic, or substances as unsaturated fatty acids that interact specifically with the elastin‐binding domain of one elastase could be useful to confer protection against elastolysis. Such substances can be further derivatized to contain a specific elastase inhibitor; these types of bifunctional agents will provide protection against elastolysis while leaving other functions of the protease intact. In this setting, S‐Gal might be an alternate target to impede the deleterious functions of elastokines in aneurysms and skin cancer progression. QDEA‐containing V14 S‐Gal peptide proved to annihilate, ex vivo and in vivo, elastokines functions on angiogenesis

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(Robinet et al., 2005). The eVect of such peptide in experimental animal models of several degenerative diseases aVecting elastic tissues need to be evaluated. Also, taking into account the lectin‐like property of S‐Gal, galactosugars as melibiose or derivatives might be similarly tested. The elastin peptides‐induced inflamm‐aging and related diseases are only one aspect of the influence of matrix‐directed proteolysis on chronic inflammation. Disease progression with aging will undoubtedly depends on the critical balance between ‘‘target’’ and ‘‘antitarget’’ cytokines, chemokines, proteases, and matrikines.

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A Phylogenetic Approach to Mapping Cell Fate Stephen J. Salipante* and Marshall S. Horwitz{ *Department of Genome Sciences University of Washington School of Medicine Seattle, Washington 98195 { Department of Medicine, Division of Medical Genetics University of Washington School of Medicine Seattle, Washington 98195

I. Development II. Fate Maps A. Fate Maps from Direct Observation B. Mapping Development with Interspecies Chimeras C. Tracing Cell Lineage Through Cell Marking D. Cell Marking with Genetic Markers III. Limited Information from Cell Marking IV. Stochastic Nature of Development in Higher Organisms V. A New Approach: Phylogenetic Fate Mapping A. Introduction to Phylogenetics B. Somatic Mutations C. Tests in Cell Culture D. Mapping Lineages in Whole Organisms VI. Challenges Dealing with Randomness in Development VII. Extended Applications of Phylogenetic Fate Mapping Acknowledgments References

Recent, surprising, and controversial discoveries have challenged conventional concepts regarding the origins and plasticity of stem cells, and their contributions to tissue regeneration, and highlight just how little is known about mammalian development in comparison to simpler model organisms. In the case of the transparent worm, Caenorhabditis elegans, Sulston and colleagues used a microscope to record the birth and death of every cell during its life, and the compilation of this ‘‘fate map’’ represents a milestone achievement of developmental biology. Determining a fate map for mammals or other higher organisms is more complicated because they are opaque, take a long time to mature, and have a tremendous number of cells. Consequently, fate mapping experiments have relied on tagging a progenitor cell with a dye or genetic marker in order to later identify its descendants. Current Topics in Developmental Biology, Vol. 79 Copyright 2007, Elsevier Inc. All rights reserved.

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This approach, however, extracts little information because it demonstrates that a population of cells, all having inherited the same label, shares a common ancestor, but it does not reveal how cells in that population are related to one another. To avoid that problem, as well as technical limitations of current methods for mapping cell fate, we, and others, have developed a new strategy for retrospectively deriving cell fate maps by using phylogenetics to infer the order in which somatic mutations have arisen in the genomes of individual cells during development in multicellular organisms. DNA replication inevitably introduces mutations, particularly at repetitive sequences, every time a cell divides. It is thus possible to deduce the history of cell divisions by cataloging somatic mutations and phylogenetically reconstructing cell lineage. This approach has the potential to produce a complete mammalian cell fate map that, in principle, could describe the developmental lineage of any cell and help resolve outstanding questions of stem cell biology, tissue repair and maintenance, and aging. ß 2007, Elsevier Inc.

I. Development Multicellular sexual organisms are composed of cells descended from a single‐cell zygote, and development is the result of cumulative cycles of mitotic cell division, migration, apoptosis, and diVerentiation. Classical embryology incorporates two exclusive views of embryogenesis distinguished by when a cell’s developmental potential for giving rise to diVerent tissues becomes apparent (Stern and Fraser, 2001): In so‐called ‘‘mosaic’’ species, embryonic cell fate is restricted during the first few cell divisions such that if an early embryonic cell (a ‘‘blastomere’’) is removed, it will begin diVerentiating in vitro, and the remaining cells of the embryo will be unable to compensate for its loss. For example, in tunicates, blastomeres separated from the two cell embryo develop into distinct half‐embryos. Contrastingly, in what are called ‘‘regulative’’ species, blastomeres remain multipotent until a later stage, when cell–cell interactions define their destiny, such that separated blastomeres yield identical twins, and the rest of the embryo remains unaVected by their loss. For example, the cells from the inner cell mass of mammalian embryos remain pluripotent, and, in fact, can serve as a source for embryonic stem (ES) cells. Nevertheless, as highlighted by recent, surprising, and controversial discoveries regarding stem cell plasticity and ‘‘transdiVerentiation’’ (Anderson et al., 2001; Blau et al., 2002; Eckfeldt et al., 2005; Orkin and Zon, 2002; Theise and Wilmut, 2003; Wagers and Weissman, 2004), classical paradigms may no longer be relevant (Lawrence and Levine, 2006; Theise and Krause, 2001).

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II. Fate Maps In order, therefore, to understand how cells determine their developmental destiny, it is useful to identify particular cells and all of their descendants and to construct ‘‘fate maps’’ that reveal what each part of the embryo becomes at later stages of development and, conversely, that allow for tracing of the embryonic origins of specific cells at later stages. A. Fate Maps from Direct Observation The simplest approach to constructing a cell fate map, first employed with tunicates more than a century ago, is to directly observe the cell divisions of an embryo as it grows (Clarke and Tickle, 1999; Stern and Fraser, 2001). Such studies reached an apex when Sulston and colleagues used a microscope to painstakingly record (Fig. 1) the lineage of each of the 671 cells produced during Caenorhabditis elegans 12‐hour embryogenesis (Sulston et al., 1983), an accomplishment rewarded, in part, with the 2002 Nobel Prize (Sulston, 2003). B. Mapping Development with Interspecies Chimeras Determining embryonic lineages in higher organisms through direct observation is impossible because the embryos are, of course, not transparent, because of their lengthy gestation, and because of the diYculty of accounting for the trillion or more cells that may be present by the time of birth. Consequently, diVerent techniques are needed. One method is to generate chimeric animals in which one part of an embryo is transplanted to the corresponding anatomical location of a diVerent embryo, where species diVerences allow for the donor tissue to be distinguished from the host. Le Douarin (2005) has pioneered this approach using quail–chick chimeras (Fig. 2). There are limits, however, to how finely embryos may be dissected and reconstructed, and graft rejection prohibits long‐term follow‐up in adults. C. Tracing Cell Lineage Through Cell Marking In essence, the construction of a chimeric embryo allows for a subset of cells to be distinguished chiefly on the basis of pigmentation. Not surprisingly, then, other, less invasive and higher resolution, experimental attempts at mapping cell fate have relied on marking just a single cell at some point during embryogenesis and then later identifying descendants of that cell

Zygote

1 Cell fate map of Caenorhabditis elegans from zygote to completion of embryogenesis (based on the observations made by Sulston et al., 1983). Figure courtesy of Nicholas Rhind and Sean Ryder, University of Massachusetts Medical School, used with permission. Figure-

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Figure 2 Fate mapping via quail to chick transplantation. Figure from Le Douarin (2005), used with permission (Copyright figure courtesy of Nicole Le Douarin, Institut d’Embryologie in Nogent‐sur‐Marne, France, and the International Journal of Developmental Biology).

