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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
Contents
Contributors
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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 I. II. III. IV. V. VI.
Introduction 2 Fluid Compartments of the Brain 3 The CP-CSF System and the Development of the CNS The CP-CSF System in Adulthood 16 Senescence of the CP-CSF System 24 Perspectives 31 Acknowledgments 35 References 35
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2 Zebrafish Genetics and Formation of Embryonic Vasculature Tao P. Zhong I. II. III. IV. V. VI.
Introduction 53 The Origin and Specification of Angioblasts 55 DiVerentiation of Arterial and Venous Endothelial Cells 59 Assembly of the Dorsal Aorta and the Cardinal Vein 69 Patterning of the Intersegmental Vessels 71 Perspectives 74 Acknowledgments 75 References 75
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3 Leaf Senescence: Signals, Execution, and Regulation Yongfeng Guo and Susheng Gan I. II. III. IV. V.
Introduction 84 Leaf Senescence-Regulating Signals 85 The Cell Death Execution Process 91 Gene Expression and Regulation During Leaf Senescence Closing Remarks 102 Acknowledgments 103 References 103
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4 Muscle Stem Cells and Regenerative Myogenesis Iain W. McKinnell, Gianni Parise, and Michael A. Rudnicki I. II. III. IV.
Developmental Origin of Muscle 113 Postnatal Muscle Repair 114 Adult Stem Cells in Skeletal Muscle 116 Interpretation of Findings from Models of Extreme Muscle Damage 125 References 127
5 Gene Regulation in Spermatogenesis James A. MacLean II and Miles F. Wilkinson I. II. III. IV.
Introduction 132 Hormonal Regulation of Transcription in the Testis 135 Transcription Factors with Testicular Functions 153 Testis-Specific Gene Expression and DNA Methylation 170 Acknowledgments 181 References 181
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6 Modeling Age-Related Diseases in Drosophila: Can this Fly? Kinga Michno, Diana van de Hoef, Hong Wu, and Gabrielle L. Boulianne I. II. III. IV. V. VI. VII. VIII.
The Challenge of Studying Age-Related Diseases 200 Why Study Human Neurodegenerative Diseases in Drosophila? 200 Huntington’s Disease and Polyglutamine Disorders 203 Parkinson’s Disease 207 Alzheimer’s Disease 208 Amyotrophic Lateral Sclerosis 212 What Can We Do with a Fly Model of Neurodegenerative Disease? 215 The Next Frontier 216 Acknowledgments 217 References 217
7 Cell Death and Organ Development in Plants Hilary J. Rogers I. II. III. IV. V.
Introduction 226 Seed and Embryo Development 228 Leaf, Stem, and Root Development 236 Flowering and Reproduction 242 Conclusions and Future Directions 247 Acknowledgments 253 References 253
8 The Blood-Testis Barrier: Its Biology, Regulation, and Physiological Role in Spermatogenesis Ching-Hang Wong and C. Yan Cheng I. II. III. IV. V.
Introduction 264 The Molecular Architecture of the BTB Models to Study BTB Dynamics 267 Regulation of BTB Dynamics 270 Concluding Remarks 285 Acknowledgments 286 References 286
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9 Angiogenic Factors in the Pathogenesis of Preeclampsia Hai-Tao Yuan, David Haig, and S. Ananth Karumanchi I. II. III. IV.
Introduction 297 Abnormal Placentation and Placental Ischemia (Stage 1) Systemic Endothelial Dysfunction (Stage 2) 300 Conclusions 306 Acknowledgments 308 References 308
Index 313 Contents of Previous Volumes
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Contributors Numbers in parentheses indicate the pages on which the authors’ contributions begin.
Gabrielle L. Boulianne (199), Program in Developmental Biology, The Hospital for Sick Children, Toronto, Ontario, Canada M5G 1X8; Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada M5G 1X8 C. Yan Cheng (263), Population Council, New York, New York 10021 Adam Chodobski (1), Department of Clinical Neurosciences, Brown University School of Medicine, Providence, Rhode Island 02903 John A. Duncan (1), Department of Clinical Neurosciences, Brown University School of Medicine, Providence, Rhode Island 02903 Susheng Gan (83), Cornell Genomics Initiative and Department of Horticulture, Cornell University, Ithaca, New York 14853-5904 Yongfeng Guo (83), Cornell Genomics Initiative and Department of Horticulture, Cornell University, Ithaca, New York 14853-5904 David Haig (297), Department of Organismic and Evolutionary Biology, Harvard University, Cambridge, Massachusetts Diana van de Hoef (199), Program in Developmental Biology, The Hospital for Sick Children, Toronto, Ontario, Canada M5G 1X8; Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada M5G 1X8 S. Ananth Karumanchi (297), Renal, Molecular, and Vascular Medicine Division, Departments of Medicine, Obstetrics and Gynecology, Beth Israel Deaconess Medical Center and Harvard Medical School, Boston, Massachusetts James A. MacLean II (131), Department of Immunology, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030 Iain W. McKinnell (113), Ottawa Health Research Institute, Ottawa, Ontario, K1H 8L6 Canada Kinga Michno (199), Program in Developmental Biology, The Hospital for Sick Children, Toronto, Ontario, Canada M5G 1X8; Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada M5G 1X8
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Gianni Parise (113), Ottawa Health Research Institute, Ottawa, Ontario, K1H 8L6 Canada Jane E. Preston (1), Institute of Gerontology, King’s College London, London, SE1 9NH United Kingdom Zoran B. Redzic (1), Department of Pharmacology, University of Cambridge, Cambridge, CB2 1PD, United Kingdom Hilary J. Rogers (225), School of Biosciences, CardiV University, CardiV, United Kingdom CF10 3TL Michael A. Rudnicki (113), Ottawa Health Research Institute, Ottawa, Ontario, K1H 8L6 Canada Joanna Szmydynger-Chodobska (1), Department of Clinical Neurosciences, Brown University School of Medicine, Providence, Rhode Island 02903 Miles F. Wilkinson (131), Department of Immunology, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030 Ching-Hang Wong (263), Population Council, New York, New York 10021 Hong Wu (199), Program in Developmental Biology, The Hospital for Sick Children, Toronto, Ontario, Canada M5G 1X8 Hai-Tao Yuan (297), Renal, Molecular, and Vascular Medicine Division, Departments of Medicine, Obstetrics and Gynecology, Beth Israel Deaconess Medical Center and Harvard Medical School, Boston, Massachusetts Tao P. Zhong (53), Departments of Medicine and Cell and Developmental Biology, Vanderbilt University School of Medicine, Nashville, Tennessee 37232
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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{ *Department of Pharmacology, University of Cambridge, Cambridge, CB2 1PD United Kingdom { Institute of Gerontology, King’s College London, London, SE1 9NH, United Kingdom { Department of Clinical Neurosciences, Brown University School of Medicine Providence, Rhode Island 02903
I. Introduction II. Fluid Compartments of the Brain A. The Sources of Brain Interstitial Fluid (ISF) and CSF, Bulk Flow of ISF, and the Relationship Between CSF and ISF B. Volume Transmission C. Protein Composition of the CSF III. The CP‐CSF System and the Development of the CNS A. Development of the CP‐CSF System B. The Role of Peptides and Other Biologically Active Substances Either Synthesized in the CP or Transported Across the BCSFB in Brain Development IV. The CP‐CSF System in Adulthood A. Transport Systems in the CP B. The Role of the CP‐CSF System in CNS Injury C. Possible Sources of Stem Cells in the CNS and Their Relation to the CP‐CSF System V. Senescence of the CP‐CSF System A. Aging Parallels with Hydrocephalus? B. CP Senescence: An AD Connection? C. CSF Turnover and Clearance VI. Perspectives Acknowledgments References
The function of the cerebrospinal fluid (CSF) and the tissue that secretes it, the choroid plexus (CP), has traditionally been thought of as both providing physical protection to the brain through buoyancy and facilitating the removal of brain metabolites through the bulk drainage of CSF. More recent studies suggest, however, that the CP‐CSF system plays a much more active role in the development, homeostasis, and repair of the central nervous system (CNS). The highly specialized choroidal tissue synthesizes trophic and angiogenic factors, chemorepellents, and carrier proteins, and is strategically positioned within the ventricular cavities to supply the CNS Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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0070-2153/05 $35.00 DOI: 10.1016/S0070-2153(05)71001-2
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with these biologically active substances. Through polarized transport systems and receptor‐mediated transcytosis across the choroidal epithelium, the CP, a part of the blood‐CSF barrier (BCSFB), controls the entry of nutrients, such as amino acids and nucleosides, and peptide hormones, such as leptin and prolactin, from the periphery into the brain. The CP also plays an important role in the clearance of toxins and drugs. During CNS development, CP‐derived growth factors, such as members of the transforming growth factor‐ superfamily and retinoic acid, play an important role in controlling the patterning of neuronal diVerentiation in various brain regions. In the adult CNS, the CP appears to be critically involved in neuronal repair processes and the restoration of the brain microenvironment after traumatic and ischemic brain injury. Furthermore, recent studies suggest that the CP acts as a nursery for neuronal and astrocytic progenitor cells. The advancement of our knowledge of the neuroprotective capabilities of the CP may therefore facilitate the development of novel therapies for ischemic stroke and traumatic brain injury. In the later stages of life, the CP‐CSF axis shows a decline in all aspects of its function, including CSF secretion and protein synthesis, which may in themselves increase the risk for development of late‐life diseases, such as normal pressure hydrocephalus and Alzheimer’s disease. The understanding of the mechanisms that underlie the dysfunction of the CP‐CSF system in the elderly may help discover the treatments needed to reverse the negative eVects of aging that lead to global CNS failure. ß 2005, Elsevier Inc.
I. Introduction The first account of ‘‘brain water’’ can be ascribed to the ancient Egyptians some 2700 years ago (Breasted, 1930). During the Renaissance period, Andreas Versalius came up with a remarkably precise description of the cerebral ventricles and the choroid plexus (CP) in humans. He also calculated that the volume of ‘‘water‐like fluid’’ that flows through ‘‘cavitae’’ and ‘‘around the brain’’ accounts for approximately one‐sixth of the total brain volume (see Clarke and Dewhurst, 1972). Studies of the human brain, using magnetic resonance imaging (MRI), revealed that the cerebrospinal fluid (CSF), or what Versalius called ‘‘water‐like fluid,’’ encompasses 18% of the total brain volume (Luders et al., 2002). Given the lack of modern technologies, it is quite surprising how accurate Versalius was in his estimates of the CSF space. For many decades the primary function of the CSF was thought to be the physical protection of the brain. It was not until the last 30–40 years that the growing body of evidence suggested a more active role of the CP‐CSF system not only in the mature brain, but also during the development of
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the central nervous system (CNS). CSF is continuously produced by the four CPs of the third, fourth, and two lateral ventricles, and flows along the ventricular system and within the subarachnoid space (SAS), both distributing CSF‐borne substances within the brain and clearing brain metabolites. In addition to its CSF secretory function, the choroidal epithelium synthesizes a large number of bioactive peptides (Chodobski and Szmydynger‐ Chodobska, 2001). Because the receptors for many of these peptides are expressed in choroidal tissue, it is possible that the peptides produced by the CP not only act on brain parenchymal cells, but also regulate the function of the CP itself. The CSF levels of various peptides and proteins produced by the CP change in several pathophysiological situations and in a number of CNS disorders, including brain injury, suggesting that the CP plays an important role in response to brain injury and, possibly, in the subsequent repair processes. The role of the CP in the transport and clearance of both endogenous molecules and xenobiotics, as well as in drug metabolism, has also been well documented (Miller et al., 2005). Therefore, CSF can no longer be considered to simply act as a ‘‘sink’’ for brain metabolites (Oldendorf and Davson, 1967). Rather, the CP‐CSF system should be viewed as an active player in maintaining CNS homeostasis. In this review, the authors will discuss the role of the CP‐CSF system in the development of the CNS and analyze its functional importance in an adult and aging brain. The understanding of the normal physiology of the CP‐CSF system and its malfunction or failure, as well as the appreciation of changes occurring in this system during normal aging, may open new avenues for designing eVective treatment strategies for CNS disorders.
II. Fluid Compartments of the Brain A. The Sources of Brain Interstitial Fluid (ISF) and CSF, Bulk Flow of ISF, and the Relationship Between CSF and ISF There are two major compartments of extracellular fluid (ECF) in the brain: the ISF and the CSF (Fig. 1). It has been postulated that ISF is secreted by the endothelial cells of the brain microvessels into the perivascular space, from where it flows through the low‐resistance pathways along the neuronal tracts and large‐diameter blood vessels. The secretion rate of ISF in the rat brain has been estimated at 0.2 l/min (Cserr et al., 1981), with the total ISF volume of 15–18% of the brain weight. In comparison to the flow of ISF, the CSF formation rate is quite rapid, amounting to 3.4 l/min in the rat (Chodobski et al., 1998b). The majority of CSF is produced by the choroidal epithelium; however, some 10–30% of the total CSF flow is thought to be associated with the bulk flow of ISF. Even though the bulk flow of ISF was
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Figure 1 Schematic diagram of the choroid plexus (CP)‐cerebrospinal fluid (CSF) system. CPs are located in all four cerebral ventricles. The CPs are composed of tightly packed villous folds consisting of a single layer of cuboidal epithelial cells overlying a central core of highly vascularized stroma. The choroidal epithelium is continuous with ependymal lining, but it is morphologically and functionally diVerent from the ependymal cells. The choroidal epithelial cells are joined by tight intercellular junctions. These epithelial tight junctions together with the arachnoid membrane form the blood‐CSF barrier. The CPs are the major source of CSF; however, 10–30% of the total CSF production is represented by the bulk flow of interstitial fluid (ISF). Under normal conditions, there is a net movement of ISF from the brain parenchyma into the CSF. The ventricular CSF is separated from the surrounding brain tissue by the ependyma, whereas the CSF outside of the ventricles is separated from brain parenchyma by pial‐ glial lining. Both the ependyma and the pial‐glial lining oVer little hindrance to the convective flow of fluid and diVusional movement of CSF‐borne substances into the brain parenchyma. CSF flows from the lateral ventricles into the third ventricle, and then continues its movement along the cerebral aqueduct and the fourth ventricle, eventually emptying into the subarachnoid space (SAS). CSF is reabsorbed from the SAS into the blood through the arachnoid villi/granulations protruding into the venous sinuses. Some CSF also drains along cranial nerves and spinal roots out to the lymphatics. Reprinted with permission from A. Chodobski and J. Szmydynger‐ Chodobska, 2001.
discovered over 20 years ago (Cserr, 1984; Rosenberg et al., 1978, 1980), the controversy still exists as to what extent the flow of this fluid contributes to the total CSF production. Promoted by the pressure gradient built up across the ventricular system, CSF flows down the neuraxis eventually emptying into the SAS. Under normal conditions, there is a net movement of ISF from the brain parenchyma into the CSF (Fig. 1), which plays an important role in the ‘‘sink’’ action of CSF (Oldendorf and Davson, 1967) and in volume
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transmission in the brain (see following). However, the direction of ISF flow can be transiently reversed in response to changes in hydrostatic or osmotic pressure, allowing CSF‐borne substances to enter the brain (Pullen et al., 1987; Rosenberg et al., 1978, 1980). B. Volume Transmission It has been postulated that both ISF and CSF play a key role in the so‐called volume transmission (Agnati et al., 1995). The concept of volume transmission has been proposed to describe the chemical communication in the CNS involving both the short‐distance (diVusional) and the long‐range (convective—thanks to the continual secretion and flow of CSF and ISF) movement of signaling molecules within the ECF space of the brain. Thus, volume transmission complements the classical mode of intercellular communication involving synaptic and gap junction‐mediated signaling. The ependymal lining of the cerebral ventricles and the pial‐glial lining at the outer surface of the brain (Fig. 1) are permeable to the high‐molecular‐ weight markers (Brightman and Reese, 1969), allowing for a free diVusional exchange between the CSF and the ISF. The penetration of brain parenchyma by CSF‐borne molecules (e.g., peptides produced by the choroidal epithelium and released into the CSF) is, however, limited to the neuropil located immediately under the ependyma or the pial‐glial lining (Ghersi‐Egea et al., 1996; Proescholdt et al., 2000). Nevertheless, a considerable body of evidence has accumulated demonstrating that the biologically active substances administered into the CSF can exert significant physiological and behavioral eVects that frequently require the activation of large and/or diverse populations of parenchymal cells. Despite intense research into this area, the underlying mechanisms of this ‘‘integrative’’ CSF function remain incompletely understood. It is possible that the biological eVects of some CSF‐borne peptides involve the receptor‐mediated retrograde transport in neurons whose axonal processes are located near the ependymal or the pial‐glial lining (Ferguson and Johnson, 1991; Ferguson et al., 1991; Mufson et al., 1999). Access to the deeper layers of brain parenchyma by CSF‐borne substances may also be facilitated by the movement of these molecules along the perivascular Virchow‐Robin spaces that are in contact with CSF (Agnati et al., 2005). C. Protein Composition of the CSF The amount of protein in the CSF is low (normally <0.5%) when compared to plasma, but the protein composition of this fluid is complex. These proteins may originate from several sources: exclusively from plasma, like
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albumin; primarily from plasma, but are also with a significant proportion synthesized intrathecally, like soluble intercellular cell adhesion molecule 1 (sICAM1); primarily from the CP, like transthyretin (TTR); or primarily from brain parenchyma, like Tau protein (for review, see Reiber, 2001). The main protein fraction in normal CSF originates from plasma and is represented by albumin (Reiber and Peter, 2001). Approximately 20% of CSF proteins are derived predominantly from the brain, but they are hardly ever brain specific. For example, TTR in CSF predominantly originates from the CP, but this protein is also synthesized peripherally in the liver; in another example, the monomer of neuron‐specific enolase (NSE) is not only a neuronal protein, but is additionally synthesized in erythrocytes and thrombocytes. The basic feature of predominantly brain‐derived proteins is their higher concentration in the CSF compared to plasma, resulting in their net flux out of CSF, whereas for peripherally produced proteins there is a net flux into the CSF. Some CSF proteins that are both brain derived and produced by peripheral organs, if present at high plasma levels, may contribute a nonnegligible fraction to their CSF concentrations. The diVerences between brain‐derived and plasma‐derived proteins are best characterized by the CSF/plasma concentration ratio and the so‐called intrathecal fraction (IF). CSF/plasma concentration ratios for brain‐derived proteins are relatively high (e.g., 1:1 for NSE to 34:1 for ‐trace protein) compared to the CSF/plasma concentration ratios for plasma‐derived proteins (e.g., 1:205 for albumin to 1:3400 for IgM). The calculated IF is very high for brain‐derived proteins (e.g., 99% for Tau protein, NSE, S‐100B, cystatin C, or ‐trace protein), but <0.1% for the proteins that, under normal conditions, are exclusively plasma derived. Brain‐derived proteins with a nonnegligible plasma‐derived fraction in the CSF, such as TTR, sICAM1, and a soluble form of angiotensin converting enzyme (ACE), have intermediate CSF/plasma concentration ratios (1:18 to 1:190), with IFs ranging between 90% and 30%, respectively.