also containing the marker. The simplest way to tag a cell is to inject a dye; although widely utilized, and in addition to technical issues related to dye diVusion, toxicity, and fading, dyes obviously become exponentially diluted every time a cell divides and are thus useful for tracking only over a limited number of cell generations. Another problem is that the manipulation of the embryo that is required for dye injection and later observation may make it diYcult to access the structures of interest without perturbing the embryo’s normal development (Clarke and Tickle, 1999; Stern and Fraser, 2001). D. Cell Marking with Genetic Markers An alternative approach for marking a cell is thus to introduce a permanent genetic change heritable by daughter cells such that it cannot be diluted through successive divisions (Clarke and Tickle, 1999; Nesbitt and Gartler, 1971; Stern and Fraser, 2001). 1. Gynandromorphs An early implementation of the strategy of mapping development from embryonic mosaicism took advantage of ‘‘gynandromorphs.’’ Gynandromorphs occur when an organism displays a mixture of female and male

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tissues within a single individual. The diVerent sexes of a species are often diVerently colored or otherwise morphologically distinct. Because gender is frequently controlled by the number of sex chromosomes, the loss of a sex chromosome through a mitotic nondisjunction event can alter the appearance of tissues in which this has happened. Should sex chromosome aneuploidy arise developmentally, then all of the descendants of the cell suVering the mutation will be marked by a change originating from their switch in gender. Examples of gynandromorphs among butterflies are especially striking because they exhibit a segmental asymmetry revealing the point in their development at which mosaicism was generated (Fig. 3); in fact, the novelist— and lepidopterist—Vladimir Nabokov recalled in autobiography (Nabokov, 1966) how, as a child, he once encountered such a specimen. Gynandromorphs occur in other species, including plants (Hake and Freeling, 1986; Stewart and Dermen, 1979) and humans (Malan et al., 2006). Sturtevant was the first to exploit gynandromorphs (Sturtevant, 1929) in order to systematically chart development. He made use of a Drosophila strain in which females heterozygous for sex‐linked genes responsible for recessive yellow pigmentation occasionally randomly lost an X chromosome at early cell divisions, thus imparting a color change that allowed for tracking of the hemizygous cell and its progeny (Fig. 4). Later eVorts made use of unstable ring X chromosomes (Lewis, 1963) or employed X‐rays to induce sex chromosome aneuploidy in flies (Stern et al., 1963), enabling the construction of reasonably complex cell fate maps (Gehring et al., 1976; Janning, 1978). A somewhat similar system, later developed in the mouse, utilized chemical mutagenesis to inactivate a recessive gene encoding a lectin‐binding protein in the small intestine in heterozygous animals, where clonal progeny could be followed by their lack of staining with lectin conjugated to a histochemical marker (Winton et al., 1988). 2. Genetically Engineered Markers The ability to manipulate the genome via transgenic and gene‐targeted recombination has greatly facilitated cell lineage studies. One approach has taken advantage of construction of a transgenic mouse containing a ‐galactosidase‐ encoding LacZ marker gene functionally inactivated through a partial tandem duplication (Eloy‐Trinquet et al., 2000); occasionally, spontaneous intra‐allelic mitotic recombination regenerates the marker and thereby allows for tagging of clonal descendants. Analysis of thousands of embryos marked this way has helped to define the patterning of myotomes (Fig. 5). Another transgenic approach has utilized a mouse ubiquitously expressing a gene for green fluorescent protein (GFP) inactivated with a stop codon, whose reversion following chemically induced mutagenesis similarly tags a clonally descended population of cells (Ro and Rannala, 2004).

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Figure 3 Gynandromorphs and other genetically mosaic organisms. (A) Segmented colony of Saccharomyces cerevisiae showing Ade2 telomeric position eVect (courtesy of Daniel Gottschling, Fred Hutchinson Cancer Research Center, used with permission). (B) Leaf from Ctenanthe oppenheimiana ‘‘Tricolor’’ demonstrating chloroplast chimerism (specimen courtesy of Douglas Ewing, University of Washington greenhouse). (C) Antherina suraka gynandromorph (courtesy of Heinz Rothacer, Aigle, Switzerland, used with permission). (D) Papilio dardanus (African swallowtail) gynandromorph (courtesy of Alexandra Freeman, Doctoral Thesis, Oxford, used with permission). (E) Papilio aegeus (orchard swallowtail) gynandromorph (courtesy of Ismor Fischer, University of Wisconsin, Madison, used with permission). (F) Zerene eurydice (California dogface) gynandromorph (used with permission, Copyright courtesy of T.W. Davies, California Academy of Sciences). (G) Homarus americanus (Maine lobster) gynandromorph (Copyright courtesy of Bob Semple, Department of Fisheries and Oceans, Canada). (H) Homarus americanus (Maine lobster) gynandromorph (used with permission: Copyright courtesy of Abigail Curtis, the Bangor Daily News). (I) Blattela germanica (German cockroach) gynandromorph (courtesy of Joseph Kunkel, University of Massachusetts, Amherst). (J) Pheucticus ludovicianus (rose‐breasted grosbeak) gynandromorph (courtesy of Robert Mulvihill, Powdermill Avian Research Center, Rector, Pennsylvania). (K) Callinectes sapidus (Chesapeake Bay blue crab) gynandromorph (used with permission, Copyright courtesy of Virginia Institute of Marine Science).

A diVerent innovation has involved genetically marking cells through retrovirus‐mediated transfer of a transgene encoding ‐galactosidase or another colored marker (Lemischka, 1993; Price et al., 1987; Soriano and Jaenisch, 1986). DiVusion of the marker from one cell to the next can still

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pose a problem, even with genetically marked cells, where cell fusion, for example, can lead to misinterpretation of results (Vassilopoulos et al., 2003; Wang et al., 2003). Moreover, ‘‘autofluorescence’’ and endogenous ‐galactosidase activity can artifactually complicate the interpretation of GFP‐ and LacZ‐based cell‐marking studies (Newsome et al., 2004), respectively. These methods all select cells to be followed on the basis of their position in the embryo at the time of labeling, but sometimes cells giving rising to a particular tissue or structure may not all be in the same place at a particular time during development; instead, spatially scattered cells converging toward related fates may share expression of common developmental markers (Stern and Fraser, 2001). The earliest experiments addressing this problem employed a transgenic marker under the control of a tissue‐specific promoter (Ornitz et al., 1985). Newer strategies have made use of a tissue‐specific, ‘‘binary’’ gene‐targeting system, in which, for example, one transgenic line expressing Cre recombinase under a tissue‐specific promoter is mated to a second line in which a marker gene (typically GFP) is interrupted by a LoxP‐ flanked (‘‘floxed’’) fragment that is removed by Cre‐mediated recombination (Matsuoka et al., 2005; Zong et al., 2005). The marker gene then becomes activated in only those cells where the promoter driving Cre expression is active regardless of position in the embryo.

III. Limited Information from Cell Marking Collectively, the cell‐marking techniques described above have been informatively employed to analyze cell lineage during development in a wide variety of species and tissues reported in landmark studies, including, for example, in the analysis of hematopoiesis (Lemischka et al., 1986), myogenesis (Eloy‐Trinquet and Nicolas, 2002; Schienda et al., 2006), neurogenesis (Turner et al., 1990; Zong et al., 2005), and neural crest formation (Matsuoka et al., 2005). There is no doubt regarding their importance. However, it should be emphasized that cell‐marking studies do not yield the same amount of information as direct observation. A point not often appreciated is that cell‐marking studies demonstrate which cells are descended from a common ancestor, but cannot reveal the hierarchy of Imaginal disks: LAT, antennal; L1, L2, L3, leg; LGE, genital; LHA, haltere; LLD, labial; LFB, fat body; LGN, gonad; LHI, hindgut; LMA and LMP, anterior and posterior Malpighian tubules; LOE, esophagus; LPV, proventriculus; LRG, ring gland; LVA and LVP, anterior and posterior ventriculus; MH, mouth hooks; SG, salivary gland; TA and TP, anterior and posterior spiracles. [Figures 4 and 10 reproduced from Janning (1978), Copyright Results and Problems in Cell DiVerentiation, with kind permission of Springer Science and Business Media.]