III. The CP‐CSF System and the Development of the CNS A. Development of the CP‐CSF System The formation of cerebral ventricles, the meninges, and the CPs takes place early during embryogenesis. The CP diVerentiates from the ependymal cells lining the ventricular walls and, in fact, is frequently considered to be a specialized cuboidal epithelium of ependymal lineage (Ek et al., 2005). The fourth ventricle CP develops first, followed by the CPs of both lateral ventricles and the third ventricle CP (Dziegielewska et al., 2001). The sequence of these events is quite uniform across mammalian species;
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however, the timing of the appearance of individual CPs varies among the species and is most likely related to the length of gestation. Interestingly, the neural tube is filled with fluid before the CPs are formed, raising the question as to whether any CP‐like cells capable of fluid secretion are present prior to the morphogenesis of the CP. Regardless of its source, the fluid filling the neural tube is rich in factors that are necessary for normal neurogenesis. For more information on embryogenesis and morphogenesis of the CP, the reader may refer to other reviews (Dziegielewska et al., 2001; Ek et al., 2005). The CSF formation rate appears to increase gradually during ontogenesis (Evans et al., 1974; Holloway and Cassin, 1972; Johanson and Woodbury, 1974). In sheep, for example, a progressive increase in CSF production takes place both before and after birth (Evans et al., 1974). In rats, the maximum rate of brain growth is observed after birth (Dobbing and Sands, 1979). In these animals, the gradual increase in CSF formation during their postnatal development (Johanson and Woodbury, 1974) correlates well with postnatal morphological changes occurring in the choroidal epithelium (Keep and Jones, 1990; Keep et al., 1986) and with the maturation of the choroidal capability to transport Kþ and Cl– (Parmelee and Johanson, 1989; Preston et al., 1993). Blood flow to the CP, a limiting factor in the CSF production (Cserr, 1971), has also been found to increase gradually in rats after their birth (Szmydynger‐Chodobska et al., 1994). These changes in blood flow to the CP likely reflect a progressive adjustment of the choroidal vasculature to steadily increasing secretory capabilities of the maturing choroidal epithelium. In addition to its CSF secretory function, the choroidal epithelial cells form the physical and functional barrier between the blood and the CSF known as the blood‐CSF barrier (BCSFB). The tight junctions between adjacent epithelial cells appear to be quite well developed in immature CP (Ek et al., 2005), suggesting that, in a growing brain, the properties of this epithelial barrier are largely similar to those typical of adult BCSFB. The apical surface area of the choroidal epithelium appears to be only two times smaller than that of the blood‐brain barrier (BBB), suggesting that the BCSFB plays a much more important role in maintaining brain homeostasis than was previously thought (Keep and Jones, 1990). CSF is not only important for the physical protection of a growing brain, but it also appears that the maintenance of suYcient CSF pressure within the ventricular system is essential for normal development of the CNS. Indeed, in an elegant study on chicken embryos, it was demonstrated that the slow drainage of the CSF from the ventricular system causes significant abnormalities in the neuronal organization of a developing brain (Desmond and Jacobson, 1977). Spina bifida, one of the most common malformations of human CNS resulting from the failure of fusion of the caudal neural tube,
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has also been demonstrated to have serious consequences for the normal development of the cerebral cortex (Miyan et al., 2001). In addition to maintaining optimal hydrostatic pressure, circulating CSF exerts ‘‘nourishing’’ eVects on the developing brain by supplying critical growth factors and other biologically active substances (see later discussion).
B. The Role of Peptides and Other Biologically Active Substances Either Synthesized in the CP or Transported Across the BCSFB in Brain Development Both immature and adult CPs synthesize a large number of neuropeptides, growth factors, and cytokines. In an embryonic brain, the CPs almost completely fill the ventricular cavities whose size is disproportionately large compared to the thin layer of neuroepithelium (Netsky and Shuangshoti, 1975). Therefore, the diVusional distances for CSF‐borne bioactive substances (produced by the choroidal epithelium and released into the CSF and/or transported from the blood into the CSF across the BCSFB) to their putative targets in the developing neural tissue are much shorter than those found in an adult brain. This raises the intriguing possibility that the embryonic CP plays an active role in the development of the CNS. 1. Transthyretin TTR, also referred to as prealbumin, is a carrier protein for thyroxine (T4), the main hormone synthesized by the thyroid gland. The biologically active principle of T4 is triiodothyronine (T3). The latter hormone is primarily derived from the local deiodination of T4, which is mediated in the brain by type II deiodinase (van Doorn et al., 1986). The mRNA levels and the activity of this enzyme are tightly regulated in the CNS (Burmeister et al., 1997) so that the T3 concentration in the brain is maintained at a relatively stable level. An interest in TTR among developmental biologists has been prompted by observations that the thyroid hormones are indispensable for normal growth of the CNS (Anderson, 2001; Bernal and Nunez, 1995; Oppenheimer and Schwartz, 1997). The liver and the CP have been identified as two major sources of TTR. It has been noted, however, that transcriptional regulation of the TTR gene in the CP diVers from that in the liver (Yan et al., 1990). The choroidal epithelium, in which synthesis of TTR begins at an early stage of brain development (Cavallaro et al., 1993), was initially considered as the only source of this carrier protein in the CNS. However, more recent studies have shown that, in addition to the CP, TTR can be produced in the hippocampus, most likely by neurons, in response to various experimental manipulations (Long et al., 2003; Stein and Johnson,
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2002; Stein et al., 2004). Interestingly, some peripherally produced TTR finds its way into the CNS (Terazaki et al., 2001), but the mechanisms by which blood‐borne TTR enters the brain and the physiological significance of this phenomenon are presently unclear. Using the isolated perfused CP and the primary choroidal epithelial cell cultures, researchers (Schreiber et al., 1990; Southwell et al., 1993) demonstrated that TTR is secreted into the CSF. Based on these findings, a model for T4 transport into the brain across the choroidal BCSFB was proposed. They theorized that newly synthesized TTR binds T4, either within the choroidal epithelium or in the CSF, immediately after its release into this fluid, and TTR‐T4 complexes reach the brain parenchyma via the CSF pathways. An enzymatic conversion of T4 to T3 then occurs locally within neuropil. This hypothesis is, in part, supported by observations that extracellular markers injected into the lateral ventricle of the rat are rapidly distributed within the CSF space (Ghersi‐Egea et al., 1996; Proescholdt et al., 2000). However, in another study (Dratman et al., 1991) in which the radiolabeled T4 was administered directly into the ventricular CSF, a markedly restricted distribution of the tracer within the brain parenchyma was observed. The latter finding indicates that transport across the BCSFB is not the major mechanism by which T4 is delivered into the CNS. This conclusion is also consistent with observations made in TTR knockout mice. These animals appear to be phenotypically normal, viable, and fertile (Episkopou et al., 1993). Research (Palha et al., 2000) shows that even though concentrations of T4 and T3 in the CP of TTR‐deficient mice are significantly lower compared to wild‐type controls, the levels of these hormones in the brains of TTR‐null mice are normal. These results suggest that TTR is neither critical for T4 entry into the brain, nor is it necessary to maintain optimal T3 levels in the CNS. Although further studies will be needed to clarify the physiological significance of CP‐derived TTR in the delivery and central homeostasis of thyroid hormones, it is important to note that TTR produced by the choroidal epithelium may have other important functions, such as the regulation of metabolism of ‐amyloid in the CNS (see following). Using TTR knockout mice, it has also been demonstrated that TTR, most likely of CP origin, exerts significant behavioral eVects (Sousa et al., 2004). 2. Insulin‐Like Growth Factor 2 Insulin‐like growth factor 2 (IGF2) is highly expressed in the CP and leptomeninges, even at early stages of mammalian development, and it continues to be synthesized by these tissues in adult brain (Bondy et al., 1992; Hynes et al., 1988; Logan et al., 1994). In contrast, the message for IGF2 has not been detected in either cells of neuroepithelial origin at any
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stage of CNS development or in normal mature brain. Interestingly, in both humans and rodents, there is a biallelic expression of the IGF2 gene in the CP and leptomeninges, whereas in many other tissues, this gene is expressed only from the paternal allele (DeChiara et al., 1991; Ohlsson et al., 1994; Overall et al., 1997). The reason for the parental imprinting of the IGF2 gene is unclear, but it is possible that its biallelic expression in the CP and leptomeninges is necessary for normal development of the CNS. Studies in rats have demonstrated that prior to CP morphogenesis (E10), the message for IGF2 is abundant in the mesenchymal component of the CP primordium, whereas no IGF2 mRNA could be detected in the primordial CP epithelium (Cavallaro et al., 1993). The mesenchymal levels of IGF2 expression decrease during morphogenesis of the CP and IGF2 mRNA is absent in the stroma of adult CP. In contrast to the choroidal stroma, in diVerentiating CP epithelium, IGF2 mRNA gradually increases as embryogenesis progresses. These observations suggest that during early CNS development, mesenchyma‐derived IGF2 acts to promote the diVerentiation of choroidal epithelial cells, whereas epithelium‐derived IGF2 may be involved in the development of other parts of the brain and may also play a role in the normal functioning of adult CNS and/or repair after injury (see later discussion). This hypothesis is consistent with an early expression of receptors for this growth factor in both the CP and other brain areas (Bondy et al., 1992; Kar et al., 1993). There are two types of insulin‐like growth factor receptors (LeRoith et al., 1993; Sara and Hall, 1990). The type I receptor (IGF1R) has the highest aYnity for insulin‐like growth factor 1 (IGF1), but it also recognizes IGF2 and binds insulin at higher concentrations. The type II receptor (IGF2R) has a higher aYnity for IGF2 compared to IGF1 and does not recognize insulin. The IGF1R mediates the mitogenic and neurotrophic eVects of IGFs, and the binding of these ligands to IGF1R results in autophosphorylation of tyrosine residues in the intracellular part of this receptor (LeRoith et al., 1993; Sara and Hall, 1990). Interestingly, IGF1R is not only present in the developing CP, but is also highly expressed in the choroidal epithelium of adult animals (Nilsson et al., 1992), suggesting the autocrine/juxtacrine actions of IGF2 on mature CP. Because adult choroidal epithelial cells have a very slow turnover rate (McDonald and Green, 1988), it is unlikely that, under normal conditions, IGF2 has a mitogenic eVect on choroidal epithelium. However, this growth factor may play a critical role in promoting a rapid recovery of choroidal tissue following the ischemic insult (Johanson et al., 2000). The IGF2R is structurally unrelated to the IGF1R and does not possess tyrosine kinase activity. The functional importance of IGF2R is not completely understood, but it is likely that this receptor plays a role in controlling the levels of IGFs in extracellular fluids by binding and subsequently degrading these growth factors (Haig and Graham, 1991).
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Data obtained in primary cultures of choroidal epithelial cells (Holm et al., 1994; Nilsson et al., 1996) support the idea that IGF2 is secreted from the choroidal epithelium into CSF. In addition to IGF2, the CP produces, and most likely secretes, a number of insulin‐like growth factor binding proteins (IGFBPs), with IGFBP2 and IGFBP4 being expressed at the highest levels (Holm et al., 1994; Stenvers et al., 1994; Walter et al., 1997). These proteins not only act as carriers for IGFs, but also play an important role in modulating the biological activity of these growth factors (Clemmons et al., 1993; Sara and Hall, 1990). Both IGF2 and IGFBP2 could be detected in myelinated nerve tracks in the brain, that is, in the areas that are remote from the site(s) of their synthesis (Logan et al., 1994). This and other studies (see later) thus provide evidence suggesting that CP‐derived IGF2, together with its binding proteins, can act distally on their target cells in various areas of the CNS after being delivered via the CSF pathways. 3. Transforming Growth Factor‐b Superfamily The members of the superfamily of transforming growth factor‐ (TGF‐) have been recognized as important regulators of various cell functions, including proliferation, diVerentiation, and survival. There are three isoforms of TGF‐: TGF‐1, TGF‐2, and TGF‐3. Only TGF‐3 is expressed in embryonic CP (Pelton et al., 1991), whereas all isoforms of TGF‐ are expressed in the epithelial cells of an adult CP (Knuckey et al., 1996). The biological importance of CP‐derived TGF‐ during brain development is not completely understood, but it is likely that this growth factor plays a role in controlling the neuronal organization of developing CNS. Indeed, researchers (Chesnutt et al., 2004) using small interfering RNA to silence the expression of SMAD4, a critical element in TGF‐ signaling, found in the chick embryo that the members of the TGF‐ superfamily are essential for normal pattern formation and the specification of neural progenitor populations in the dorsal neural tube. TGF‐ has also been shown to play an important role in both the induction and survival of dopaminergic neurons in the midbrain (Farkas et al., 2003). Further studies are clearly needed to more precisely define the role of CP‐derived TGF‐ in the developing CNS. Bone morphogenetic proteins (BMPs) are another subfamily of proteins belonging to the TGF‐ superfamily. Several members of this subfamily are present in the embryonic mouse brain, with BMP4, BMP5, BMP6, and BMP7 being expressed in the CP (Furuta et al., 1997). BMPs play an essential role in the development of CP, because in type I BMP receptor mutant mice, the lateral ventricle CPs are greatly reduced or fail to form (He´ bert et al., 2002). Interestingly, the disruption of BMP signaling does not appear to aVect the development of the rest of the telencephalon. BMP6 and BMP7 continue to be expressed in the adult CP (Charytoniuk et al., 2000).
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These CP‐derived BMPs may play a role in neuronal repair processes following ischemic brain injury (Charytoniuk et al., 2000). Recently, a new member of the TGF‐ superfamily, growth/diVerentiation factor 15 or macrophage‐inhibiting cytokine 1 (GDF‐15/MIC‐1), has been cloned. High levels of mRNA for this growth factor have been found in the CP of both newborn (P0) and adult rats (Schober et al., 2001). The hypothesis based on the concept of volume transmission has proposed that GDF‐15/MCI‐1, following its release into the CSF, acts on developing neurons and/or glial cells in brain parenchyma. Again, further studies will be necessary to ascertain the function of this CP‐derived protein in both the developing and mature CNS. 4. Fibroblast Growth Factors Although several members of the family of fibroblast growth factors (FGFs) are expressed in the developing brain, only FGF7 (keratinocyte growth factor) and FGF2, also known as basic FGF, are expressed in the embryonic CP (Finch et al., 1995; Raballo et al., 2000). By comparison, four isoforms of FGF receptor, FGFR1–FGFR4, are present in immature choroidal tissue, with FGFR1, FGFR2, and FGFR4 being expressed on the epithelial cells and within the choroidal mesenchyma, and FGFR3 having nuclear localization (Reid and Ferretti, 2003). These observations suggest that CP‐derived FGFs act as the autocrine and/or juxtacrine/paracrine regulators of CP development. At the same time, volume transmission may be involved in the regulation of CP growth by other members of the FGF family expressed in other parts of the developing brain. For instance, FGF8, originally identified as androgen‐induced growth factor, is expressed in the commissural plate of the embryonic rodent brain and has been found to play an essential role in the normal development of the CP (Theil et al., 1999). Although direct evidence has yet to be established, it is possible that CP‐derived FGF2 not only aVects the development of the CP, but also controls the growth of other parts of the CNS. Indeed, Fgf2 knockout mice, though viable and fertile, exhibit significant abnormalities in the cytoarchitecture of the cerebral cortex (Ortega et al., 1998; Raballo et al., 2000). 5. CP‐Derived Chemorepellents DiVusible chemorepellents play a critical role in axon guidance during the development of the CNS. Studies (Hu, 1999; Nguyen‐Ba‐Charvet et al., 2004; Tamada and Murakami, 2004) have demonstrated that the CP has the ability to synthesize and release such chemorepellents, suggesting that this tissue can provide the guidance cues for growing axons. Two members of the Slit protein family of chemorepellents (Wong et al., 2002), SLIT2 and
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SLIT3, and a secreted member of the semaphorin family (Raper, 2000), semaphorin 3F (SEMA3F), have been found to be expressed in the CP. Using explants of choroidal tissue, including those isolated from Slit2–/– mice, as well as employing COS cells expressing SLIT2, several groups have demonstrated that CP‐derived SLIT2 repels the precursors of olfactory interneurons (Hu, 1999; Nguyen‐Ba‐Charvet et al., 2004; Tamada and Murakami, 2004). The repellent activity of the CP toward olfactory bulb axons was attenuated in the presence of the soluble form of Roundabout (Robo), a receptor for the Slit proteins (Tamada and Murakami, 2004). By comparison, CP‐derived SEMA3F has been demonstrated to repel the axons from the epithalamic and hippocampal explants obtained from the embryonic rat brain (Tamada and Murakami, 2004). When the soluble form of neuropilin 2 (NRP2), a receptor for SEMA3F, was included in the assays, the repellent activity of the CP toward the epithalamic and hippocampal axons was reduced. These observations suggest that CP‐derived SLIT2 and SEMA3F may control axonal growth in various regions of the developing brain. In this context, it is important to note that in their axon guidance actions, the chemorepellents produced by the CP may interact with other CP‐derived growth factors. Indeed, NRP2 and neuropilin 1 (NRP1) do not only function as the receptors for semaphorins (Raper, 2000), but they also bind vascular endothelial growth factor (VEGF) that is highly expressed in the choroidal epithelium (Chodobski et al., 2003). A study has shown that the biological eVects of semaphorin 3A (SEMA3A), a chemorepellent binding to NRP1, are antagonized by VEGF (Bagnard et al., 2001). These authors have also demonstrated that the SEMA3A actions require the presence of type I VEGF receptor. These results indicate that axonal guidance involves specific interactions at both the ligand and receptor levels. 6. Retinoic Acid Retinoic acid (RA), an active derivative of retinol (vitamin A), is essential for normal development of the CNS. It plays an important role in anteroposterior and dorsoventral patterning of neuronal diVerentiation, and its major sites of action are the hindbrain and the anterior part of the spinal cord (Maden, 2002). RA also controls the development of interneurons and motor neurons along the dorsoventral axis. Both RA overexposure and retinol deficiency cause major malformations of all hindbrain structures, frequently resulting in hydrocephalus. During the process of neuronal diVerentiation, RA regulates the expression of a large number of genes by binding to two classes of its receptors that operate as ligand‐activated transcription factors (Bastien and Rochette‐Egly, 2004). Two classes of enzymes, the alcohol dehydrogenases (ADHs) and the retinaldehyde dehydrogenases (RALDHs), are involved in RA synthesis. RA synthesis is also facilitated
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by the cellular retinol‐binding proteins (CRBPs) that sequester retinol and present it to specific dehydrogenases (Ottonello et al., 1993). The CP has the ability to produce RA. This tissue expresses ADHs, though conflicting data have been reported with regard to which members of the ADH family are present in the CP (Galter et al., 2003; Martinez et al., 2001). In addition, the CP expresses RALDH2 (also known as the A2 member of the aldehyde dehydrogenase 1 family; ALDH1A2) and CRBP1, with the latter protein appearing to enhance the enzymatic activity of RALDH2 in the choroidal tissue (Ruberte et al., 1993; Yamamoto et al., 1998). The fourth ventricle CP is closely associated with the cerebellum throughout its development and into the mature CNS. Unlike the developing CP, which is highly active in producing RA, the growing cerebellum has a rather limited capability to synthesize RA (Yamamoto et al., 1996). Normal development of the cerebellum is extremely sensitive to an imbalance in the levels of retinoids, as, for example, an excess of RA has been found to have potent teratogenic eVects on this part of the brain (McCaVery et al., 2003). Based on these observations, it has been proposed that the fourth ventricle CP plays a key role in the development of the cerebellum by being an important source of RA for this hindbrain structure (Yamamoto et al., 1996). These authors have found biphasic changes in the choroidal activity of RALDH2 in both mice and rats, with the first peak of enzymatic RALDH2 activity occurring at E18, followed by the second peak at days 6– 8 of postnatal development. These changes in the choroidal RALDH2 activity correlate well with distinct developmental events in the cerebellum. The first peak in RALDH2 activity observed at an embryonic stage of CNS development coincides with the somatic and axonal diVerentiation of Purkinje cells, whereas the postnatal peak is paralleled by the dendritic arborization of Purkinje cells and diVerentiation of granule cells. Interestingly, RA is not only critical for the development of the cerebellum, but it also appears to play an important role in the growth of the CP, given that the insuYcient dietary intake of vitamin A aVects the development of choroidal tissue (see discussion in Ruberte et al., 1993). Although the enzymatic activity of RALDH2 decreases substantially in the choroidal tissue of the mature brain, an adult CP still maintains the ability to produce RA. The physiological significance of this choroidal function remains unclear, however. 7. Leptin In previous sections, we have analyzed the ability of the choroidal epithelium to produce various bioactive peptides and RA, and discussed how these CP‐derived substances may aVect brain development. Here, the possible role of the BCSFB‐mediated transport of leptin (LEP) in the development of the CNS will be discussed. LEP, the product of the obesity (ob) gene, was
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discovered through positional cloning (Zhang et al., 1994). This hormone is mainly synthesized by white adipose tissue and its serum levels depend on the percentage of body fat (Considine et al., 1996; MaVei et al., 1995). LEP plays a critical role in the regulation of energy balance of the body by acting on several groups of hypothalamic neurons (Ahima and Flier, 2000). The LEP receptor (LEPR) has been identified through expression cloning and found to be present at exceptionally high levels in the CP (Tartaglia et al., 1995). Later on, it was determined, however, that the choroidal LEP receptor is a short isoform of LEPR that has a limited signaling capability compared to the long isoform expressed in the hypothalamus (Bjørbæk et al., 1997; Ghilardi et al., 1996). It has therefore been proposed that the choroidal LEPR plays a role in the receptor‐mediated transport of LEP from the blood into the CSF. This idea is supported by studies in which in situ rat brain perfusion and perfused sheep CP models were used to demonstrate the transport of radioiodinated LEP across the BCSFB (Thomas et al., 2001; Zlokovic et al., 2000). Reduction in the capacity of the choroidal LEP transport and/or impairment of the transport of this hormone across the BBB has been proposed to cause LEP resistance, leading to obesity. Further discussion on this subject will be presented in a later section. It has been recognized for some time that obese Lepob/ob mice that do not produce functional LEP have several abnormalities of the CNS, such as reduced brain weight and brain DNA content, as well as altered dendritic orientation and abnormal myelination (Ahima et al., 1999; Bereiter and Jeanreneaud, 1979, 1980; Steppan and Swick, 1999; Sena et al., 1985; van der Kroon and Speijers, 1979). It has also been shown that in these animals, the total brain protein content is lower compared to wild‐type littermate mice (Ahima et al., 1999). Specific analysis of several neuronal and glial proteins has demonstrated that the levels of expression of various synaptic proteins, such as synaptosome‐associated protein of 25 kDa (SNAP‐25), syntaxin 1 (STX1), and synaptobrevin, are reduced in the cerebral cortex, hippocampus, and hypothalamus of Lepob/ob mice. Similar results were obtained in obese diabetic (db) Leprdb/db mice, in which obesity results from an abnormal splicing of LEPR (Lee et al., 1996). These changes in expression of synaptic proteins were associated with either LEP deficiency or impaired LEP signaling and not with obesity itself, as they were not observed in obese, LEP‐resistant agouti (Ay/a) mice. When immature Lepob/ob mice had received daily intraperitoneal injections of recombinant LEP, researchers observed that the brain levels of expression of SNAP‐25 and STX1 were restored. LEP replacement therapy has also been found to improve the locomotor activity of Lepob/ob mice, which did not appear to be secondary to the loss of body weight resulting from LEP administration, but was instead most likely mediated by LEP itself (Ahima et al., 1999). These observations strongly suggest that LEP plays an important role in the
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development of the CNS. The widespread expression of the long form of LEPR observed in various structures of the embryonic rat brain (Udagawa et al., 2000) is consistent with the preceding hypothesis. In a recent study, scientists investigated the role of LEP in the development of neuronal projections from the arcuate nucleus (ARC), one of the major hypothalamic areas involved in the regulation of food intake (Bouret et al., 2004). These authors have shown that in Lepob/ob mice, neuronal projections from the ARC are permanently disrupted and that LEP replacement in adult mutant mice does not reverse this anatomical defect. Interestingly, there is a prominent surge in serum LEP levels in rodents during their first 2 weeks of postnatal development (Ahima et al., 1998; Morash et al., 2001). Accordingly, in neonatal Lepob/ob mice treated with recombinant LEP it was found that, unlike adult mutants, immature Lepob/ob mice respond to exogenous LEP with normal development of ARC projections (Bouret et al., 2004). LEP has also been shown to induce neurite outgrowth from the ARC in organotypic cultures of this hypothalamic nucleus obtained from P6 wild‐type mice. These findings indicate that LEP is a critical factor in the development of hypothalamic neuronal pathways involved in the control of energy balance. It is important to note, however, that LEP may not only exert direct eVects on immature neurons, but may also influence the development of the CNS by regulating the levels of other hormones, such as glucocorticoids (Ahima et al., 1999), that are known to aVect brain development (Matthews, 2000). Although it is likely that circulating LEP is transported into the brain during development, findings suggest that this protein can also be synthesized centrally in areas such as the hypothalamus, cerebral cortex, and cerebellum (Morash et al., 1999, 2001). Further studies will be needed to clarify the physiological importance of this central LEP synthesis for both immature and adult CNS.
IV. The CP‐CSF System in Adulthood In the previous sections, the role of CSF‐borne substances and the involvement of the CSF pathways in the development of the brain was discussed. In this section, the focus will be on transport/clearance properties of the mature BCSFB and the possible role of the CP‐CSF system in CNS injury.
A. Transport Systems in the CP The exchange processes at the choroidal BCSFB are tightly controlled and involve complex regulatory mechanisms. Choroidal epithelial cells are equipped with a number of transporters that are localized to both the apical
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and basolateral membranes. This polarized distribution of transport systems is essential for the bidirectional movement of various substances across the choroidal epithelial barrier. 1. Transport of Glucose and Amino Acids The Naþ‐independent glucose transporter, GLUT1, is expressed exclusively on the basolateral membrane of choroidal epithelial cells (Kumagai et al., 1994). The transport of glucose across the BCSFB was studied by researchers (Deane and Segal, 1985) who used a model of the perfused sheep CP. These authors estimated the concentration of glucose in the newly formed CSF based on the rate of CSF secretion and the net flux of this sugar across the choroidal epithelium. The concentration of glucose in this fluid was found to be 45–60% of that in plasma, suggesting that the low glucose levels observed in bulk CSF are related to the entry process and not to the cerebral metabolism of this sugar. An uptake of amino acids across the apical (CSF‐facing) membrane of choroidal epithelium was initially demonstrated using an in vitro preparation of the CP (Caruthers and Lorenzo, 1974) and an in vivo ventriculo‐cisternal perfusion technique (Davson et al., 1982). The existence of an apical, Naþ‐ dependent uptake of small neutral and charged amino acids was later confirmed by other researchers who employed either primary cultures of choroidal epithelial cells (Villalobos et al., 1997) or conditionally immortalized choroidal epithelial cells (Kitazawa et al., 2001; Terasaki and Hosoya, 2001). Studies employing the perfused sheep CP have shown that the uptake of amino acids across the opposite, basolateral membrane of choroidal epithelium is strictly equilibrative and mediated by the L‐transport system (for large, neutral, and branched amino acids) and by the ASC system (for small, neutral amino acids) (Preston and Segal, 1990). Such distribution of amino acid transporters (equilibrative transporters in the basolateral membrane and concentrative transporters in the apical membrane) allows for the maintenance of a steep amino acid gradient between the CSF and the plasma, which may play a role in the removal of amino acids having neurotransmitter activities, such as glycine, from the CSF. 2. Transport of Nucleosides Another group of transporters that are present in the choroidal epithelium are nucleoside transporters (for review on this topic, see Redzic, 2005). It appears that the distribution of nucleoside transporters in the choroidal epithelium is polarized and that this polarization is essential for the clearance of nucleosides from CSF. Studies (Wu et al., 1992, 1994) demonstrated that, in the rabbit CP, nucleosides are transported across the apical membrane
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against the concentration gradient and in the presence of inwardly directed Naþ gradient. These authors have also provided the functional evidence that in the choroidal tissue, both purine and pyrimidine nucleosides are substrates for a single Naþ‐nucleoside cotransport system, later designated the cib (concentrative, insensitive to NBTI, broad specificity) system. The cib transport system is now known to be represented by the transporter CNT3 (Gray et al., 2004; Ritzel et al., 2001). Studies in which an in situ perfused ovine CP model was used provided functional evidence that the uptake of purine nucleosides across the basolateral membrane of the choroidal epithelium is strictly equilibrative (Redzic et al., 1997). An analysis of adenosine uptake in primary cultures of rat choroidal epithelial cells confirmed that the distribution of nucleoside transporters is polarized, with the concentrative transport occurring exclusively across the apical (CSF‐facing) membrane (Redzic et al., 2005). It is thus possible that the concentrative transporters expressed in the rat choroidal epithelial cells, rCNT2 (detected at both the mRNA and protein level) and rCNT3 (detected at the mRNA level), are confined to the apical membrane of choroidal epithelium. In contrast, the equilibrative transport, which is presumably mediated by the equilibrative nucleoside transporter 1 (rENT1), was only detectable across the basolateral membrane of choroidal cells (Redzic et al., 2005). This pattern of distribution of nucleoside transporters, together with the well‐known rapid metabolism of adenosine within the choroidal epithelium (Pardridge et al., 1994; Redzic et al., 1997), suggests that CP plays a key role in both preventing circulating adenosine from entering into the CSF and in removing this nucleoside from CSF. This aspect of choroidal function may be critical for central signaling, given that adenosine can act as a neuromodulator. 3. Removal of Xenobiotics An important aspect of the BCSFB function is protection of the brain from toxins and xenobiotics. This subject has been discussed in an excellent review (Miller et al., 2005). The choroidal epithelium expresses a number of transport proteins involved in the eZux of CSF‐borne lipophilic compounds, such as etoposide and vinca alkaloids, and organic anions, such as p‐aminohippurate, benzylpenicillin, and cimetidine. Pharmacokinetic and immunohistochemical studies have demonstrated the presence of the multidrug resistance (MDR) gene product MDR1 P‐glycoprotein (Pgp) and the multidrug resistance‐ associated protein 1 (MRP1) in the choroidal epithelium (Rao et al., 1999; Wijnholds et al., 2000). Studies performed on mutant mice that lacked MRP1 have shown that their CSF levels of etoposide following intravenous (IV) administration of this compound are 10‐fold higher than those observed in mice expressing MRP1 (Wijnholds et al., 2000). A considerable blood‐ to‐CSF concentration gradient across the choroidal epithelium has been
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found in humans after peripheral infusion of Tc‐sestamibi, a membrane‐ permeant radiopharmaceutical whose transport is mediated by both Pgp and MRP (Rao et al., 1999). It has also been reported that an eYcient eZux system for organic anions exists in the CP (Nagata et al., 2002). Using Western blotting and immunohistochemistry, these authors determined that the organic ion transporter (OAT) 3, but not OAT1, is expressed in the rat CP and that OAT3 is localized to the apical (CSF‐facing) membrane of the choroid epithelial cells. 4. Transport of Peptides a. Leptin. The eVect of LEP on CNS development has been previously mentioned. In an adult organism, one of the key functions of this hormone is the maintenance of energy balance (Ahima and Flier, 2000). Convincing evidence has been provided that circulating LEP is transported across the BCSFB (Thomas et al., 2001; Zlokovic et al., 2000). However, the short isoforms of LEPR, thought to be involved in the transport of LEP into the brain, are also expressed in brain microvessels (Hileman et al., 2002). Consistent with these observations, LEP has also been found to cross the BBB (Banks et al., 1996, 2000; Zlokovic et al., 2000), and the importance of LEP transport across the BBB versus BCSFB is presently a matter of debate. (The reader will find a more detailed discussion of this subject in Chodobski et al., 2005.) Although the rodent models of obesity, such as Lepob/ob and Leprdb/db mice, are commonly used to study this disease, their genetic equivalents are rarely observed in humans (O’Rahilly et al., 2003). Rather, the frequent finding in obese individuals is elevated LEP concentrations in serum (Considine et al., 1996; MaVei et al., 1995). It has therefore been proposed that obesity in humans is associated with LEP resistance caused by defective LEP transport into the brain. This hypothesis is supported by a number of clinical and animal studies (Banks and Farell, 2003; Banks et al., 1999; Caro et al., 1996; Hileman et al., 2002); however, the mechanisms underlying this defect in LEP signaling are not completely understood. In this context, it is important to note that LEP resistance may also involve defective signal transduction in the hypothalamic LEPR (El‐Haschimi et al., 2000). Further studies are likely to enhance our insight into LEP resistance, thus facilitating the identification of new potential targets for the treatment of obesity. b. Prolactin. Prolactin (PRL), a hormone synthesized and secreted by the anterior pituitary, is commonly known for its involvement in mammary gland development and lactation. Interestingly, the amino acid sequence of PRL shows similarity with two other hormones: growth hormone (GH) and placental lactogen (PL). Because of their structural homology and
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a similarity of many biological features, these three proteins are called the PRL/GH/PL family. Recently, these hormones were linked to a more extended group of proteins referred to as hematopoietic cytokines (GoYn et al., 2002). PRL elicits a variety of biological responses in diVerent target tissues by binding to its cognate receptor PRLR. Several isoforms of PRLR have been identified, including short, intermediate, and long isoforms, all of which belong to the class I cytokine receptor family (Clevenger and Kline, 2001; GoYn and Kelly, 1997). In addition to PRL, the PRLR binds two other ligands, PL and GH (GoYn et al., 1996), which complicates the understanding of the biological eVects produced by PRL. Both radioligand binding analysis and in situ hybridization histochemistry have demonstrated a particularly high concentration of PRLR in the CP (Brooks et al., 1992; Lai et al., 1992). Consistent with these findings, research (Walsh et al., 1978) showed that, following its IV infusion, 125I‐PRL heavily labeled the CP, whereas the cerebral vasculature was free of radioiodinated PRL. Based on these observations, the authors suggested that circulating PRL is transported into the brain across the BCSFB. This conclusion is supported by other studies from the same laboratory (Walsh et al., 1987) showing the presence of a saturable transport of PRL from the blood to the CSF. Further research, however, will be needed to determine the physiological significance of PRL transport across the BCSFB. c. Clearance of CSF‐Borne Oligopeptides. The choroidal epithelium is equipped with the enzymatic and peptide‐transport systems that play important roles in the processing/degradation of CSF‐borne peptides and the clearance of peptide degradation products. Many peptidases are present in the CP, with some of these enzymes (e.g., ACE and neprilysin) being highly expressed on the apical membrane of choroidal epithelial cells (summarized in Smith et al., 2004). Because of the large apical surface area of choroidal epithelium (previously discussed), these enzymes are in contact with a considerable amount of bulk CSF. Consequently, the choroidal peptidases are likely to play a significant role in the processing and degradation of CSF‐borne peptides. The physiological significance, substrate specificity, and transport kinetics of oligopeptide transporters present in the choroidal BCSFB have been reviewed (Smith et al., 2004). Two oligopeptide transporters, each being a member of a separate family of transporters, have been shown to be expressed in the CP. The first transporter, PEPT2 (Berger and Hediger, 1999), is responsible for the symport of dipeptides and tripeptides along an inwardly directed proton gradient. Functional and immunocytochemical experiments performed in primary cultures of rat choroidal epithelial cells have shown that PEPT2 is expressed apically in choroidal cells and mediates
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the accumulation of a model dipeptide, glycylsarcosine, across the apical membrane of choroidal epithelium (Shu et al., 2002). These results suggest that PEPT2 plays a role in the clearance of oligopeptides and endogenous peptidomimetics from CSF. Another oligopeptide transporter identified to be expressed in the CP, though not yet functionally characterized in this tissue, is peptide/histidine transporter PTH1 (Yamashita et al., 1997). Similar to PEPT2, PTH1 transports dipeptides and tripeptides; however, PTH1 is also able to transport the amino acid L‐histidine. Further studies are needed to define the functional importance of this choroidal transporter.