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relationships between cells. The distinction is not trivial. For example, suppose that an embryonic cell is tagged with a marker and that a short while later four cells are now found to be tagged with that marker. While it is clear that all four cells are descended from the original labeled cell, there are actually a minimum of 15 diVerent patterns through which the progenitor cell could have divided to yield those four descendants (Fig. 6). In fact, by extrapolation from a phylogenetic algorithm (Hall, 2004), at a time when there are n cells descended from a single, labeled progenitor cell, the number of potential lineages is described by the equation shown in Table I (adapted from Hall, 2004). Even after just a few cell divisions, producing a small quantity of marked cells, the number of possible diVerent permutations of lineages through which they may be related becomes incomprehensibly large. As a point of reference, there are approximately 40 generations of cell divisions occurring between fertilization and birth during mouse embryogenesis (Frumkin et al., 2005). Note further that this equation does not account for lineages in which there is the death of an intermediary generation, something that commonly occurs during development. For example, in C. elegans,

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apoptosis claims over 10% of somatic cells (Lettre and Hengartner, 2006). Thus, this equation actually understates the number of possible lineages. In spite of the fact that a tremendous number of diVerent possible lineage histories can all lead to the same net result, interestingly, it appears that, in C. elegans and other organisms, evolutionary forces shape the actual lineage to be close to the simplest of all those that are possible (Azevedo et al., 2005). Of course, cell‐marking studies can incorporate more than one type of label [e.g., red fluorescent protein was added to GFP in observations of neurogenesis (Zong et al., 2005)] or can label multiple, diVerent cells [e.g., the fate map of myotomes drew on examination of about 3000 individual mouse embryos (Eloy‐Trinquet et al., 2000)]. Nevertheless, only so many diVerent markers and repeated observations can be made, and for experiments replicated thousands, or even millions, of times, the ‘‘number of particles in the universe’’ divided by a million is still absurdly large—the point being that cell marking reveals surprisingly scant detail about cell lineage in comparison to fate maps produced from simply watching embryos develop.

IV. Stochastic Nature of Development in Higher Organisms Direct observation proved to be a tractable approach for constructing the cell fate map of C. elegans not just because of the worm’s transparency, short duration of embryogenesis, and small number of cells, but also because lineage assignment from one individual worm to the next is invariant (Sternberg and Felix, 1997; Sulston et al., 1983). Given that the number of theoretically possible cell hierarchies grows so precipitously as a function of increasing number of cells, it is perhaps not surprising that in mammals and other more complicated organisms, lineage specification is not completely deterministic, and development is partly stochastic (Sternberg and Felix, 1997).

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The random component to development becomes apparent at two levels of organization. First, the cells of the inner cell mass of early mammalian embryos are pluripotent, and studies employing dye injection (Lawson et al., 1991), retroviral marking (Soriano and Jaenisch, 1986), and mixing of embryonic cells from diVerently pigmented mouse strains (Gardner, 1978; Saburi et al., 1997) indicate that embryonic cells labeled as late as at the gastrulation stage (when the germ layers are formed) mix to such an extent that they can be distributed anatomically almost anywhere later during development. Specific examples of cell mixing at more defined developmental stages include the seeding of hematopoietic stem cells from distinct regions of the yolk sac and embryo via the circulation (Delassus and Cumano, 1996; Orkin and Zon, 2002) or the migration of pigmented melanocytes from the neural crest (Christiansen et al., 2000). Fish also demonstrate this phenomenon (Kimmel and Warga, 1987). Second, cellular diVerentiation is also stochastic. This was first made clear in a subtly elegant study (Till et al., 1964), whose impact took sometime to become appreciated (Levenson and Housman, 1981), of the colony formation potential of hematopoietic stem cells, where the number of colony forming units present in equivalent samples of stem cell colonies did not follow the Poisson distribution expected from sampling errors, but rather demonstrated a gamma distribution indicative of a random process that could be simulated by the Monte Carlo method, wherein the probability of a stem cell diVerentiating was twice as likely as that of dividing. Similar subsequent experiments have confirmed stochastic models of commitment to diVerentiation in melanocytes (Bennett, 1983), neural crest cells (BaroYo and Blot, 1992), lymphocytes (Davis et al., 1993), skeletal myoblasts (Lin et al., 1994), adipocytes (Steinberg and Brownstein, 1982), keratinocytes (Steinberg and Brownstein, 1982), and intestinal goblet cells (Paulus et al., 1993). Some have gone so far as to argue (Arias and Hayward, 2006; Golubev, 1996; Huang et al., 2005; Moller and Pagel, 1998; Rao et al., 2002; Veitia, 2005) that the ultimately stochastic nature of cell fate may be an inescapable consequence of the transcription of genes, their epigenetic regulation, and the networks into which they assemble because the molecular interactions that control them unavoidably reduce to quantum mechanical noise (Monod, 1971; Schrodinger, 1956). It is thus clear that even if a complete cell fate map could be drawn for the trillion or so cells of a particular mammal, in the same way it has been done for the far simpler worm, an element of chance would always remain when using that map to track the course of development in another individual. Breakthroughs in cloning animals provide striking proof (Archer et al., 2003) that genetic identity does not necessarily equate with morphological identity (Fig. 7).

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Figure 7 Cloned cattle (clone is on left) demonstrating diVerences in pigmentation and venous patterning among other things. (Used with permission, Copyright courtesy of Frank Robinson, Lodi, California.)

This is not to say that the randomness itself cannot be constrained through genetic controls. A particularly intriguing mouse mutation is ‘‘disorganization’’ (ds) in which the same locus in the same genetic background yields a dizzying array of birth defects among diVerent individuals (Robin and Nadeau, 2001). A ‘‘two‐hit’’ somatic mutation mechanism, analogous to that occurring with tumor suppressor genes in cancer, has been proposed to explain its remarkably variable expressivity, but this hypothesis awaits validation pending the molecular genetic characterization of the ds gene, which remains uncloned nearly 50 years following its first description. Development in higher animals likely therefore represents a compromise between what can be eYciently scripted in the genome and the eVort required to control the entropic noise intrinsic to its underlying biochemistry. The similarities in embryogenesis between invariantly developing organisms, such as nematodes, and ‘‘indeterminate’’ organisms, such as mammals, may actually be greater than their diVerences. C. elegans probably follows the same basic principles as in other organisms, including definition of cell fate through cell–cell interactions, and it has been argued that the variability in development of higher organisms is merely a consequence of the greater number of cell–cell interactions (Schnabel, 1997). Conversely, there may be circumstances in which even indeterminate organisms use invariant lineages at later points in embryogenesis (Kimmel and Warga, 1986).