B. The Role of the CP‐CSF System in CNS Injury It has been demonstrated that in various types of brain injury, such as traumatic brain injury, ischemia, and subarachnoid hemorrhage (SAH), the CSF levels of several growth factors are elevated. The source(s) of these factors is still a matter of debate. However, considering the fact that transcripts for many growth factors have been identified in the CP (Chodobski and Szmydynger‐Chodobska, 2001), it is likely that they originate, at least in part, from the CP. Studies by several groups (Borlongan et al., 2004a,b; Ide et al., 2001; Matsumoto et al., 2005) demonstrated that transplantation of choroidal cells to both traumatized spinal cord and ischemic brain have significant neuroregenerative and neuroprotective eVects. Protection of striatal cholinergic neurons by choroidal grafting in a rodent model of Huntington’s disease has also been shown (Borlongan et al., 2004c). Furthermore, in various in vitro assays, both neuronal survival and neurite outgrowth have been found to be promoted by conditioned media from choroidal cultures or by the coculturing of choroidal epithelial cells with neurons (Borlongan et al., 2004a; Chakrabortty et al., 2000; Kimura et al., 2004). However, the nature of biologically active factors produced by the choroidal epithelium in response to injury and the mechanisms underlying the beneficial eVects of CP grafting are not fully understood. In the following paragraphs, the possible roles of selected, CP‐derived growth factors in brain injury will be analyzed. 1. Insulin‐Like Growth Factor 2 The idea that an increase in CSF concentration of various growth factors observed after brain injury is a result of their augmented production by choroidal epithelium is supported by the study of Walter et al. (1999). These authors reported that after a localized brain injury, there was a transient increase in IGF2 concentration in the CSF that peaked at 7 days post‐ injury. These changes in the CSF IGF2 level were paralleled by increased
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concentrations of immunoreactive IGF2 in the aVected neuropil. Because the message for IGF2 was not increased in any parenchymal cells until later after the injury, it is highly likely that the IGF2 protein detected in the injured parenchyma is of CP/leptomeningeal origin. Interestingly, in the chronic phase (7–14 days after injury), the levels of IGF2 in the CSF declined, and this growth factor appeared to be predominantly synthesized by astrocytes located in the injured parenchyma. These observations suggest the existence of two major sources of IGF2 in the injured brain, with distinct, time‐dependent synthetic activities. In the acute post‐injury phase, IGF2 appears to be largely produced by the choroidal epithelium and leptomeninges, exerting endocrine‐like eVects on its target cells in both the injured neuropil and other unaVected areas of the brain. However, during the late post‐injury period, the production and, possibly, biological actions of IGF2 are mainly confined to the injured parenchyma. 2. Transforming Growth Factor‐b As discussed above, TGF‐ plays an important role in promoting neuronal survival. For example, studies in rodents have demonstrated that the intracerebroventricular (ICV) administration of a moderate dose (4 ng) of recombinant TGF‐1 one hour prior to the induction of transient forebrain ischemia has a significant neuroprotective eVect on pyramidal neurons in the CA1 hippocampal region (Henrich‐Noack et al., 1996). Interestingly, dose of TGF‐1 both approximately 10 times lower or higher did not aVect neuronal survival in the CA1 region. Using a similar rat model of transient forebrain ischemia, researchers (Knuckey et al., 1996) showed an increase in the message for all three isoforms of TGF‐ in the CP at 1–2 days after the insult. Increased choroidal expression of TGF‐1 in other models of brain injury, such as hypoxia‐ischemia and localized cerebral injury, has also been observed (Klempt et al., 1992; Logan et al., 1992). Clinical studies of patients with SAH have demonstrated biphasic changes in the concentration of TGF‐1 in the CSF (Flood et al., 2001). The first peak in TGF‐1 levels (1–2 days post‐SAH) was associated with the disruption of the BBB, whereas the second peak (9–10 days post‐SAH) was not. Researchers have attributed the first peak to the release of TGF‐1 from platelets, a rich source of this growth factor, whereas the second peak has been suggested to result from the central production of TGF‐1 in areas, such as the choroidal epithelium (Flood et al., 2001). This conclusion was supported by the observations that in CPs from SAH patients collected at 10–12 days after SAH, TGF‐1 was expressed at much higher levels than in the choroidal tissues obtained from control subjects. Based on these observations, it is tempting to speculate that in response to injury, larger amounts of TGF‐ are secreted from the choroidal epithelium into the CSF, from which
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this growth factor is transported to its parenchymal target cells to exert neuroprotective eVects. Further studies are needed to test this hypothesis. 3. Fibroblast Growth Factor 2 Originally known as a strong mitogenic and angiogenic factor, FGF2 has been gaining attention as a growth factor with neurotrophic properties (Abe and Saito, 2001). FGF2 has also been shown to have the ability to modulate synaptic transmission. In an adult CP, the mRNA and protein for both FGF2 and its receptor, FGFR1, have been identified (Fuxe et al., 1996; Gonzalez et al., 1995). The receptors for FGF2 are expressed apically on the choroidal epithelium (Szmydynger‐Chodobska et al., 2002) and their activation may play a role in the FGF2‐mediated inhibition of CSF formation (Hakvoort and Johanson, 2000; Johanson et al., 1999b). Another study (Mufson et al., 1999) demonstrated a retrograde neuronal transport of FGF2 from the ventricular CSF, which was suggested by these authors to have important functional implications for the treatment of neurological disorders. This idea is supported by observations that FGF2 administered into the cerebral ventricles has a significant neuroprotective eVect in focal cerebral ischemia in rodents (Koketsu et al., 1994; Ma et al., 2001). Intracerebroventricular infusion of the recombinant FGF2 has also been found to stimulate the diVerentiation of progenitor cells in the subventricular zone (SVZ) into neurons in both young adult and aged rodents (Jin et al., 2003; Kuhn et al., 1997). Considering the anatomical location of the SVZ and its proximity to the CSF, one can speculate that CP‐derived FGF2 likely promotes neurogenesis in the SVZ. However, FGF2 may also exert adverse eVects on neurons. For example, high doses of FGF2 infused into the cerebral ventricles have been found to promote neuronal apoptosis in the caudate‐putamen (Chodobski et al., 1998a), possibly through the caspase‐ dependent mechanisms and downregulation of BCL2 expression (Burchill and Westwood, 2002; Wang et al., 1998). Elevated levels of FGF2 in the CSF have been observed in various CNS disorders, including moyamoya syndrome, Chiari malformation, and hydrocephalus (Malek et al., 1997), suggesting a broad range of biological actions of this peptide in the CNS. These putative actions of FGF2 await further investigation.
C. Possible Sources of Stem Cells in the CNS and Their Relation to the CP‐CSF System Historically, the possibility of neurogenesis in adult mammalian brain has been rejected. Only recently has both the turnover of neuronal cells in the adult CNS been shown and a pool of pluripotent stem cells been identified in
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the SVZ, just under the ependymal lining of the lateral ventricles (Lois and Alvarez‐Buylla, 1993; see also review by Galli et al., 2003). It has been demonstrated that these SVZ cells, when maintained under appropriate conditions, may undergo diVerentiation into neurons or astrocytes (Galli et al., 2003; Lois and Alvarez‐Buylla, 1993). Researchers (Doetsch et al., 1997) have described the topographical organization of the SVZ and identified three distinct types of SVZ cells: type A, with an ultrastructure of migrating neuronal precursor cells; type B (B1 and B2), with characteristics of astrocytes; and type C, with ultrastructural characteristics of immature cells. Interestingly, type B cells frequently have direct contact with CSF by protruding between the ependymal cells (Alvarez‐Buylla and Garcia‐Verdugo, 2002). Although our knowledge about the mechanisms regulating the diVerentiation, migration, and integration of new neuronal cells in adult CNS remains incomplete, the close location of pluripotent SVZ cells to the CSF space suggests that CP‐derived growth factors, such as FGF2 (see previous discussion) and heparin‐binding epidermal growth factor‐like growth factor (Mishima et al., 1996), influence the fate of these cells (Jin et al., 2003). In 2001 intriguing work was reported (Ide et al., 2001). It was demonstrated that the fourth ventricle CP excised from the brain of an adult rat and grafted into the dorsal funiculus of rat spinal cord can promote axonal growth in the host. Later that year, the same group (Kitada et al., 2001) showed that the choroidal epithelial cells harvested from adult mice, cultured for 4–6 weeks, and then grafted into the prelesioned spinal cord of the same species have the ability to diVerentiate into astrocytes. Within 1 week following the grafting of choroidal cells, some transplanted cells were found to stain positively for glial fibrillary acidic protein (GFAP), an astrocytic marker. After 2 weeks, these GFAP‐positive cells demonstrated the morphological characteristics of astrocytes and appeared to be fully integrated in the host tissue. Although in their spinal cord injury model, Kitada et al. (2001) were not able to show that the transplanted choroidal cells diVerentiate into neurons, another study (Li et al., 2002) suggests that the choroidal epithelium has the potential to diVerentiate into neurons after focal cerebral ischemia. In this latter study, a small population of bromodeoxyuridine‐positive cells, presumed to represent proliferating cells, was found to costain for the neuronal marker NeuN in the lateral ventricle CP ipsilateral to the injured hemisphere. Further studies will be needed to confirm these preliminary observations and evaluate their physiological significance.
V. Senescence of the CP‐CSF System Descriptions of age‐related changes to the CP‐ventricular system have drawn many parallels with CNS pathologies. While it is clear that certain aspects of CP‐CSF senescence are not in themselves a disease state, the links between
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the aging system and two pathologies, normal pressure hydrocephalus (NPH) and Alzheimer’s disease (AD), have been outlined in the following.