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V. A New Approach: Phylogenetic Fate Mapping We (Salipante and Horwitz, 2006), as well as Shapiro and colleagues (Frumkin et al., 2005), have recently and independently proposed a new approach whose goal is to retrospectively generate a single‐cell‐resolution fate map of a mouse or human—opaque organisms with overwhelming numbers of cells whose lengthy embryogenesis is not amenable to the sort of direct observational studies that have proved so useful for the worm. We exploit stochastic processes occurring during development by taking advantage of the fact that nearly every time a cell divides, random mutations arise in its genome. The inheritance of mutations by daughter cells oVers a record of cell divisions in which the order that the mutations have occurred during development reflects a cell’s ancestral embryonic relationships. Cells that exhibit the most closely related patterns of somatic mutation share the most recent common ancestry, analogous to the manner in which species with closely related genomes are separated by less evolutionary time than those containing more divergent sequences. If it were possible to identify the somatic mutations diVerentiating one cell of a multicellular organism from another, then it should be possible to infer how the cells are related to one another. Prior studies along related lines, and leading up to this approach, have included analysis of somatic hypermutation at immunoglobulin loci to deduce relationships between lymphocytes (Michael et al., 2002) and postzygotic methylation patterns at CpG sites to reconstruct the ancestry of intestinal crypt cells (Kim and Shibata, 2004). We envision an expansive and systematic approach, in which we treat the cells of an adult mouse as members of an asexual population, individualized by somatic mutation, yet descended from a common founder (the zygote), and have adapted phylogenetic methods developed originally for evolutionary and microbial population biology in order to trace their embryonic lineage. Deciphering cell fate from inherent genetic mosaicism could oVer a simple and passive means for reconstructing the lineage of any cell, from any location, and at any time, during the development of a single individual. Such a map would include the hierarchical lineage ancestries that cell‐marking studies cannot provide. And while the map generates a probabilistic landscape, such a view is, in fact, compatible with the stochastic processes that underlie the biological basis of development itself. A. Introduction to Phylogenetics ‘‘Phylogenetics’’ refers to the study of evolutionary relatedness among species. Over a span of time, meiotic mutations gradually alter the genetic constitution of organisms, leading to the emergence of new species. Yet, mutations also act as a record of evolutionary relationships in which

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genomes exhibiting closely related sequence patterns are more likely to share a recent common ancestry than those separated by a greater number of changes (Fig. 8A). Because sequences continue to mutate over time, ancestral evolutionary states are usually not available, and correctly assigning past lineage relationships may become problematic when the same mutation has arisen independently more than once (homoplasy), or with reversion of mutation to a previous state (Fig. 8B). A number of phylogenetic algorithms have been developed in order to deduce evolutionary lineages despite these complications. Some, such as the ‘‘neighbor‐joining’’ method, establish relationships based on the arithmetic number of diVerences between taxa. Others, including ‘‘maximum likelihood’’ (Felsenstein, 1981) and ‘‘Bayesian’’ methods (Ronquist and Huelsenbeck, 2003), attempt to fit the data in accordance with an evolutionary model. Each approach to phylogenetic reconstruction has its merits and drawbacks, and the question of which is ‘‘best,’’ with respect to accuracy or reliability, or other metrics, remains a subject of debate (Hall, 2005; Suzuki et al., 2002; Taylor and Piel, 2004). Phylogenetic analyses are typically used to determine the evolutionary lineages of extant genes or organisms based on genomic DNA sequence. Nevertheless, the same algorithms can be applied to deduce relationships within any population in which quantifiable changes occur over time.

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For example, phylogenetic programs have been used to reconstruct the histories and relatedness of languages (Gray and Atkinson, 2003; Gray and Jordan, 2000; Rexova et al., 2006) and written texts (Bennett et al., 2003; Spencer et al., 2004) based on similarities and diVerences in grammar and word usage. Similar cladistic relationships have been inferred for archeological artifacts sharing morphological features (O’Brien et al., 2001), and even among human populations that have been compared with respect to cultural beliefs and customs (Collard et al., 2006). In the same fashion, we have employed phylogenetics to build a cell fate map from mitotic mutations accumulating over the course of development.

B. Somatic Mutations DNA replication errors occur with surprising regularity (Drake et al., 1998), and insertion or deletion mutations at short, repetitive DNA sequences are particularly common (Boyer et al., 2002; Streisinger et al., 1966). In fact, contemporary approaches to human genetic linkage analysis have relied on variable number of tandem repeat (VNTR) polymorphisms in microsatellite markers or, more recently, single nucleotide polymorphisms (SNPs). The meiotic mutations responsible for these polymorphisms occur suYciently often such that most families exhibit distinguishably informative alleles, and it is actually not uncommon to detect newly arising mutations in these markers over the course of the two or three generations observed in a ‘‘genome‐wide screen’’ (Ellegren, 2000). For certain sorts of repetitive sequences, such as the triplet repeats involving genes responsible for neurodegenerative diseases (La Spada, 1997), meiotic mutations are inevitable and occur with certainty. In addition to meiotic instability, repetitive DNA sequences also undergo a high frequency of mitotic mutation (Drake et al., 1998; La Spada, 1997). Mononucleotide repeats appear particularly vulnerable to mitotic error and, in particular, polyguanine repeat sequences (Boyer et al., 2002; Diamant et al., 2004) undergo mitotic mutation at frequencies reported at about 10–4 per cell per generation, and even higher in our experience (Salipante and Horwitz, 2006). The DNA replication fidelity of repetitive sequences is under the control of the DNA mismatch repair system, and deficiency of mismatch repair causes the hereditary nonpolyposis colorectal carcinoma (HNPCC) syndrome with attendant microsatellite sequence instability in tumors (Kunkel and Erie, 2005). From an experimental point of view, the somatic mutation frequency is capable of manipulation via mismatch repair‐deficient genetic backgrounds (Frumkin et al., 2005) or through chemical mutagens inactivating this pathway (such as cadmium; Clark and Kunkel, 2004) or predisposing to

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insertion or deletion (such as acridine‐based intercalating agents; Ferguson and Denny, 1990).

C. Tests in Cell Culture As an initial test of phylogenetic approaches to constructing fate maps, both we and Shapiro’s group determined if it was possible to reconstruct the lineage of cells grown in tissue culture and passaged through single‐cell bottlenecks. Shapiro and coworkers (Frumkin et al., 2005) began by estimating the range of parameters for which phylogenetic reconstruction of embryonic development from somatic mutations should be theoretically possible. In computer simulations assuming 40 cell division generations from zygote to birth and using marker mutation frequencies based on those reported for microsatellite sequences in DNA mismatch repair‐deficient cells, they concluded that cell lineage reconstruction could be completed without errors for any cell in a mouse using fewer than 800 markers. They also noted that ‘‘there is not much point in performing tedious and time‐consuming analysis’’ and, in fact, demonstrated experimentally that they could actually reconstruct the lineage ‘‘trees’’ of mismatch repair‐deficient human adenocarcinoma cells grown in tissue culture with perfectly accurate branching orders by genotyping as few as between 20 and 40 microsatellite repeat markers, using the neighbor‐joining phylogenetic algorithm. We performed similar studies (working independently of them and before their work became known to us through publication) in which we took a single NIH3T3 mouse embryonic fibroblast, allowed it grow for 22 population doublings (by repeatedly ‘‘splitting’’ the cells on a single tissue culture plate), and then seeded a single cell from this population on a second generation of plates, and so on, in order to construct a branched tree, grown over approximately 4 months time, where the real phylogeny was known to us (Fig. 9A). At the end of each generation, we collected the cells from each plate, extracted the DNA, and ‘‘genotyped’’ multiple diVerent marker sequences to determine if they had undergone mitotic mutations capable of delineating the relationships between how the diVerent plates of cells had been passaged. Our experiments diVered from Frumkin et al. (2005) in the following respects: (1) We used mouse cells instead of human cells. (2) The cells we used are thought to have an intact DNA mismatch repair system; so to compensate for a lower mutation frequency, we genotyped markers containing polyguanine repeat sequences because they undergo mitotic mutations at high frequency. In fact, we observed a mutation frequency for some markers as high as about 10–2 mutations per cell per generation. (3) Our cells grew for a constant number of divisions (22) between generations,