A. Aging Parallels with Hydrocephalus? 1. Ventriculomegaly and CSF Drainage One of the first changes to the CSF axis in later life is an increase in CSF volume, particularly ventricular volume that increases by 25–30% in 50‐ to 80‐year‐old people compared to 20‐ to 40‐year olds (PfeVerbaum et al., 1994), but also SAS volume (Narr et al., 2003). As a proportion of total intracranial space, the CSF occupies only 7–9% of intracranial volume in adolescence, but starts to increase soon after the second decade of life, reaching 20–33% of intracranial volume by age 71–80 (Courchesne et al., 2000). These changes are exaggerated in neuropathological conditions, including schizophrenia (Narr et al., 2003) and dementia, but in healthy aging, elevated intracranial volume is largely assumed to be secondary to brain atrophy and in one study, total brain volume was smaller in 71‐ to 80‐year olds than in a healthy 2‐ to 3‐year‐old child (Courchesne et al., 2000). Grey matter volume changes are prominent and this volume diminishes by around 5% per decade after adolescence; white matter is relatively preserved but still falls by 13% between the fourth and eighth decades. Accompanying the increased CSF volume, there is evidence for increased resistance to CSF drainage in healthy middle and later life (Albeck et al., 1998), probably as a result of a combination of calcification of the arachnoid villi, thickening of the arachnoid membrane (Bellur et al., 1980), and central vascular hypertension (Rubenstein, 1998). It is striking that the gross changes seen in the aging CSF system resemble changes in NPH characterized by increased intracranial CSF volume, but which, in NPH, is thought to be secondary to increased outflow resistance (Rout) rather than brain atrophy (Boon et al., 1998; Borgesen et al., 1982). Prevalence of NPH increases significantly with age and the pathophysiology is largely unknown (Eide et al., 2003), but risk factors include previous cerebral diseases or trauma, such as meningitis and hemorrhage (Silverberg et al., 2003), and cerebrovascular disease (Boon et al., 1999). Among patients with NPH, Rout significantly increases with age (Czosnyka et al., 2001; Eide et al., 2003), as it does in healthy subjects. 2. CSF Secretion Given the ‘‘closed’’ nature of the CSF circulatory system, any increase in resistance to drainage, Rout, would be expected to elevate intracranial pressure (ICP), following the relationship: ICP ¼ Rout CSF secretion
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rate sinus pressure (Davson et al., 1987), but no correlation between ICP and age is seen in healthy subjects or NPH patients (Czosnyka et al., 2001; Eide et al., 2003). Indeed, for NPH, ICP is usually within the normal range, at least during the day (Eide et al., 2003). An explanation may lie in the downregulation of CSF secretion rate in both healthy aging and NPH. Comparing CSF secretion rates in patients with NPH, Parkinson’s disease (PD), AD, and acute hydrocephalus, observers (Silveberg et al., 2003) found that NPH patients had the lowest secretion rates, at just over 0.2 ml/min, comparable to AD patients (mean age 72; Silverberg et al., 2001), whereas secretion rates in PD (mean age 69; Silverberg et al., 2001) and acute hydrocephalic patients were almost double NPH levels (Silverberg et al., 2002). Decreased CSF production rate in NPH has also been seen in studies (Czosnyka et al., 2001) in humans and in animal models of chronic hydrocephalus (Marlin et al., 1978; Sahar et al., 1971). In animal models, ion transport, fundamental to CSF secretion, which could be analyzed and reduced transfer of both Naþ (Marlin et al., 1978) and Cl– (Knuckey et al., 1993) seems to underlie diminished CSF secretion, at least in kaolin‐hydrocephalus models. Measurements of CSF secretion in aging are less consistent in humans; an early study using the invasive Masserman technique suggested that secretion rates halved from 0.4 to 0.2 ml/min with age, comparing two cohorts averaging 29 and 77 years, respectively (May et al., 1990). Other studies have not seen such definitive changes; describing a mild, but nonsignificant change with age in PD patients from 0.47 to 0.40 ml/min (Silverberg et al., 2001). Noninvasive techniques generally provide higher overall estimates for CSF secretion, but there have not been suYcient studies looking at the oldest age groups (e.g., 75þ). However, using MRI, secretion rates of 0.68 ml/min and 0.69 ml/min in a group of young (average 30 years) and older (average 69 years) subjects was measured (Gideon et al., 1994). Animal studies have provided more consistent findings, with age‐ related decrease in CSF secretion in rats and sheep (Preston, 2001; Wilson et al., 1999). In these models, like in NPH, ion transport deficits are seen in the reduced blood to CSF transport of Naþ in the rat in vivo (Smith et al., 1982), reduced Naþ, Kþ‐ATPase activity and mRNA expression in the rat CP (Kvitnitskaia‐Ryzhova and Shkapenko, 1992; Masseguin et al., 2005), and in the sheep, reduced Naþ uptake and eZux (Chen et al., 2005b), and Cl– eZux (Preston, 1999). The aging sheep CP is also less sensitive to inhibitors of CSF secretion, such as ouabain and acetazolamide (Chen et al., 2005b), and to an upregulator of Kþ/Cl– transport, NEM (Chen et al., 2004). In addition, studies have shown reduced expression of carbonic anhydrase II (providing the HCO3– for Cl– exchange) and aquaporin 1, consistent with reduced water flux across the apical CP membrane (Masseguin et al., 2005).
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a. Vasopressin. Potential underlying mechanisms for such changes are not clearly defined, but will reside both in the general age‐related changes to the CNS and vasculature impacting on brain mass, central sinus pressure, and CP perfusion, as well as to CP‐specific decrements. Arginine vasopressin (AVP) is likely to play a part in the CP‐specific changes. With increasing age (from 25 to 75 years old), AVP levels in plasma rise more than six‐fold in humans (Frolkis et al., 1982) and this was confirmed in later studies (Johnson et al., 1994; Lucassen et al., 1997). CSF levels are also seen to rise in the aging rat, although not in water deprivation state (Frolkis et al., 1999), and there is an increase in the number of AVP‐positive fibers penetrating the third ventricle and forming ‘‘axoventricular’’ contacts (Frolkis et al., 1999). AVP has inhibitory eVects both on CP perfusion and CSF secretion (Chodobski et al., 1998b; Faraci et al., 1988) and induces ‘‘dark’’ epithelial CP cells in young tissue and reduced Cl– flux via the V1 receptor (Johanson et al., 1999a). Similar morphological changes are seen in aged mouse CP, where dark cell numbers are increased (Sturrock, 1988), and in rat, Cl– eZux is reduced (Preston, 1999). Interestingly, CP dark cells are also evident in animal models of hydrocephalus (Shuman and Bryan, 1991) and it has been suggested that the presence of dark cells is indicative of CSF ‘‘resorption’’ in a situation in which there is excess ventricular CSF (Weaver et al., 2004). b. Fibroblast Growth Factor 2. Age‐related changes in FGF2 levels in CP or CSF have not been systematically assessed. What little data there is suggests no changes to either serum or CSF levels with healthy aging in humans (Johansson et al., 2003), although CSF elevations are seen in the neurological disorder amyotrophic lateral sclerosis (Johansson et al., 2003) and in brain parenchyma in AD (Stopa et al., 1990). Data for normal brain levels vary depending on cell type and whether protein immunoreactivity or mRNA is assessed (Belluardo et al., 2004; Cintra et al., 1994; Lolova, 1991), but overall there is a trend for an age‐related increase in baseline FGF2 mRNA in aging rat striatum and cerebral cortex (Belluardo et al., 2004). Nicotine treatment produces significant upregulation of FGF2 mRNA in the cerebral cortex, hippocampus, and substantia nigra of both adult and aged rats (12 and 24 months compared to 3 months) via nicotinic receptors and is neuroprotective (Belluardo et al., 2004). Although elevated FGF2 may be beneficial for neuronal survival following trauma and stimulate neurogenesis (Jin et al., 2003), the evidence that it can induce hydrocephalus via increased resistance to CSF drainage (Johanson et al., 1999b) suggests that high brain levels may contribute to dysfunction of CSF dynamics in aging and disease. FGF2 also induces CP epithelial dark cells and reduces CSF secretion rate, possibly via interaction with AVP (Szmydynger‐Chodobska et al., 2002), and so may exacerbate the AVP eVect on CP.
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c. Cytokines. Marked elevation in CSF cytokines interleukin‐1, tumor necrosis factor‐ (TNF‐), and TGF‐ are seen in various neurodegenerative states with underlying inflammation (Flood et al., 2001; Sjogren et al., 2004; Tarkowski et al., 2003a) and a particularly marked elevation in TNF‐ is seen in NPH with accompanying neuronal degeneration and ependymal disruption (Tarkowski et al., 2003b). However, only TGF‐ levels seem to show any correlation with age, increasing in CSF between 50 and 83 years (Sjogren et al., 2004). Production is regulated by other cytokines, including TNF‐, and TGF‐ acts as an anti‐inflammatory cytokine, inhibiting, by negative feedback, production of proinflammatory cytokines, such as TNF‐. The pattern of cytokines is fundamentally diVerent in aging CSF compared to NPH and probably indicates the relative preservation of brain parenchyma in healthy aging.
B. CP Senescence: An AD Connection? 1. Morphological and Biochemical Changes Whether reduced CSF secretion is a cause of the altered volume transmission seen with aging or a consequence of it, there is abundant additional evidence for global dysfunction of the CP. Pronounced morphological changes occur, including multiple intracellular accumulations of lipofuscin (Serot et al., 2000; Wen et al., 1999), psammoma bodies (Jovanovic et al., 2004), amyloid Biondi bodies (Eriksson and Westermark, 1990; Miklossy et al., 1998), and calcification (Modic et al., 1980). The epithelial cells become flattened and the microvilli are shortened (Serot et al., 2003) reducing the apical surface area, and many of these changes are exaggerated in AD. Extracellular fibrosis of the stroma and thickening of the basement membrane (Serot et al., 2000, 2001; Shuangshoti and Netsky, 1970) would additionally act to impede fluid movement across the tissue in either direction. Generalized oxidative damage to CP nuclear DNA is seen (Nakae et al., 2000), with increased mitochondrial dysfunction in aged humans (Cottrell et al., 2001), decline in activity of enzymes of anaerobic and oxidative respiration (Ferrante and Amenta, 1987), and elevation in CSF lactate (Yesavage et al., 1982), all of which point to declines in energy transduction, essential for maintenance of normal CP function. 2. Protein and Peptide Synthesis TTR is a major protein synthesized by the CP and secreted preferentially into CSF. Published data on TTR changes in CSF with age appear conflicting, and elevated (Kleine et al., 1993; Serot et al., 1997), stable
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(Garton et al., 1991; Kunicki et al., 1998; Vatassery et al., 1991), and declining levels have been reported (Zheng et al., 2001). What is consistent is the finding of reduced CSF TTR in patients with AD (Riisoen, 1988; Serot et al., 1997). In animal models, we have described reduced TTR in aged sheep CSF (Chen et al., 2003). In these animals, there is no change in TTR mRNA expression in the CP (Chen et al., 2005a) or changes in plasma TTR levels, but there was a reduction in de novo protein synthesis by the aging CP and a reduction in newly synthesized TTR monomers both in the CP and in the newly secreted CSF in old sheep (Chen et al., 2005a). TTR has several roles within CSF, including being a chaperone protein for T4 (previously described) and retinoic acid (see Zheng et al., 2001), and importantly, prevents ‐amyloid peptide (A) fibril formation and hence neurotoxicity (Schwarzman et al., 2004). The importance of T4 in maintaining mature neuronal metabolism and of amyloid in the pathogenesis of AD has led to the suggestion that lack of suYcient TTR contributes directly to age‐ related cognitive decline (Rubenstein, 1998). Evidence to suggest that the CP‐CSF axis plays a role in A homeostasis with potential for modulating the onset and course of AD includes nicotine‐ and estrogen‐induced TTR increase in brain and CP (Li et al., 2000; Tang et al., 2004) and may help explain the protective eVect of cigarette smoking and estrogen in dementia (Graves et al., 1991; Tang et al., 1996). In a transgenic mouse carrying a mutant form of human amyloid precursor protein, the mechanism of protection appears to lie in upregulation of TTR given that administration of antibody against TTR results in neuronal loss and apoptosis (Stein et al., 2004). The CP also expresses mRNA for other proteins involved in amyloid handling or with a protective role, including IGF2, which is neuroprotective in vitro (Zheng et al., 2000), IGFBP2, which aVects the action of IGF2 (Walter et al., 1999), gelsolin, which inhibits A fibrillogenesis and prevents neurotoxicity (Matsumoto et al., 2003; Qiao et al., 2005; Ray et al., 2000), and the apolipoprotein J receptor, LRP2 (Kounnas et al., 1994), which mediates cellular clearance of A1–40 (Hammad et al., 1997). We know little about the eVect of late life on production of these compounds, although it is known that production of IGF is partially dependent upon the presence of GH (Cohen et al., 1992), which falls in plasma in later life (Arnold et al., 1999), with reduced density of GH binding to CP (Lai et al., 1993). Because IGF2 may also have a role in promoting choroidal epithelial cell growth (Nilsson et al., 1996), any reduction in activity or content in the CP could have significant consequences for tissue repair and cell turnover. More generally, the global decline in de novo protein synthesis by aged sheep CP and the subsequent reduction in new proteins secreted into CSF (Chen et al., 2005a) could compromise all of these maintenance and protective mechanisms.
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C. CSF Turnover and Clearance For the aging CSF system, like NPH, any one of the gross changes described (increased intracranial CSF volume, increased Rout, or decreased CSF secretion) would be suYcient to increase the turnover time of the CSF leading to stagnation of the fluid and impairment of volume transmission. In aging all these factors are likely to be present, and the rate at which CSF is replaced falls from around 4 times each day to 1–2 times a day in humans (Rubenstein, 1998; Silverberg et al., 2003) and from 12 to 3 times a day in rats (Preston, 2001). As a consequence, the clearance from CSF of a range of compounds slows with increasing age. For example, after lumbar injection of radioiodinated human serum albumin (RIHSA), clearance from aged human CNS is slow and most RIHSA can be detected in the brain 1–2 days later, in contrast to younger subjects when most RIHSA had been cleared (Henriksson and Voight, 1976). Compounds smaller than albumin also show reduced clearance. The rate of removal of ICV‐injected radiolabeled A fell by 90% between 3 and 30 months of age, and at the same time, brain accumulation of the labeled A increased from 7% to 49% (Preston, 2001). Similarly, the old senescence‐accelerated mouse, SAMP8, shows slower eZux of radiolabeled A1–42 from brain after ICV injection compared to young SAMP8 or control mice (Banks et al., 2003). Many other proteins, particularly of large molecular size, have increased CSF/plasma ratios with age. For example, albumin in humans and sheep and IgG in humans (Blennow et al., 1993; Chen et al., 2003; Garton et al., 1991; Reiber, 2001). Brain‐derived proteins have a more varied pattern (Reiber, 2001) presumably because their removal depends on brain ISF flow and the eVect of the local BBB as well as CSF drainage. Because studies on both sheep and rats show no significant change in BCSFB permeability for large compounds (Chen et al., 2005b; Preston, 2001), the elevation of CSF proteins is consistent with reduced CSF turnover (Reiber, 2001) rather than an increased BCSFB or BBB permeability. A further consequence of stagnation in CSF turnover is impaired delivery of compounds by CSF to brain. Direct studies are lacking; however, studies of FGF2 administration into lateral ventricles of young and old mice suggest that this may be a factor. FGF2 administration results in increased neurogenesis in the SVZ and hippocampal dentate gyrus in both young and old mice, demonstrating the potential for neuronal replacement even into old age (Jin et al., 2003). However, the eVects in old mice were mostly seen in the ipsilateral SVZ after unilateral ICV injection, while in young mice, similar increases in neurogenesis were observed in both hemispheres after unilateral injection (Jin et al., 2003). Transiently higher levels of growth factor in the ipsilateral ventricle immediately after injection is put forward as an explanation by the authors, and this is consistent with alterations in CSF dynamics
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and stagnation of ventricular pools in the old rats using CSF as a vehicle for CNS distribution. It is notable that the old SVZ is capable of continuing to act as a nursery for neurogenesis after FGF2 administration; that this is limited in vivo may relate to availability of CSF‐delivered growth factors, as well as neuronal function in late life. The consequences of restoring CSF turnover on cognitive function in AD have been investigated (Silverberg et al., 2003). A ventricular shunt was established to increase CSF drainage and this was successful in stabilizing the cognitive deficit so that at 12 months after shunt, no change in Mattis dementia rating scale was seen. In the nonshunted group, cognitive decline followed its expected course (Silverberg et al., 2003). From human and animal studies of healthy later life and disease, it is clear that there is senescence‐related dysfunction in multiple aspects of the CP‐CSF axis. CP function, CSF secretion, and CSF clearance are all negatively aVected, resulting in reduced volume transmission with detrimental eVects both on the removal of compounds from the CSF/CNS and on the capacity to act as a vehicle for distribution of essential compounds throughout the CNS. The consequences for global CNS function need now to be systematically addressed.
VI. Perspectives The experimental and clinical data previously discussed support the importance of the CP‐CSF system in CNS development, homeostasis and repair, and aging. The CP is a highly specialized tissue, strategically positioned within the ventricular cavities to provide the CNS with a variety of biologically active factors that are essential for normal brain function. Shown in Table I, the CP synthesizes a number of neurotrophic and angiogenic factors, chemorepellents, and carrier proteins, as well as cell‐associated and secreted enzymes, and enzyme inhibitors. The CP is also equipped with many transport systems that not only control the entry of nutrients and other essential substances from the periphery into the brain, but that also play important roles in the clearance of toxins and brain metabolites. The diVusible growth factors and chemorepellents secreted by the CP appear to be critically involved in neurogenesis and axonal guidance during the development of the CNS. Therefore, a better understanding of the role that the CP‐CSF system plays in brain development is likely to benefit in new treatments of congenital CNS disorders. Various functions of immature choroidal tissue subside in the adult CNS; however, after brain injury, the CP resumes its ability to promote neuronal survival and to restore the brain microenvironment. A growing body of evidence suggests that the CP can act as a nursery for neuronal and astrocytic progenitor cells. The demonstrated
Table I
Examples of Peptides and Proteins Synthesized in the Choroid Plexus (CP) Peptide/Protein
Growth factors Bone morphogenetic proteins 4–7 (BMP4–7) Fibroblast growth factor 2 (FGF2)
Growth/diVerentiation factor 15 or macrophage‐inhibiting cytokine 1 (GDF‐15/MIC‐1) Insulin‐like growth factor 2 (IGF2)
Heparin‐binding epidermal growth factor‐like growth factor (HB‐EGF) Transforming growth factor‐ (TGF‐)
Putative Functions
References
Autocrine and/or juxtacrine/paracrine regulation of CP development. May be involved in postischemic neuronal repair processes in the hippocampus Autocrine and/or juxtacrine/paracrine regulation of CP development. May play a role in the development of the cerebral cortex. In adult animals, it promotes the diVerentiation of progenitor cells in the subventricular zone (SVZ) into neurons. Inhibits cerebrospinal fluid (CSF) formation. Overproduction of FGF2 may cause fibrosis of the arachnoid villi and ventricular enlargement Survival‐promoting factor for dopaminergic neurons
Charytoniuk et al., 2000; Furuta et al., 1997; He´ bert et al., 2002
Early expression of IGF2 in the choroidal stroma may promote the diVerentiation of choroidal epithelial cells, whereas epithelium‐derived IGF2 may be involved in the development of other parts of the brain. May play a role in repair processes after brain injury Promotes neurogenesis in the SVZ
Bondy et al., 1992; Cavallaro et al., 1993; Logan et al., 1994; Walter et al., 1999
Neuroprotection in ischemic brain injury
Diaz‐Ruiz et al., 1993; Justicia et al., 2001
Gonzalez et al., 1995; Jin et al., 2003; Johanson et al., 1999b; Kuhn et al., 1997; Ortega et al., 1998; Raballo et al., 2000; Reid and Ferretti, 2003
Schober et al., 2001; Strelau et al., 2000
Jin et al., 2003; Mishima et al., 1996
Transforming growth factor‐ (TGF‐; isoforms 1, 2, and 3; TGF‐3 is expressed in embryonic CP, whereas TGF‐1–3 are expressed in adult CP)
Vascular endothelial growth factor (VEGF)
Chemorepellents Semaphorin 3F (SEMA3F) Slit proteins (SLIT2 and SLIT3) Cytokines and chemokines Interleukin‐1 (IL‐1) Tumor necrosis factor‐ (TNF‐) Cytokine‐induced neutrophil chemoattractants 1 and 2 (CINC1 and CINC2) Neuropeptides Adrenomedullin (ADM)
Arginine vasopressin (AVP) Endothelin 1 (ET1)
May be involved in normal pattern formation and the specification of neural progenitor populations in the dorsal neural tube. May play a role in the induction and survival of dopaminergic neurons in the midbrain. Exerts neuroprotective eVect on the CA1 hippocampal region. Postinjury overproduction may result in hydrocephalus Antagonizes the actions of chemorepellents. Promotes neurogenesis in the SVZ and dentate gyrus. Provides neuroprotection in ischemic brain injury
Chesnutt et al., 2004; Farkas et al., 2003; Flood et al., 2001; Henrich‐Noack et al., 1996; Knuckey et al., 1996; Pelton et al., 1991; Tada et al., 1994
During embryonic development, repels the axons of the epithalamic and hippocampal neurons During embryonic development, SLIT2 repels the precursors of olfactory interneurons
Tamada and Murakami, 2004
Host‐defense response to infection Host‐defense response to infection Promote neutrophil migration across the blood‐CSF barrier (BCSFB)
Quan et al., 1998, 1999 Quan et al., 1999; Tarlow et al., 1993 J. Szmydynger‐Chodobska and A. Chodobski (unpublished observations)
Controls CSF formation and/or other CP functions by up‐regulating choroidal 3’,5’‐cyclic monophosphate synthesis. Modulates the permeability of the blood‐brain barrier Inhibits CSF formation
Kis et al., 2001, 2003; Kobayashi et al., 2001; Takahashi et al., 1997
May play a role in the development of cerebral vasospasm after subarachnoid hemorrhage
Bagnard et al., 2001; Chodobski et al., 2003; Scha¨ nzer et al., 2004; Sun et al., 2003
Hu, 1999; Nguyen‐Ba‐Charvet et al., 2004; Tamada and Murakami, 2004
Chodobski et al., 1997, 1998b; Johanson et al., 1999a Takahashi et al., 1998; Zimmermann and Seifert, 1998 (Continued )
Table I
Continued Peptide/Protein
Carrier proteins Apolipoprotein J (apoJ)/clusterin (CLU) Gelsolin (GSN) Transthyretin (TTR)/prealbumin
Enzymes Alcohol dehydrogenases (ADH1 and ADH4 or ADH3—conflicting data) Matrix metalloproteinases 2 and 9 (MMP2 and MMP9) Prostaglandin H synthase and prostaglandin D synthase/‐trace protein (PGHS and PGDS) Retinaldehyde dehydrogenase 2 (RALDH2/ALDH1A2) Enzyme inhibitors Cystatin C (CST3) Tissue inhibitor of metalloproteinase 3 (TIMP3)
Putative Functions
Binds ‐amyloid peptide (A) and prevents its aggregation and polymerization Inhibits the fibrillization of A and promotes the disaggregation of preformed ‐amyloid fibrils May play a role in the transport of thyroxine (T4) across the BCSFB. Inhibits aggregation of A
References
Aronow et al., 1993; Matsubara et al., 1996 Matsumoto et al., 2003; Ray et al., 2000 Cavallaro et al., 1993; Dickson and Schreiber, 1986; Schreiber et al., 1990; Schwarzman et al., 2004; Southwell et al., 1993
Convert retinol to retinal, the precursor of retinoic acid (RA). RA produced by the fourth ventricle CP may play a role in the development of the cerebellum Facilitate leukocyte migration across the BCSFB
Galter et al., 2003; Marinez et al., 2001; Yamamoto et al., 1996
By sequential actions, produce prostaglandin D2 (PGD2) from arachidonic acid. May play a role in controlling CP function because the receptors for PGD2 are expressed in the CP Converts retinal to RA. RA produced by the fourth ventricle CP may play a role in the development of the cerebellum
HoVmann et al., 1996; Thrikawala et al., 1998; Wright et al., 1999
Neuroprotection in focal cerebral ischemia Inhibits CP‐derived MMP2 and MMP9. Blocks the binding of VEGF to its type II receptor in the CP
Olsson et al., 2004; Tu et al., 1990, 1992 Butler et al., 1999; Pagenstecher et al., 1998; Qi et al., 2003
Strazielle et al., 2003
Yamamoto et al., 1996, 1998
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neuroprotective capabilities of choroidal transplants open new and exciting avenues of research that may lead to designing novel therapies for ischemic stroke, neurotrauma, and neurodegenerative diseases. Furthermore, understanding the mechanisms that lead to the senescence‐related dysfunction of the CP‐CSF axis may help find the treatments needed to reverse the negative eVects of aging that lead to global CNS failure. Accordingly, the challenge for the coming decades is to learn how to bring into clinical practice the potential of the CP to control brain homeostasis and assist in the repair processes of the CNS. It is hoped that this analysis of the various aspects of CP function will attract the broader attention of researchers outside the CP field.
Acknowledgments The authors wish to thank Andrew PfeVer and Alan Miller for their help in the preparation of this manuscript. This work was supported by a grants from the NIH NS39921 and NS49479 (to A.C.) and by research funds from the Neurosurgery Foundation and Lifespan, Rhode Island Hospital (to A.C. and J.S.C.).
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Zebrafish Genetics and Formation of Embryonic Vasculature Tao P. Zhong Departments of Medicine and Cell and Developmental Biology Vanderbilt University School of Medicine Nashville, Tennessee 37232
I. Introduction II. The Origin and Specification of Angioblasts III. DiVerentiation of Arterial and Venous Endothelial Cells A. Delta‐Notch Signaling B. The Role of grl/hey2 Gene C. The Role of VEGF D. Plasticity of Arterial and Venous Endothelial Fate IV. Assembly of the Dorsal Aorta and the Cardinal Vein V. Patterning of the Intersegmental Vessels VI. Perspectives Acknowledgments References
The embryonic vasculature develops in a conserved manner in all vertebrates. Endothelial progenitor cells diVerentiate from mesodermal cells, then migrate and assemble into the dorsal aorta and the cardinal vein. This primitive circulatory loop undergoes sprouting and branching via a two‐step navigation mechanism to form the trunk vascular network. Various studies using several model systems have uncovered a number of signaling mechanisms that regulate these complex processes. A genetic approach in zebrafish has led to identification of mutations and molecules that are responsible for specification of endothelial progenitor cells, diVerentiation of arterial and venous cells, and patterning of the dorsal aorta and intersegmental vessels. These studies highlight the unique utilities and benefits of the zebrafish system for studying development of embryonic blood vessels. ß 2005, Elsevier Inc.
I. Introduction In the vertebrate embryo, the dorsal aorta and the posterior cardinal vein form in the trunk to comprise the circulatory loop prior to a first heartbeat. The endothelial tubes of the great vessels in the embryonic proper assemble Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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through migration and de novo coalescence of endothelial progenitors (angioblasts), a process termed vasculogenesis (Risau, 1996; Risau and Flamme, 1995). Subsequent angiogenesis occurs, where intersomatic vessels sprout from the dorsal aorta and the cardinal vein to form anastomotic arterial and venous systems, allowing blood to circulate in the developing embryo. In the embryonic yolk sac, angioblasts first assemble and form a complex capillary plexus via vasculogenesis. This primitive plexus undergoes pruning and remodeling to develop into the vitelline artery and vitelline vein. In zebrafish embryos, vascular development occurs in a conserved manner compared to other vertebrates (Isogai et al., 2001). The zebrafish yolk sac does not develop a separate vascular system. Instead, part of the cardinal venous system, the duct of Cuvier, comes together to form the common cardinal vein (Fig. 1). Although studies in many vertebrate systems, including chick, frog, and mouse, have contributed greatly to our understanding of embryonic vascular development, zebrafish, a relatively new model organism, oVers distinct advantages for studying vessel formation in vivo. Zebrafish embryos develop externally and are optically clear, providing noninvasive and high‐resolution observation of virtually the entire vascular system at every stage of embryonic development. Furthermore, early development of zebrafish embryos is dependent upon oxygen diVusion, which allows for observation and analysis of vascular developmental events prior to blood circulation. The greatest strength of the zebrafish system is its genetic accessibility due to its small size, fecundity, brief generation time, and easy handling of mutagenesis. Large‐scale mutagenesis screens can be conducted to identify mutations that perturb formation and patterning of embryonic vasculature and other organs (Chen et al., 1996; Staininer et al., 1996; Weinstein et al., 1996). In addition, mutated genes can be rapidly mapped into a small genetic interval, and can be positionally cloned in a rapid way (Talbot and Hopkins, 2000). Because genetic screens make no presuppositions about the roles of genes in biological processes, it is possible to identify novel genetic pathways and developmental mechanisms. Discovery of gridlock (grl ) mutation and identification of grl/hey2 gene as a novel hairy‐related basic helix‐loop‐helix (bHLH) transcription factor in arterial endothelial development proves the strength of zebrafish as an excellent genetic system studying vascular development (Weinstein et al., 1995; Zhong et al., 2000, 2001). Vascular development includes a series of cellular and molecular events, from endothelial cell diVerentiation to blood vessel assembly and morphogenesis. This review discusses some of those important events that have been analyzed mostly in zebrafish, and also compares some of these events to those in other organisms. This discussion begins with specification of endothelial progenitors, then diVerentiation of arterial and venous cells, and finally leads up to assembly and patterning of axial and intersegmental vessels.
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Figure 1 Anatomy of the zebrafish embryonic vasculature. (A) Diagram of a circulatory loop at the zebrafish embryo at 30 hpf. (B) Fluorescent confocal microangiogram showing the zebrafish embryonic vasculature at 60 hpf. AA, Aortic arch; ACV, anterior cardinal vein; CA, caudal artery; CctA, cerebellar central artery; CCV, common cardinal vein (duct of Cuvier); CV, caudal vein; DA, dorsal aorta; DLAV, dorsal longitudinal anastomotic vessel; DLV, dorsal longitudinal vein; H, heart; MsV, mesencephalic vein; NCA, nasal ciliary artery; PCeV, posterior cerebral vein; PCV, posterior cardinal vein; PHBC, primary hindbrain channel; PHS, primary head sinus; PrA, prosencephalic artery; Se, intersegmental vessel; SIV, subintestinal vein; VA, ventral aorta. Panel B reprinted with permission from Isogai et al., 2001.
II. The Origin and Specification of Angioblasts Angioblasts are defined as endothelial precursors that have certain characteristics of endothelial cells and have not yet formed a lumen (Risau and Flamme, 1995). In zebrafish embryos, angioblasts, detected by vascular endothelial growth factor receptor‐2 (VEGFR‐2/flk1), arise and segregate from the lateral plate mesoderm at the 5‐somite stage (Fig. 2A) (Fouquet et al., 1997; Liao et al., 1997; Thompson et al., 1998). Laser‐activated cell lineage analysis indicates that these angioblasts give rise to the primordia of the dorsal aorta, the cardinal veins, or even the intersegmental vessels (Fig. 2F) (Childs et al., 2002; Zhong et al., 2001). Angioblasts arise from
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Figure 2 Formation of the dorsal aortae and the cardinal veins in zebrafish embryos. (A) Dorsal view of flk1‐expressing angioblasts (arrows) arising in the lateral plate mesoderm as bilateral strips at the 5‐somite stage. (B) Dorsal view of angioblasts appearing to migrate toward the midline at the 10‐somite stage. (C) Migratory separation of arterial angioblasts (white arrows) from venous angioblasts (black arrows) at the 15‐somite stage. (D) Primordia of the presumptive aortic bifurcation, aortae (white arrows), and cardinal veins (black arrows) at the 18‐somite stage. (E) Elaboration of the primordial structure in the aortic bifurcation, the aortae, and the cardinal veins. (F) Schematic drawing showing arterial and venous angioblasts migrate and assemble into the dorsal aorta and the cardinal vein.
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the lateral plate mesoderm, migrate to the midline, and assemble into the primordial of the dorsal aorta and the cardinal vein (Fig. 2A–E). Some of the angioblasts exit from axial vessels to populate the intersegmental vessels at late stages. In avian embryos, angioblasts detected by QH1 antibodies originate from the lateral plate mesoderm (CoYn and Poole, 1988; Pardanaud et al., 1987). Splanchnopleural mesoderm, a ventral compartment of the lateral plate mesoderm in contact with endoderm, populates the ventral part (floor) of the dorsal aorta and endothelial lumens of vessels in visceral organs (Pardanaud et al., 1996). Splanchnopleural mesodermal cells also diVerentiate into hematopoietic stem cells residing in the floor of the aorta. Angioblasts derived from the somitic paraxial mesoderm only contribute to vessels in the neural tube, limb buds, and the dorsal‐lateral part (roof) of the aorta, and never invade the visceral organs. In contrast, the somatopleural mesoderm, a dorsal layer of the lateral plate mesoderm in contact with ectoderm, produces few angioblasts (Pardanaud et al., 1996). Thus, angioblasts in the embryo proper originate primarily from mesodermal cells in association with endoderm. In the yolk sac, the earliest sign of angioblast formation is the appearance of blood islands that emerge from the extra‐ embryonic mesoderm in association with visceral endoderm (Haar and Ackerman, 1971). The external cells of blood islands that flatten diVerentiate into endothelial cells, while the internal round cells form hematopoietic stem cells. Based on simultaneous emergence of angioblasts and hematopoietic cells in blood islands, it was postulated that angioblasts and hematopoietic cells originate from a common precursor, termed the ‘‘hemangioblast’’ (His, 1900; Murray, 1932; Wagner, 1980). Several lines of evidence support the existence of such progenitors with dual potential. Both angioblasts and hematopoietic progenitors express similar sets of transcription factors and surface receptors. Development of both angioblast and hematopoietic lineages is impaired in embryos bearing a mutation or a dominant negative form of one of the relevant genes (for reviews, see Dieterlen‐Lievre, 2000; Orkin and Zon et al., 1997). Numerous anatomical studies have shown that direct interactions between endoderm and mesoderm might be required for angioblast induction (Pardanaud and Dieterlen‐Lievre, 1993; Pardanaud et al., 1989; Wilt, 1965). However, studies in Xenopus combining molecular and embryological techniques with carefully designed experimental strategies demonstrate that endoderm is not required for the initial induction of angioblasts (Vokes and Krieg, 2002). The signals for angioblast specification are likely to be mesodermal, and it is possible that angioblast specification occurs by an inherent patterning mechanism. Indeed, a crucial molecular insight into specification of angioblasts came from studies of VEGFR‐2/flk1 and its ligand VEGF. Targeted inactivation of flk1 in mouse embryos causes embryonic lethality and absence of endothelial and hematopoietic cells,
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illustrating that flk1 is essential in inducing angioblast specification (Shalaby et al., 1995). The zebrafish schwentine mutation encodes a truncated flk1 gene (Habeck et al., 2002). Surprisingly, in schwentine mutant embryos angioblast specification still proceeds, although formation of intersomitic and subintestinal vessels is disrupted. This phenotype is not as severe as defects in flk1‐inactivated mouse. These discrepancies might be explained by the truncated flk1 receptor receiving and retaining some VEGF signaling, so that vasculogenesis can still proceed in schwentine mutant embryos. Alternatively, a second copy of flk1 in the zebrafish genome may compensate for the defective flk1 in schwentine mutant embryos (Len Zon, personal communication). In developing embryos, VEGF is expressed in paraxial mesoderm adjacent to angioblasts (Cleaver et al., 1997; Liang et al., 2001). Inactivation of a single VEGF allele results in impaired vasculogenesis in the yolk sac and defective development of the dorsal aorta (Carmeliet et al., 1996; Ferrara et al., 1996). This severe haploid‐insuYcient phenotype suggests that a critical threshold of VEGF protein is required for the flk1 receptor in early vasculogenesis. However, inactivating VEGF activity in both zebrafish and mouse embryos still leads to the presence of angioblasts and endothelial cells. These results suggest that additional VEGF ligands (i.e., VEGFB and VEGFC ) may interact with the flk1 receptor in vasculogenesis (Conway et al., 2001). Alternatively, unknown mesodermal‐derived factors may promote angioblast specification via the flk1 receptor. Zebrafish cloche mutation is the second mutation that blocks angioblast specification (Stainier et al., 1995). cloche mutant phenotypes are similar to the phenotypes caused by targeted inactivation of flk1 in mouse embryos, which fail to develop angioblastic and hematopoietic lineages (Liao et al., 1998). Transcripts of endothelial and hematopoietic markers, including flk1, scl, gata1, tie1, and tie2, are absent in cloche embryos at very early stages, and faint expression of these markers at later stages is seen in the restricted caudal region. Cell transplantation studies reveal that the cloche mutation acts in a cell‐autonomous manner for generation of endothelial lineages, and in a non‐cell‐autonomous fashion for development of hematopoietic lineage at early stages (Stainier et al., 1995). These findings suggest that cloche might act as a cellular intrinsic factor for inducing angioblast formation. Because of the diYculties of mapping the cloche mutation that locates in a telomeric region, the molecular nature of cloche has not been revealed. Nevertheless, gain‐of‐function studies help to place cloche in a putative molecular pathway. Forced expression of scl, a bHLH transcription factor, in cloche embryos restores expression of endothelial marker flk1 and hematopoietic marker gata1, and increases the number of blood and endothelial cells (Liao et al., 1998). These findings suggest that cloche acts upstream of scl for hemangioblast specification, which in turn controls diVerentiation of angioblastic and hematopoietic lineages.