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whereas Frumkin et al. (2005) grew the cells for as many as 127 cell divisions between each generation. (The longer the cells grow, the more likely it is that mutations will accumulate.) (4) We used a Bayesian approach for phylogenetic reconstruction, whereas Frumkin et al. (2005) used a neighbor‐joining method. Nevertheless, our results were in good agreement, and we obtained slightly less than perfect topological accuracy by genotyping 28 polyguanine markers (Fig. 9B), not all of which displayed length polymorphisms in every sample. At first glance, the reconstruction appears somewhat diVerent from the known phylogeny, but it is worth emphasizing that the genetic data infers relationships between isolates at greater detail than could be known to us while actually growing the cells in tissue culture. For example, among some related isolates (such as A1–A7), the reconstruction has introduced bifurcations that more accurately portray the sequence of clonal expansion of a progenitor cell within a plate of cells. It is also possible to use the mutation frequency in order to provide a scale calibrated to the number of cell divisions. During embryonic development, intermediate lineages may be lost

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through apoptosis or may otherwise be unavailable to sample in the same way that cells can be while growing in tissue culture. We therefore also determined if it is possible to accurately reconstruct the tissue culture lineage by performing the genetic analysis on just the final generation. The actual tree is a bit simpler when represented this way (Fig. 9C) and the phylogenetic reconstruction, even in the absence of information regarding all the preceding generations, is still fairly correct (Fig. 9D) and introduces just one error (at H1–H2) and one polytomy (at E1–E2). Cells grown in tissue culture are generally regarded as clones of one another; nevertheless, these experiments conclusively demonstrate that somatic mutation generates suYcient genetic diversity to allow for delineation of an in vitro ancestry spanning over a period of time as short as that of just a few days.

D. Mapping Lineages in Whole Organisms Frumkin et al. (2005) next applied this approach to mapping cell lineages in plants, which may be of particular interest for validation purposes, because, conceivably, there is less cell migration than in animals and the branching of the phylogenetic ‘‘tree’’ should therefore parallel the actual branches of the plant specimen under study. They initially surveyed one microsatellite locus in 28 tissue samples taken from a black locust tree and interestingly found that three of the samples shared an allele with a distinguishable number of repeats diVerent from the other 25 samples and that these 3 samples resided adjacently along the same terminal branch of the tree’s limb. Apparently because genomic DNA sequences necessary for identifying other microsatellite repeats in this species were lacking, they pursued more extensive studies in a strain of the plant Arabidopsis thaliana deficient for the DNA mismatch repair gene, MSH2. They examined 23 tissue samples and genotyped 22 microsatellite markers. Although they were able to statistically significantly correlate genetic diVerences with the branching anatomy of the plant, they apparently lacked suYcient information to produce a phylogenetic reconstruction of the cellular lineages within the plant. One problem with extracting DNA from patches of tissues is that a mutation would need to occur early enough developmentally to be present in the majority of the sampled cells, thus limiting the power to map cell fate to earlier points in development. A second issue is that a sample is likely to be composed of a heterogeneous population of diVerent types of tissues arising from diVerent germ layers and with distinct developmental histories. In order to circumvent these diYculties, it is necessary to determine the somatic genotype of single cells from an organism. This also presents a challenge, however, because a single cell contains just one copy of the genome

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that can obviously only be used for just a single PCR genotyping reaction. Fortunately, the amount of available template DNA can be dramatically increased through the use of ‘‘whole genome amplification’’ (Stoecklein et al., 2002). We (Salipante and Horwitz, 2006) therefore isolated single cells from diVerent tissues of an individual adult mouse and subjected each cell to whole genome amplification. We examined each cell microscopically so that we could reasonably confidently assign histology. We collected 84 mouse cells, most of which were hepatocytes from the liver, and then genotyped each cell for 31 diVerent polyguanine markers. It is important to note that no two cells had identical genotypes for all 31 markers; thus, every cell had become individualized by somatic mutations over the course of development. While algorithms could construct a phylogeny for all 84 cells, there were branches where the statistical confidence of the predicted tree was low or where there was inadequate information to infer a branching order (resulting in a ‘‘polytomy’’). However, when we selected the approximately half of those cells exhibiting the most divergent genotypes, we were able to use Bayesian methods to reconstruct an embryonic phylogeny exhibiting reasonably confident assignment of branching order (Fig. 10). An intriguing feature of this ‘‘proof‐of‐principal’’ fate map is that there are two main clades of hepatocytes, determined to 99% confidence (‘‘prior probability’’ in Bayesian terms), each predominantly clustering in the same anatomic regions, and only one of which has the potential to contribute to cells representing lineages sampled from other organs. Accordingly, a recently constructed fate map of embryonic liver development (Tremblay and Zaret, 2005), using the canonical approach of dye injection of single cells, interestingly determined that in cultured mouse embryos, in contrast to nonmammalian vertebrates, two distinct types of endoderm progenitor cells, lateral and medial, arise from spatially separated embryonic domains and converge to generate the embryonic liver bud. While our approach awaits rigorous validation, it does appear capable of returning biologically meaningful results.

VI. Challenges Dealing with Randomness in Development So far, we have applied this approach to just a single individual. Yet unlike the nematode’s deterministic embryogenesis, mammalian development exhibits stochastic elements, and much of embryogenesis is marked by seemingly random cell mixing and migration and commitment to diVerentiation. Thus, fate maps of individual organisms may vary in several respects. There may be dissimilar contributions of precursor cells to tissues, resulting in diVerent lineage relationships for cells that comprise those tissues (Fig. 11A). Individuals may exhibit identical embryonic cell lineages, but alternative fate decisions may result in the production of diVerent cell types from all of, or

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Salipante and Horwitz R. Cerebellum-1 R. Cerebellum-2 Neuron R. Cerebral cortex L. Cerebellum-1 L. Kidney-1 L. Kidney-2 Renal tubular epithelial cell R. Kidney-1 R. Kidney-2 Cardiac myocyte Heart, L. Ventricle L. Parotid Serous cell 0.56 R. Parotid Liver, L. Lobe-10 0.75 Liver, L. Lobe-11 Liver, L. Lobe-6 0.88 Liver, caudal lobe-7

10 Cell divisions

Liver, caudal lobe-6 Liver, R. Lobe-6 Liver, R. Lobe-4 0.93 0.66 Liver, R. Lobe-7 0.63 Liver, R. Lobe-3 0.98 Liver, R. Lobe-1 Liver, R. Lobe-9 Liver, R. Lobe-2 Liver, R. Lobe-8 Liver, M. Lobe-8 Liver, M. Lobe-9

0.72 0.99

Liver, caudal lobe-4 Liver, caudal lobe-2 Liver, R. Lobe-5 0.60 0.98

Liver, caudal lobe-3 Liver, caudal lobe-1 Liver, L. Lobe-1 Liver, L. Lobe-5

0.79

0.99

Hepatocyte

Liver, L. Lobe-9 Liver, M. Lobe-6

0.89

Liver, M. Lobe-7 0.87

0.56

0.57 0.84

Liver, M. Lobe-5 Liver, L. Lobe-4 Liver, L. Lobe-3 Liver, L. Lobe-2 0.64 Liver, L. Lobe-8 Liver, M. Lobe-4

0.82

Liver, L. Lobe-7 Liver, M. Lobe-2 Liver, M. Lobe-1 Liver, M. Lobe-3

Figure 10 Cell fate map showing lineage relationships between diVerent types of cells sampled from diVerent anatomic regions of an adult mouse. Each terminal node is a single cell. Cells taken from the same site are denoted by a dash followed by a number. The shading corresponds to laterality and subdivisions within each organ. Posterior probabilities for the Bayesian reconstruction (which can be conceived as confidence values) are indicated alongside their corresponding nodes. R., right; L., left. [Reproduced from Salipante and Horwitz (2006), used with permission, Copyright National Academy of Sciences.]