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Studies in zebrafish suggest that scl acts synergistically with lmo2, a LIM domain transcription factor, in converting non‐axial mesoderm into angioblasts and endothelial cells (Gering et al., 2003). Misexpression of scl mRNA in zebrafish embryos induces ectopic expression of endothelial markers flk, fli, and flt4 in somitic paraxial mesoderm. The restriction of angioblast induction to somitic paraxial mesoderm correlates with scl‐induced lmo2 expression in this region. Furthermore, coinjection of lmo2 and scl mRNA extends endothelial fate into head, cardiac, and pronephric duct mesoderm at expense of myocardial and pronephros fates. Expression of both scl and lmo2 can induce angioblast specification and endothelial diVerentiation in non‐axial mesodermal cells. Although these gain‐of‐function studies in zebrafish indicate an important role of scl and lmo2 in angioblast specification, loss‐of‐function studies in mice do not reveal such function. Targeted inactivation of scl or lmo2 in mice causes deficiency in hematopiesis and angiogenesis, but not in angioblast formation (Porcher et al., 1996; Visvader et al., 1998; Warren et al., 1994; Yamada et al., 2000), suggesting that scl and lmo2 are dispensable for angioblast specification at early stages. Indeed, knockdown of scl using antisense morpholinos in zebrafish results in a loss of primitive and definitive hematopoietic lineages, but does not cause absence of angioblasts and endothelial cells (Dooley et al., 2005). Interestingly, the expression of gata2 and lmo2, early hematopoietic genes, is not aVected in embryos reducing scl activities, suggesting that scl is not required for induction and specification of hematopoietic progenitor cells.
III. Differentiation of Arterial and Venous Endothelial Cells It has long been assumed that identity of arterial endothelial and venous endothelial cells is defined primarily by anatomical sites and physiological conditions, and their diVerences may be imposed after, and depend upon, onset of function. A large body of evidence has been collected suggesting that identity of arterial and venous endothelial cells is determined prior to blood circulation, and the fate of arterial and venous endothelial cells can be determined before blood vessel assembly (Lawson et al., 2001, 2002; Wang et al., 1998; Zhong et al., 2000, 2001; for reviews, see Torres‐ Vazquez et al., 2003). Since the initial finding of ephrinB2, a transmembrane ligand, and EphB4, a cognate receptor for ephrinB2, marking arterial and venous endothelium, respectively, many signaling molecules have been documented to be expressed in either arteries or veins in vertebrate embryos (Table I), and multiple signaling pathways have been discovered regulating diVerentiation and development of arterial and venous endothelium.
60 Table I
Tao P. Zhong Markers of Arteries and Veins
Gene Artery specific EphrinB2 deltaC Dll4 Jagged1 Jagged2 Hey1 grl/Hey2 HeyL Neuropilin‐1 Notch1 notch1b Notch3 Notch4 notch5 tbx20 Connexin37 CD44 Vein specific EphB4 flt4 tie2 dab2
Species
Reference
Chick, zebrafish, mouse Zebrafish Mouse Mouse Mouse Mouse Zebrafish, mouse Mouse Chick, mouse Mouse Zebrafish Mouse Mouse Zebrafish Zebrafish Mouse Mouse
Wang et al., 1998; Lawson et al., 2001 Lawson et al., 2001 Shutter et al., 2000 Villa et al., 2001 Villa et al., 2001 Leimeister et al., 1999; Nakagawa et al., 1999 Zhong et al., 2000; Leimeister et al., 1999 Leimeister et al., 1999; Nakagawa et al., 1999 Moyon et al., 2001; Fischer et al., 2004 Villa et al., 2001 Zhong et al., unpublished Villa et al., 2001 Villa et al., 2001 Lawson et al., 2001 Ahn et al., 2000 Duarte et al., 2004 Fischer et al., 2004
Zebrafish, mouse Zebrafish Chick Zebrafish
Wang et al., 1998; Lawson et al., 2001 Thompson et al., 1998 Movon et al., 2001 Song et al., 2004
A. Delta‐Notch Signaling Delta‐Notch signaling has been indicated in regulating arterial‐venous diVerentiation and vascular development. Evidence of involvement of Notch pathway in arterial endothelial development includes expression of Notch receptors and their ligands in arterial endothelial cells as well as diVerentiation defects in arterial endothelial cells. In mouse, notch1, notch3, and notch4 receptor, jagged1, jagged2 and delta‐like4 (dll4) ligand are all expressed in arteries and not in veins (Del Amo et al., 1992; Krebs et al., 2000; Myat et al., 1996; Reaume et al., 1992; Shutter et al., 2000; Uyttendaele et al., 1996; Villa et al., 2001). Among those, notch 1, notch 4, dll4, jagged1, and jagged2 are found in arterial endothelial cells. notch3 and jagged1 are detected in arterial smooth muscle cells (Villa et al., 2001). In zebrafish, deltaC, notch5, and grl, a component of the Notch pathway, are expressed in arterial endothelial cells (Lawson et al., 2001; Zhong et al., 2000). Zebrafish notch1b is observed to be expressed in arterial endothelial
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cells (TPZ, unpublished observation). In the zebrafish genome, notch1a and notch1b are duplicated copies of the notch1 gene (Westin and Lardelli, 1997). Thus, zebrafish notch1b could be the functional counterpart representing the mammalian notch1 gene in arterial endothelial diVerentiation and development. The role of Notch signaling in arterial‐venous diVerentiation and vascular development has been revealed in zebrafish and mouse. Zebrafish mutant mind bomb (mib) encodes a ubiquitin ligase that is essential for signaling cells to activate Notch receptors in neighboring cells (Itoh et al., 2003). mib mutants have a neurogenic phenotype due to a failure of lateral inhibition. The interaction of mib with Delta promotes its ubiquitination and internalization, which facilitates transendocytosis of the Notch extracellular domain and cleavage of the Notch intracellular domain, resulting in activation of downstream genes in neighboring cells. During vascular development, mibta56b mutants display disorganization of axial vessels and cause cranial hemorrhage. Some mutant embryos also develop reduced dorsal aorta, arterial‐venous shunts, and ectopic intersomitic sprouts (Lawson et al., 2001). Molecular characterization of mib using a series of vascular markers show that expression of some arterial gene markers, including efnb2 and notch5, is absent, whereas expression of other arterial markers, including grl and tbx20, is not altered (Lawson et al., 2001). deltaC expression is complexed, and it is reduced in the anterior domain of dorsal aorta but not in the posterior region of aorta. These findings suggest that endothelial cells in mib mutants have established initial arterial fate with presence of some arterial markers, but are defective in subsequent arterial diVerentiation and maintenance with loss or reduction of other arterial markers. In zebrafish, flt4 is expressed in arterial and venous endothelium prior to 24 hpf, and its expression is gradually restricted to venous endothelium after 24 hpf. In mib mutant embryos, flt4 is ectopically expressed in arterial endothelium. Furthermore, interfering Notch signaling by misexpression of dominant negative Su(H), a transcriptional mediator of Notch signaling, causes absence of arterial markers, and in some cases, leads to loss of region of the dorsal aorta and expands contiguous region of cardinal vein (Lawson et al., 2001; Zhong et al., 2001). These data support the idea that arterial fate is specified and maintained via repressing venous fate. Thus, it is possible that redundancy of mib locus in zebrafish might hamper the role of Notch signaling for arterial endothelial formation at early stage. It will be interesting to test whether inactivating functions of both mib and mib‐related gene can cause defects in formation of arterial endothelial cells. Consistent with roles of Notch pathway in vascular development in zebrafish, jagged1 mutant, notch1 mutant, and notch1/notch4 double mutant mice display severe defects in vascular development, including defects in yolk sac vascular remodeling, reduction of the dorsal aorta, massive hemorrhages,
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and embryonic lethality (Krebs et al., 2000; Xue et al., 1999). Re‐ examination of notch1 mutant mice has revealed absence or strongly reduced expression of arterial markers ephrinB2, neuropilin1, and CD44, suggesting notch1 is required for arterial endothelial diVerentiation and development (Fischer et al., 2004). Studies in several groups suggest that dll4 is required in a dosage‐sensitive manner for normal arterial diVerentiation and development (Duarte et al., 2004; Gale et al., 2004; Krebs et al., 2004). Heterozygous dll4þ/ mutant mice already show arterial defects, growth retardation, and haploinsuYcient lethality. The most notable phenotype is the reduction of the size of arteries, including the dorsal aortae, the vitelline artery, and the umbilical artery, as well as expansion of the cardinal veins. Homozygous dll4/ mutant mice develop more severe and precocious vascular defects than heterozygotes, with more pronounced growth retardation and atresia of the aorta. For example, the dorsal aortae, the vitelline artery, and the internal carotid artery are often absent or reduced to capillary remnants, whereas the cardinal veins are expanded to a large degree, resulting in fusion of greatly expanded veins with drastically reduced aortae. In placentas, the initial arterial network has formed; however, the major central placental arteries appear to undergo degeneration and regression. Further characterization of dll4/ embryos with residual dorsal aortae indicates that expression of arterial markers ephrinB2 and connexin37 is absent, whereas venous marker EphB4 is ectopically expressed in arterial endothelium. These arterial defects are also shown in embryos lacking activity of Su(H) in endothelial lineage (Krebs et al., 2004). Thus, the Notch signal pathway regulates the patterning and development of arteries, through the regulation of arterial endothelial diVerentiation, maintenance, and possible proliferation. It is notable that the striking phenotypes resulting from strain‐ dependent haploinsuYciency of dll4 indicate that arterial endothelial development is dosage dependent for dll4. Such severe vascular defects associated with haploinsuYciency have been only noted for VEGF. This suggests that arterial endothelial development may be exquisitely controlled by levels of critical ligands, such as VEGF and dll4. The involvement of Notch pathway in vascular development poses a mechanistic question as to how the Delta‐Notch cascade regulates arterial endothelial diVerentiation and development. In the central and peripheral nervous systems, Notch signaling regulates cell fate through mechanisms of lateral inhibition among a group of cells with equivalent developmental potential, or directive induction between two groups of cells with distinctive diVerentiative capacities. It is best known for selecting cells to be neuroblasts or sensory organ precursor cells through a process of lateral inhibition in Drosophila (Artavanis‐Tsakonas et al., 1999; Lewis, 1998; Mumm and Kopan, 2000). In a group of cells with equal developmental potentials, cells expressing delta (or serrate) ligands communicate with neighbors via notch
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receptors. The interactions trigger the Notch signaling cascade, which transduces into the nucleus via Su(H) to activate a transcriptional repressor E(spl), which in turn downregulates delta in given cells. Such Delta‐ Notch‐mediated lateral inhibition coordinates the fate of adjacent cells, resulting in diVerent developmental status (Artavanis‐Tsakonas et al., 1999). In vasculogenesis, angioblasts arise from the lateral plate mesoderm, and are fated to be arterial and venous cells prior to vessel formation (Ondline and Krieg, 1998; Zhong et al., 2001). The Notch‐mediated lateral inhibition would require that delta and notch be expressed in arterial and venous angioblasts in a complementary manner prior to vessel assembly. Currently, we do not know expression patterns of delta and notch in angioblasts at early stages due to their extremely weak expression at the lateral plate mesoderm. Thus, it remains unknown whether Notch‐mediated lateral inhibition determines choices of arterial or venous cell fate at early stages. However, we do know, after blood vessel formation, transcripts of notch receptors, delta ligands, and downstream grl/hey2 are all detected in arterial endothelium. Thus, at late stages, it is possible that delta (or other ligands) may cell‐autonomously activate Notch signaling in a positive feedback loop in arterial endothelial cells so that an arterial fate can be maintained and a venous fate can be continuously repressed. Such Notch‐mediated cell‐ autocrine mechanism for arterial diVerentiation at late stages may be distinct from that for the Notch‐mediated arterial specification at early stages. It is absolutely critical to examine dynamic expression of notch receptors and their ligands prior to vessel assembly at the lateral plate mesoderm. This would be particularly important to decipher mechanisms in specifying arterial and venous angioblasts at early stages. B. The Role of grl/hey2 Gene The zebrafish grl gene is related to the recently described hey2/hrt2/ hesr1/ chf1 in mouse, belonging to a novel group of the hairy and E(spl)‐related bHLH family (Chin et al., 2000; Kokubo et al., 1999; Leimeister et al., 1999; Nakahawa et al., 1999; Zhong et al., 2000). The hey gene family consists of three members, hey1, grl/hey2, and heyL, in human, mouse, and zebrafish, and a single Drosophila counterpart (Kokubo et al., 1999; Winkler et al., 2003; Zhong et al., 2000). All members of the hey family contain the bHLH motif, Orange domain, and YRPW protein–protein interaction domain (Leimeister et al., 1999; Zhong et al., 2000). The carboxyl‐terminal motif is critical for interactions of Hairy proteins with Groucho, a transcriptional corepressor (Paroush et al., 1994). Whether Grl interacts with Groucho or other proteins via YRPW domain has not been determined. In mouse, members of the hey gene family show partly overlapped but also
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complementary expression patterns. For example, three members of the hey gene family are all expressed in the presomitic mesoderm, somites, and aorta. But in the heart, hey1 and grl/hey2 are expressed in a complementary fashion, in which expression of hey1 and grl/hey2 is restricted to the atrium and ventricle, respectively (Leimeister et al., 1999; Nakagawa et al., 1999). In contrast to this, in zebrafish, grl/hey2 is the only hey gene that is expressed in the aortae (Fig. 3A) and the heart; hey1 is expressed in the presomitic mesoderm and somites, and heyL is expressed in the ventral neural tube (Winkler et al., 2003; Zhong et al., 2000). The members of hey gene family are expressed in distinctive patterns in zebrafish. Particularly, grl/hey2 is expressed in angioblasts at the lateral plate mesoderm prior to vessel formation, and thereafter is restricted to arterial endothelial cells within the formed aortic bifurcation and the dorsal aortae (Winkler et al., 2003; Zhong et al., 2000). Therefore, grl/hey2 is expressed at the correct time and appropriate region for regulating arterial endothelial fate and subsequent diVerentiation in zebrafish. The grl/hey2 gene is suggested to mediate Notch signaling in regulating arterial diVerentiation and vascular development (Fischer et al., 2004; Zhong et al., 2001). Two conserved Su(H)/rbp‐1‐ binding sites (CGTGGGAA) have been identified in the upstream regulatory region of zebrafish grl and its mouse homologue hey2, suggesting that grl/hey2 acts as a transcriptional target of Su(H) (Maier and Gessler, 2000; Nakagawa et al., 2000; Zhong et al., 2001). Indeed, transcriptional activity of hey2 is regulated by Notch signaling in cultured cells (Iso et al., 2001, 2002; Nakagawa et al., 2000). Expression of grl/hey2 gene is downregulated in zebrafish and mouse embryos lacking notch activity (Fischer et al., 2004; Leimeister et al., 2000a,b; Zhong et al., 2001). However, to directly prove grl/hey2 acts as an immediate target of notch signaling in arterial endothelial development, dissection of the grl upstream regulatory region using transgenesis will be needed. The zebrafish grl hypomorphic mutation causes artery‐specific defects, including disruption of the aortic bifurcation, leading to a blockage of blood flow into trunk and ‘‘coarctation’’‐like phenotype (Fig. 3B, C) (Weinstein et al., 1995). The aortic defective phenotype caused by the grl mutation can be rescued by VEGF and certain chemicals (Peterson et al., 2004). In addition to morphological defects in the aortic bifurcation, grl mutants manifest the defective aorta with a decreased lumen size revealed by EGFP expression under control of an flk1 promoter. Characterization of grl mutant embryos derived from homozygous mutant pairs has revealed that the morphologically defective aortae express early arterial markers, notch1b and grl, but not late arterial markers, including deltaC, notch5, ephrinB2, and tbx20 (FL and TPZ, unpublished observations). These data indicate that endothelial cells in grl mutants, like in mib mutants, have acquired the initial arterial fate but are defective in subsequent arterial diVerentiation, maintenance, and
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Figure 3 The arterial endothelium is disrupted in zebrafish embryos reducing grl/hey2 activity. (A) Longitudinal section of anterior region of the embryo showing that grl is expressed in the aortae, and the aortic bifurcation, where the bilateral aortae merge at the midline to form the dorsal aorta. (B) Confocal microscopic image of the aortic bifurcation and venous branches in TG(flk:EGFP) embryos. (C) Confocal microscopic analysis of grlm145 mutant embryos [TG (grlm145 /grlm145; flk:EGFP/þ)] showing a disrupted aortic bifurcation and normal venous branches. (D) Dorsal view of anterior region of the embryo by flk1 in situ staining showing the aortic bifurcation and the anterior cardinal veins laterally (arrowheads). (E) flk1 staining showing no evident aortic bifurcation and aortae in grl‐antisense‐morpholinos‐injected embryos, but the anterior cardinal veins (arrowheads) are present and relatively expanded. ACV, Anterior cardinal vein; CCV, common cardinal vein; da, dorsal aorta; Gray arrowhead, aortic bifurcation; N, notochord; PCV, posterior cardinal vein; White arrow, venous branch. 24 hpf (A, D, E). 48 hpf (B, C). Bars, 50 m (A, B‐C); 100 m (D, E). Panels D and E reprinted with permission from Zhong et al., 2001.