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6. Phylogenetic Fate Maps A A B 1 2 3 4 5 6 7 8

Consensus fate map

Individuals

A B 1 2 3 4 5 6 7 8

B

A B 1 2 3 4 5 6 7 8

A B 1 2 3 4 5 6 7 8

C A B 1 2 3 4 5 6 7 8

A A or B B 1 2 3 4 5 6 7 8

A B 1 2 3 4 5 6 7 8

A B 1 2 5 6 3 4 7 8

A A or B B 1 2 3 4 5 6 7 8

Figure 11 Examples of stochastic diVerences in the development of individual organisms. Terminally diVerentiated cells (1 through 8) comprise tissues A and B. (A) DiVerences in embryonic lineages of cells that compose diVerent tissues. (B) Conserved cell lineages exhibiting diVerences in cellular diVerentiation. (C) DiVerences in early cell migration resulting in diVerences in terminal diVerentiation patterns.

portions of, those lineages (Fig. 11B). Patterns of cell migration could also vary, leading to discrepancies between cell lineage and terminal cell fate based on position eVects (Fig. 11C). In spite of these issues, it may be possible to combine information across individuals in order to produce a fate map representative of a species as a whole. A ‘‘consensus’’ fate map could be constructed by combining information from separate organisms into a single phylogenetic reconstruction. A fate map derived in this way would reflect the probability that a particular embryonic lineage will pursue a particular cell fate, a view of development that is more compatible with the actual events of mammalian embryogenesis than is a single fate map derived from examination of just one individual (Lawson et al., 1991). By comparing and contrasting the fate maps of individual organisms, it should additionally be possible to identify precisely what cell lineage pathways are variable and which are conserved throughout development.

VII. Extended Applications of Phylogenetic Fate Mapping Phylogenetic fate maps can be constructed using cells from a single individual, and because lineage information is carried by the genome itself, fate maps may be built, retrospectively, at any point during the lifecycle of that organism. These properties make the approach unique among current methods for cell fate analysis, and also oVer new opportunities to study cellular lineages involved in biological processes distinct from development, including cancer, stem cell populations, wound healing, tissue repair and maintenance, aging, adult neurogenesis, and learning.

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Acknowledgments This study is supported by NIH grants R01DK58161 (M.S.H.), T32GM007266 (S.J.S., Medical Scientist Training Program), and Poncin Scholarship Fund (S.J.S.). We thank Dr. Galya Diment for the reference on Nabokov and the many individuals and journals noted in the figure legends who graciously allowed us to reproduce figures.

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Index A Abdominal aortic aneurysms (AAAs), 100, 103, 117–119 neovascularization and aging in, 122–124 pathophysiology of, 118 Adenosine, in CBF regulation, 85 Adenoviral expression, of Cux1, 8 AER. See Apical ectodermal ridge Airway smooth muscle (ASM), 61–65, 67–70 AKT-dependent mechanism, 122


Aminobutyric acid (GABA), 76, 79 Amphiarthrosis, 2 Aneurysm model, 119–120 ANF. See Atrial naturitic factor Angiogenesis, 118–123, 131–133, 137–138 Angiotensinogen, 62 Anidjar/Dobrin rat model, 118 Anti Mullerian hormone, 44, 50 Antisense probes, and wnt gene in situ hybridization, 9–10, 14 Apical ectodermal ridge (AER), 6 Arterial diseases, elastin fragmentation, 117–118 Arthritis, 1, 3 Articular cartilage, 1–5, 20, 22–28 development of, 22–23 ASM. See Airway smooth muscle ASM hyperplasia, 68 Astrocyte-neuron lactate shunt, 81 Astrocytes anatomical position of, 77 Ca2þ signaling, 78 calcium elevation in, 80 excitatory amino acid transporters (EAAT), 81 G-protein–coupled receptors activation, 79 glutamate release and synaptic transmission, 80 glutamate-dependent Ca2þ elevation, 79 as sensors of synaptic activity, 82 sodium-dependent glutamate uptake in, 75 Astrocytic Ca2þ elevations, in neurovascular coupling, 75, 78–82, 86, 90

Astrocytic endfeet, Kþconductance, 77, 83–84 Astrocytic foot processes, 76, 81–82 Atrial naturitic factor (ANF), 64 B ‘‘Bayesian’’ methods, 172 BMP receptor 1a (BMPr1a), 23–24 Bone morphogenetic protein (BMP), 11 Brachydactyly type A1, 7 Brachypodism, (gdf5 Au3 null), 7, 11 Brain blood flow, 77 regulation of synaptic transmission, 78–79 slices studies, 82–83 Brain mechanisms involved in lordosis behavior action of opioid peptide enkephalin, 43, 45 anterior pituitary gland, 42 dual roles LHRH (GnRH) neurons, 42 MIS type I receptor, 50–53 MIS type II receptor, 45–53 Mullerian Inhibiting Substance (MIS), 44–52 neuropeptides, 43, 46 opioid peptide enkephalin, action of, 43, 45 ovulatory surge of luteinizing hormone, 42 progesterone (P) receptor and ir cells density, 44, 42 sexual diVerentiation of, 41–42 testosterone and metabolites, 42–43 in sexual behavior, 53–54 C Ca2þ in astrocytic endfeet photolysis, 83 channel for PKCa activation, 18 chelators, 19 waves in astrocytes, 79, 82–83

185

186 Caenorhabditis elegans, cell fate map, 159–160 CAM model, 121 cAMP degradation, 63 cAMP response element (CRE), 62 cAMP response element-binding protein (CREB), 62 cAMP synthesis, 63 cAMP-PDE signaling cascade in ASM, 68. See also Airway smooth muscle Cartilage diVerentiation, 12 Cartilage formation, role of tenascin-C, 20 Cartilage oligomeric matrix protein (COMP), 20. See also Joint formation -Catenin, 10–11, 128 CD44rel, 10 CD8þ TIL, cytolytic activity, 134 Cell culture tests, for construction of fate map, 174–176 Cell fate map construction, 159 adult mouses, 178 Cell marking and genetically engineered markers, 162–165 and gynandromorphs, 161–162 Cerebral blood flow (CBF), 76, 78, 84–87, 90 and coupling of neuronal activity, 86 role of astrocytes in regulation of, 82 role of endothelial cells in, 77 CFTR. See Cystic fibrosis transmembrane conductance regulator Chondrocytes (chondrification), 4, 6–7, 10 Chondrogenesis, 19–21, 27 Chondrogenous cells, 2, 23, 28 Chordin, 5, 10–11 Chorioallantoic membrane model, 120 Chronic obstructive pulmonary disease (COPD), 64, 68–69 CNS arousal states, 37 Col2a1 promoter/enhancer, 11 Collagen expression, 23 model, 121 type II (Col2), 5, 9 type III, 9 Connexin 43 CREB-binding protein (CBP), 62 Cyclooxygenase (COX) inhibitors, 83 Cyclooxygenase-2 (COX-2) metabolites, 75, 77

Index Cysteine proteases in elastolysis, 109 Cystic fibrosis transmembrane conductance regulator (CFTR), 62 Cystine accumulation in astrocytes, 80 Cystine/glutamate exchanger, 80 ‘‘Cysteine switch’’ motif, 101 Cytokines, (IL-6, 8, 10), 69–70 D Diacylglycerol (DAG), 79, 87 Drosophila cut gene, Cux1, 8 fate map, 164 E EAAT. See Excitatory amino acid transporters EBP. See Elastin-binding protein EDHF. See Endothelium-derived hyperpolarizing factor EETs mitogenic eVect, in gial cells, 89 Elastases in human, 106 in inflammatory cells, 107–108 Elastic fibers development, 105 formation and degradation, 103–111 fragmentation and aging, 103 Elastin content of, 103–104 degradation, 99 degradation mechanism by elastases, 110–111 gene expression of, 104 longevity, 103 mediated angiogenesis, 120–121 synthesis of, 104 Elastin-binding protein (EBP), 105, 113–116, 120 Elastin peptides (EP), 120–121, 123–125, 128–129, 135–137 biological activities of, 111–116 biological eVects of, 112 proangiogenic eVect of, 118–119 receptor system, 111–114 Th-1 polarization, 126 Elastolysis, 117–126 Embryonic cartilage, 4, 22 Endochondral ossification, 11, 22 Endothelial NO synthase (eNOS), 77

187

Index Endothelin, vasoconstrictor agents, 77 Endothelium-derived hyperpolarizing factor (EDHF), 77 EP. See Elastin peptides EP eVect on Th-1/Th-2 orientation in melanoma, 135 Epiphyseal chondrocytes, 12. See also Chondrocytes Epoxyeicosatrienoic acid (EETs), 75, 77, 87 Estrogen receptor (ER), 38, 42–43 actions in hypothalamic neurons, 38 Eudiarthrosis joints, 2 Exchange protein directly activated by cAMP (Epac), 63 Excitatory amino acid transporters (EAAT), 81

F fgf8 expression, 6 Fibrillin-1, 105 Fibroblast growth factor 10 (fgf10), 6 Fibrocartilage joints, 2 Fibulin-5, 105 Flaccid paralysis, 16–17 Frameshift mutations, 11 Frizzled (Fz) gene family, 10, 65 Functional brain imaging techniques, 76 Functional magnetic resonance imaging (fMRI), 76

G Gap junctions, 78–81, 86 GEFs. See Guanine-nucleotide exchange factors Gdf5 gene, 5 expression, analysis of, 1, 11 Gdf5 homologues, contact, 13 Gdf5:gdf6 double mutant, 12 Gdf6 gene, 12 Gdf6 null mutant mice, 12 gdf7 gene, 12 Gdf7 mutation, in mice, 12 Gelatinase A gene, 109 Generalized arousal of CNS biophysical and molecular mechanisms for, 54 histamines and NDMA, 54–55 Generalized arousal transmitters and hormone, interactions, 55

Genes properties ( ERand ER ), 38, 41 Glial cells classification in CNS, 76 Glucose transporter (GLUT)-1, 81 G-proteins, 67, 115 Green fluorescent protein (GFP), 162 Growth factor gdf5, 9 Growth factor signaling, 1 Guanine-nucleotide exchange factors (GEFs), 63 H HA in joint cavitation, importance of, 15 HA synthase (HAS), isoforms, 18 HA synthesis, translational inhibitors, 18 HAbinding proteins (HABP), 15 HAS genes, 18–19 Hemiarthrosis joints, 2 Hemichannels, 78 Higher organisms, nature of development, 168–169 Histamine H1 receptor antagonism, 54 Homeodomain, transcription factors, 7 Hox genes, 7–8 Hox homeobox genes, 8 Hoxa9-13 and Hoxd9-13, gene expression patterns of, 7–8 Human matrilysin (MMP-7) gene, 109 Hyaline articular cartilage, 2 Hyaluronan (HA), 5, 16–17 20-Hydroxyeicosatetraenoic acid (20-HETE), 83 Hyperphalangy, 9 Hypertrophy, 5, 7 I Indian hedgehog (ihh), 7 Inflammatory aneurysms, 118 Inositol-1,4,5-trisphosphate (IP3), 79 Interferon- (IFN- ), 70, 101 Interspecies chimeras, mapping development, 159 Interzone specific gene products, 5, 11 Intrinsic histone acetyltransferase activity, 62 Isobutyl methyl xanthum (IBMX), 65 J Joint biology, postnatal BMPr1b and BMPs2/4 signaling, 24

188 Joint biology, postnatal (cont.) collagen type II expression, 24 cre recombinase, 23 gdf5 promoter, 23 levels of aggrecan, 24 Notch-signaling protein family, 24, 26, 28 progenitor/stem cell population, 24, 27 transforming growth factor, 24 Joint cavitation and exogenous HA oligosaccharides, 15 CD44 levels, 15 eVects of mechanical stimuli on developing joints, 15, 17 embryonic limb movement, 14 extracellular-signal-related kinases (ERK1/2), 19 formation, 5 gdf5 protein expression, 16 HA, unsulfated GAG, 15, 18 HA–HABP interactions, 15 hyaluronan-related events during, 16 local extracellular matrix (ECM) composition, 14 and mechanical stimuli, 13, 15 and mechanical strain, 16–17 mitogen-activating protein kinase (MAPK) signaling, 18–19 uridine diphosphoglucose dehydrogenase (UDPGD), 15 Joint formation cartilage oligomeric matrix protein (COMP), 20 cell–matrix interactions, 19–22 ECM components, 19 fibronectin, an ECM glycoprotein, 21 glycoproteins, tenascin-C, 19–20 integrins, ECM cell surface receptors, 22 syndecan-3, role of, 21 thrombospondins adhesive glycoproteins, 20 Joints immobilization in ovo, 13 Joint interzone formation, 9–13 Joint patterning and specification, 5 L Ligand-activated transcription factors, 38 LIM homeodomain transcription factor, Lmx1, 6 Limbic-hypothalamic system of estrogenbinding neurons, 38

Index Lordosis behavior, 37–44, 53–54 brain mechanisms involved in action of opioid peptide enkephalin, 43, 45 anterior pituitary gland, 42 dual roles LHRH (GnRH) neurons, 42 MIS type I receptor, 50–53 MIS type II receptor, 45–53 Mullerian Inhibiting Substance (MIS), 44–52 neural circuit for, 40 neuropeptides, 43, 46 opioid peptide enkephalin, action of, 43, 45 ovulatory surge of luteinizing hormone, 42 progesterone (P) receptor and ir cells density, 44, 42 sexual diVerentiation of, 41–42 testosterone and metabolites, 42–43 neuropeptides and neurotransmitters, 46 LRP5/6, 10 M MAGP-1, 105 MAP-kinase pathway, 19 Marfan syndrome, 105 Matrigel model, 121 Matrikines, 102–103. See also Proangiogenic matrikines Matrilysin (MMP-7), 101 Matrix metalloproteinases (MMPs), 100 homologies of, 101 as matrix decryptases, 102 regulation of, 101 Mechanical stimuli, role of, 13. See also. Joint cavitation Meckel’s cartilage,hyperplasia of, 8 MEK/Erk pathway, 115–116 Melanoma and elastin, 127–135 Membrane-type matrix metalloprotease-1 (MT1-MMP), 109 MEROPS database, 101 Mesenchymal condensation, 5 Metalloproteinases, expression of, 23 Metobotropic glutamate receptors (mGluR), 78 MIS. See anti Mullerian hormone MMP-1 gene transcription, 115 Morphogen sonic hedgehog (shh), 6

189

Index Mosaic organisms and gynandromorphs, 163 Mutant digits, 7 Mutations in ihh, 7 N ‘‘Neighbor-joining’’ method, 172 Neural circuit for lordosis behavior, 40 Neurometabolic coupling, 81 Neuromuscular blockers, 16 Neuron–Astrocyte interactions and gap junctions, 78–79 Neuronal NO synthase (nNOS), 77–78 Neuropeptides on lordosis behavior, 46 Neurotransmitters on lordosis behavior, 46. See also Lordosis behavior Neurovascular coupling role of astrocytes, 82–89 vasoactive mediators of, 84 Neurovascular unit, 76–78 N-methyl-D-aspartate (NMDA) receptor, 80 Noggin, 5, 9, 13 Nonandrogenic genetic influence, sexual diVerentiation, 37 Notch-signaling pathway, 25. See also Joint biology O Organisms mapping lineages in, 176 Osteoarthritis, 2 Osteoclast-specific transcription factor, 117 P p110 phosphatidylinositol-3 kinase (PI3K), 116 Perivascular endfeet, 75, 82–83 Phosphodiesterases (PDEs) and ASM growth and migration, 68–69 and ASM tone and contractility, 67–68 calcium/calmodulin-dependent PDE, 64 cAMP and cGMP isoforms, 64–65 chemokine secretion, 69–70 and cilomilast, 69 classification of, 64 cytokine secretion, 69–70 developmental and physiological responses, 65–67 degradation of cAMP in ASM cells, 62 as regulators of cAMP,cGMP-activated signaling pathways, 62

isoforms in ASM, 68 in regulating ASM function, 61 inhibition abrogates cytokine-induced eVects in ASM, 63 gene family, 64 modular structure of, 64 and nonselective inhibitor, 65 Phospholipase A2 (PLA2) inhibitor, 83 Phospholipase C (PLC), 79 Phylogenetic algorithm of tagged cell, 167 fate mapping, 171–177, 179 introduction, 171–173 somatic mutations in polyguanine sequences, 175 Phylogeny of NIH3T3 mouse, 175 PI3K/AKT/eNOS/Erk1/2 signaling, 120 PKC activity, HA release, 18 Polymorphonuclear neutrophils (PMN), 118 Potassium channel, cardiac pacemaker cells, 63 Prechondrogenic mesenchyme, 10 Proangiogenic matrikines, 119–120 Prostaglandin E2 (PGE2), 69, 83 Prostaglandin F2 (PGF2), 77 Protein kinase C (PKC), 18, 43, 115 Proteoglycans, 21, 23, 102, 105 Pthrp expression domains, 7 Purinergic P2X receptors, 80

R Ras-Raf-MEK1/2-Erk1/2, phosphorylation cascade, 115 Regulated on activation, normal T-cell expressed and secreted (RANTES), 70, 102 Reactive nitrogen species (RNS), 100 Reactive oxygen species (ROS), 100 Rheumatoid arthritis, 2

S S-Gal complex, 114 S-Gal occupancy, cell signaling pathways, 115–116 Schizarthrosis joints, 2 Secondary cartilaginous fusion, 5 Selective inhibitors development, PDE, 68

190 Sexual diVerentiation behavior development of, 37 molecular mechanisms, 39 Short DNA sequences phylogenies, 174 Smooth muscle cells (SMC), 64, 77, 112, 115–116, 136 Somatic mutations in polyguanine sequences, phylogeny construction, 175 Somatopleural mesoderm, 6 Somitocoele cells, 9 Sonic hedgehog (shh), 6 sox9 expression, 5, 11 Spino-midbrain-spinal neural circuit, 38 Switch-oV enzyme latency, 101 Synarthrosis, 2 Synovial joint fluid, 1–3, 5 formation, inhibition, 12 membrane, 2 morphogenesis, interzone, 5 tissues, 2 T T-box transcription factors, 6 T-Lymphocytes polarization and elastin peptides, 124–125 Transforming growth factor, 24 Transgenic marker, fate map, 166 Tripartite synapse, 77, 80 Tromboxan A2 (TXA2), 77

Index Tropoelastin molecules coacervation of, 103–105 sequences of, 113 synthesis and association, 103–105 ‘‘Two-hit’’ somatic mutation mechanism, 170 Tumor necrosis factor- (TNF), 62, 66, 70, 101 Tumor-infiltrating lymphocyte (TIL), 134, 135 U Uridine diphosphoglucose dehydrogenase (UDPGD), 15. See also Joint cavitation V Vasomotion, 82–83, 87, 90 W wnt family members, 6, 9–10 wnt14 adenovirus, micromass assays, 10 wnt14 and gdf5, expression of, 10, 12 wnt14 (Wnt9a) gene expression, 9 Z Zebra fish gdf5 (contact) expression, 13–14 Zone of polarizing activity (ZPA), 6–7

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-Franc¸ois 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

191

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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

Contents of Previous Volumes

193

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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Contents of Previous Volumes

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

Contents of Previous Volumes

195

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 Joaquı´n 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

Contents of Previous Volumes

197

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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Contents of Previous Volumes

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

Contents of Previous Volumes

199

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

Contents of Previous Volumes

201

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

Contents of Previous Volumes

203

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

Contents of Previous Volumes

205

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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Contents of Previous Volumes

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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209

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-Franc¸ois 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

Contents of Previous Volumes

211

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

Volume 76 1 BMP Signaling in the Cartilage Growth Plate Robert Pogue and Karen Lyons

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

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

4 Wnt Signaling: A Key Regulator of Bone Mass Roland Baron, Georges Rawadi, and Sergio Roman-Roman

5 Eukaryotic DNA Replication in a Chromatin Context Angel P. Tabancay, Jr. and Susan L. Forsburg

6 The Regulatory Network Controlling the Proliferation–Meiotic Entry Decision in the Caenorhabditis elegans Germ Line Dave Hansen and Tim Schedl

7 Regulation of Angiogenesis by Hypoxia and Hypoxia-Inducible Factors Michele M. Hickey and M. Celeste Simon

Volume 77 1 The Role of the Mitochondrion in Sperm Function: Is There a Place for Oxidative Phosphorylation or Is this a Purely Glycolytic Process? Eduardo Ruiz-Pesini, Carmen Dı´ez-Sa´nchez, Manuel Jose´ Lo´pez-Pe´rez, and Jose´ Antonio Enrı´quez

Contents of Previous Volumes

213

2 The Role of Mitochondrial Function in the Oocyte and Embryo Re´mi Dumollard, Michael Duchen, and John Carroll

3 Mitochondrial DNA in the Oocyte and the Developing Embryo Pascale May-Panloup, Marie-Franc¸oise Chretien, Yves Malthiery, and Pascal Reynier

4 Mitochondrial DNA and the Mammalian Oocyte Eric A. Shoubridge and Timothy Wai

5 Mitochondrial Disease—Its Impact, Etiology, and Pathology R. McFarland, R. W. Taylor, and D. M. Turnbull

6 Cybrid Models of mtDNA Disease and Transmission, from Cells to Mice Ian A. Trounce and Carl A. Pinkert

7 The Use of Micromanipulation Methods as a Tool to Prevention of Transmission of Mutated Mitochondrial DNA Helena Fulka and Josef Fulka, Jr.

8 Difficulties and Possible Solutions in the Genetic Management of mtDNA Disease in the Preimplantation Embryo J. Poulton, P. Oakeshott, and S. Kennedy

9 Impact of Assisted Reproductive Techniques: A Mitochondrial Perspective from the Cytoplasmic Transplantation A. J. Harvey, T. C. Gibson, T. M. Quebedeaux, and C. A. Brenner

10 Nuclear Transfer: Preservation of a Nuclear Genome at the Expense of Its Associated mtDNA Genome(s) Emma J. Bowles, Keith H. S. Campbell, and Justin C. St. John

Volume 78 1 Contribution of Membrane Mucins to Tumor Progression Through Modulation of Cellular Growth Signaling Pathways Kermit L. Carraway III, Melanie Funes, Heather C. Workman, and Colleen Sweeney

2 Regulation of the Epithelial Naþ Channel by Peptidases Carole Plane`s and George H. Caughey

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3 Advances in Defining Regulators of Cementum Development and Periodontal Regeneration Brian L. Foster, Tracy E. Popowics, Hanson K. Fong, and Martha J. Somerman

4 Anabolic Agents and the Bone Morphogenetic Protein Pathway I. R. Garrett

5 The Role of Mammalian Circadian Proteins in Normal Physiology and Genotoxic Stress Responses Roman V. Kondratov, Victoria Y. Gorbacheva, and Marina P. Antoch

6 Autophagy and Cell Death Devrim Gozuacik and Adi Kimchi