maturation. However, inactivating grl function using antisense morpholino oligonucleotides (antisense‐morpholinos) in a high dose inhibits formation of the aortic bifurcation and the dorsal aorta (Fig. 3D, E), whereas reducing grl activity using antisense‐morpholinos in a low dose phenocopies the defects in grl mutant embryos (Zhong et al., 2001). Thus, grl is required for arterial endothelial diVerentiation and development in a dosage‐sensitive manner in zebrafish. In mouse, hey genes function redundantly in vascular development because of their overlapped expression in the aorta. Targeted inactivation of hey2 causes fatal cardiomyopathy, ventricular septation defects, and high lethality, but arterial endothelial system is not aVected (Donovan et al., 2002; Gessler et al., 2002; Sakata et al., 2002). However, the combined loss of hey1 and hey2 results in reduction of the dorsal aorta, absence of vasculogenesis in the placental labyrinth, lack of vascular remodeling, and embryonic lethality (Fischer et al., 2004). Furthermore, hey1/
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hey2 double knockout leads to absence of expression of arterial markers epherinB2, CD44, and neuropilin1. Thus, the phenotypes in hey1/hey2 double knockout embryos are similar to that in zebrafish embryos reducing grl activity. It will be interesting to find out whether arterial endothelial specification is aVected in mice embryos by triple knocking out activities of hey1, hey2, and heyL. The lack of functional overlap with other hey genes in the zebrafish aortae has revealed arterial endothelial defects in zebrafish embryos deficient for single grl/hey2 gene. These defects are also similar to phenotypes of mouse embryos in absence of function of notch1 or dll4, suggesting that hey1/hey2 function in the Notch pathway for regulating arterial endothelial diVerentiation and development. It is predicted that zebrafish embryos lacking grl activity may display cardiac defects. It will be interesting to find out whether this will be the case. In humans, expression profiling reveals that grl with a group of signaling genes, including Notch receptors, are selectively expressed in human arterial endothelial cells (HUAECs), whereas expression of grl in human venous endothelial cells (HUVECs) induces expression of a group of artery‐specific genes (Chi et al., 2003). The apparently conserved expression pattern of the grl‐dependent pathway in HUAECs highlights the potential importance of this pathway in HUAEC diVerentiation. C. The Role of VEGF VEGF has been shown to be important for migration, proliferation, maintenance, and survival of endothelial cells (Carmeliet, 2000). Studies suggest that VEGF promotes arterial endothelial diVerentiation (Lawson et al., 2002; Mukouyama et al., 2002; Stalmans et al., 2002). In cultured murine embryonic endothelial cells, addition of VEGF‐120 or VEGF‐164 induces ephrinB2 expression up to 50%, whereas addition of bFGF, BMP2, PDGF, IGF‐1, NGF, NT‐3, BDNF, or Sonic Hedgehog can only promote ephrinB2‐ positive cells to ~10%. Importantly, the two VEGF isoforms induce arterial diVerentiation without promoting endothelial proliferation at the low concentration of 2.5 pM (Mukouyama et al., 2002). In zebrafish, reducing VEGF activity using antisense morpholinos causes loss of expression of arterial markers ephrinB2 and notch5, as well as leads to ectopic expression of venous marker flt4. Conversely, misexpression of VEGF‐121 mRNA or VEGF‐165 mRNA causes ectopic expression of ephrinB2 in the cardinal vein. Taken together, these results suggest that VEGF is suYcient to promote arterial endothelial diVerentiation in vivo and in vitro. However, it remains unclear how VEGF influences diVerentiation of arterial endothelial cells. One model has been proposed to illustrate the diVerential eVect of VEGF on arterial endothelial cells in zebrafish, in which some angioblasts
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that migrate medially first preferentially receive VEGF that is secreted from the somites and hypochord. These angioblasts therefore become specified as arterial angioblasts and subsequently diVerentiate into arterial endothelial cells (Torres‐Vazquez et al., 2003). Alternatively, angioblasts with prespecified arterial fate determined by intrinsic cellular mechanism respond to VEGF induction, which triggers overt arterial diVerentiation. Indeed, proportion of ephrinB2‐expressed embryonic endothelial cells that is induced by VEGF never exceeded more than 50%, suggesting that isolated ephrinB2‐ negative embryonic endothelial cells contain prespecified arterial and venous angioblasts in 50% ratio, respectively. VEGF only promotes prespecified arterial angioblasts to subsequently diVerentiate into arterial endothelial cells. Consistent with this observation, selective expression of single VEGF‐ 188 isoform in mouse retinas displays defects in arteriolar diVerentiation and outgrowth. However, arterial endothelial specification is not impaired in mutant mice (Stalmans et al., 2002). Gain‐of‐function and loss‐of‐function studies using VEGF as an entry point in zebrafish have helped to assemble several genes into a pathway that regulates arterial endothelial development. The hedgehog (Hh) signaling pathway has been shown to be important for arterial diVerentiation and development in zebrafish (Lawson et al., 2002). The zebrafish sonic‐you (syu) and you‐too (yot) encode the orthologs of sonic hedgehog (shh) and gli2, a downstream eVector of hedgehog signaling (Karlstrom et al., 1999; Schauerte et al., 1998). Mutations in syu and yot cause a loss of VEGF expression in somites, which in turn downregulates ephrinB2 expression in the aorta, suggesting that the somitic source of VEGF is important for arterial diVerentiation. Forced expression of VEGF‐121 or VEGF‐165 in yot119 mutant embryos rescues ephrinB2 expression. Furthermore, constitutively activated Notch pathway rescues expression of ephrinB2 and notch5 in embryos with reduced VEGF activity, whereas misexpression of VEGF‐121 cannot restore notch5 expression in mibta52b mutant embryos. These data suggest that VEGF acts downstream of hedgehog signaling but upstream of notch pathway for regulating arterial markers ephrinB2 and notch5 expression during arterial endothelial diVerentiation. It is unknown whether such simple epistatic relationships can be held with respect to other arterial markers. Similar to the phenotype in zebrafish embryos reducing VEGF activity, y10 mutation, encoding a defective phospholipase C gamma‐1 (plcg1), causes artery‐specific defects, including absence of sprouting of primary segmental vessels, a failure to form the aortic bifurcation, and reduction of expression of ephrinB2 and notch5 in the dorsal aorta (Lawson et al., 2003). Plcg1 functions downstream of many receptor tyrosine kinases in regulating numerous intracellular signaling pathways (Rhee, 2001; Wilde and Watson, 2001). That misexpression of VEGF causes a failure to rescue ephrinB2 and notch5 expression in y10 mutants implicates that plcg1 may act downstream
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of VEGF signaling in regulating arterial diVerentiation. However, targeted inactivation of plcg1 function in mice causes defects of erythrogenesis, vasculogenesis, and other tissues (Liao et al., 2002). It is possible that preferential expression of plcg1 in zebrafish blood vessel might cause defects restricted to arterial endothelium but not in other tissues.
D. Plasticity of Arterial and Venous Endothelial Fate Although multiple signaling molecules play important roles in regulating identity of arterial and venous endothelial cells prior to blood flow, studies using quail‐chick transplantation revealed that arterial and venous endothelial cell fate remains plastic before certain embryonic stages (Moyon et al., 2001; Othman‐Hassan et al., 2001). Embryonic dorsal aorta or cardinal vein, transplanted from quail donors before embryonic day 7, can colonize both arteries and veins in chick host embryos. These transplanted arterial and venous endothelial cells express ephrinB2, neuropilin‐1 (NRP‐1), and tie2, arterial markers in birds, when they integrate into the lining of arteries. Those cells that are integrated into venous endothelium fail to express these arterial markers. However, specified arterial and venous endothelial cells gradually lose plasticity and become committed after embryonic stage 7. This probably has to do with the vascular wall (smooth‐muscle cell layer) helping to maintaining endothelial fate. At early embryonic stages, arterial and venous endothelial fates are not fully committed, even though they are prespecified, and local vascular environment can alter vessel‐specific endothelial fate. Furthermore, arterial genes, such as ephrinB2, NRP‐1, and tie2, may not be involved in determining arterial and venous endothelial fates, but they could be regarded as diVerentiated and/or maintenance markers. Indeed, ephrinB2 is not expressed in angioblasts at the lateral plate mesoderm in zebrafish embryos (TPZ, unpublished observations). An interesting question is whether constitutive expression of notch receptors and downstream grl/hey2 transcription factors would lead to commitment of arterial endothelial fate. Conversely, it will be important to examine whether notch and grl are ectopically induced in venous endothelial cells when they convert into arterial endothelial cells after transplantation. Venous arterialization occurs when a vein segment is transposed into an arterial region in therapeutic procedures (Henderson et al., 1986; Kwei et al., 2004). The adaptation of endothelial cells to the novel vascular environment could explain how venous endothelial cells respond to the arterial environment. Indeed, notch and grl/hey2 are ectopically induced to high levels in human femoral veins when applied to arterial flow condition (G. Garcia‐Cardena and M. Gimbrone, personal communication), suggesting that venous arterialization in humans requires molecular program that involves establishment of
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arterial endothelial fate at embryonic stages. It remains to be examined whether such arterial transformation leads to a structural and functional adaptation of vascular wall.
IV. Assembly of the Dorsal Aorta and the Cardinal Vein Arterial and venous angioblasts migrate and reach the midline to assemble into endothelial cords, and subsequently, these cords elaborate into lumens of the dorsal aorta and the cardinal vein. Zebrafish genetics has indicated the importance of various midline structures and signaling pathways in organizing assembly of the dorsal aorta; however, little is known about genetic programs that only contribute to assembly of the cardinal vein. Zebrafish floating head (flh) and no tail (ntl) encode Xnot, a homeobox gene, and Brachyury, respectively (Schulte‐Merker et al., 1994; Talbot et al., 1995). In flh mutant embryos, the notochord, the dorsal aorta, and the hypochord fail to develop. Instead, the axial mesodermal tissue transfates into the paraxial somitic muscle in the region of notochord, hypochord, and aorta (Halpern et al., 1995). However, flh mutants develop flk‐expressing endothelial cells in the region of cardinal vein, and these endothelial cells form venous luminal structure (Fouquet et al., 1997). The hypochord is an endodermally derived structure that transiently lies ventral to the notochord, and immediately dorsal to the aorta of fish and amphibian embryos (Gibson, 1910). flh acts cell‐autonomously with respect to the development of notochord, floor plate, and hypochord. Transplanted wild‐type cells give rise to notochord, floor plate, and hypochord in flh host embryos (Halpern et al., 1993). Interestingly, aorta can be rescued in the region immediately underneath the transplanted wild‐type notochord (and possible hypochord). In contrast, aorta is not evident in the region where notochord cannot be rescued (Fouquet et al., 1997). Like flh, ntl mutants lack a diVerentiated notochord and hypochord, but develop the floor plate and one axial vessel. Based on histological analyses, the axial vessel is presumably the cardinal vein. Taken together, these observations reveal a strong correlation between presence of the notochord (and possible hypochord) and the dorsal aorta. Although it is not clear whether notochord or hypochord directly provides signals for organizing the aorta assembly, or whether it organizes other adjacent tissues, which then provide signals for patterning the aorta, these observations at least provide perspective and insights for assembling and patterning the aorta. VEGF plays a critical role in controlling development of the dorsal aorta (Carmeliet et al., 1996; Ferrara et al., 1996; Lambrechts and Carmeliet, 2004). The possible cellular mechanism of VEGF in organizing the aorta assembly came from studies in Xenopus (Ondine and Krieg, 1998). During
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development of Xenopus embryos, VEGF is expressed in the hypochord, somites, and endoderm. Interestingly, only the diVusible VEGF‐121 is preferentially expressed in the hypochord relative to VEGF‐165 and VEGF‐188. Furthermore, ectopic source of VEGF‐121‐expressing implants is suYcient to mediate angioblast migration. It seems plausible that a gradient of VEGF is established between the hypochord and the lateral plate mesoderm and this gradient directs migration of angioblasts into the midline to form the aorta. Further experiments are required to directly demonstrate that loss of VEGF‐121 in the hypochord impairs arterial angioblast migration, which leads to abolishment of aorta formation in embryos of Xenopus. Although loss of VEGF activity in mice causes a complete absence of dorsal aorta formation, it is not clear whether such defects are due to a potential chemoattractant function of VEGF in the midline, considering no hypochord‐ like structures exist in mammals. However, it is possible that diVerent embryonic structures (e.g., somites) express VEGF and might have taken analogous roles in mediating arterial angioblast migration and aorta assembly. Indeed, in zebrafish, VEGF expression is absent in somitic tissues of flh, ntl, and yotty119 mutant embryos, while its expression is present in the hypochord of yotty119 mutant embryos, and in glomeruli and anterior pronephric ducts in flh and ntl mutant embryos (Lawson et al., 2002; Liang et al., 2001). In addition to VEGF, sonic hedgehog signaling has been implicated in organizing assembly of the dorsal aorta in zebrafish and mouse (Brown et al., 2000; Lawson et al., 2002; Vokes and Krieg, 2004). Mutations in components of shh pathway, including in syu, yot, and smootherned (smo), the receptor for shh, block formation of the dorsal aorta but not the cardinal vein (Brown et al., 2000; Chen et al., 2001; Lawson et al., 2002; Varga et al., 2001; Vokes and Krieg, 2004). As discussed above, zebrafish syut4 and yotty119mutants cause loss of VEGF expression in somites. Overexpression of Shh mRNA in wild‐type embryos causes an upregulation of VEGF, and misexpression of VEGF in yotty119 mutant embryos rescues arterial markers ephrinB2 and notch5. Thus, it appears that shh signaling aVects artery development via VEGF. Studies in avian embryos and cultured endothelial cells suggest that shh signaling acts independently of VEGF to mediate artery assembly and vascular tubulogenesis (Vokes and Krieg, 2004). Implanted beads soaked with Shh protein induced artery assembly and vascular tubulogenesis in quail embryos lacking endoderm, whereas VEGF implantation only promotes endothelial cell proliferation. Furthermore, addition of shh to endothelial cells promotes vascular assembly but does not induce VEGF expression. These observations indicate that shh signaling plays a direct role in promoting artery tubulogenesis. Consistent with these findings, mouse embryos lacking function of smo fail to develop lumens of the dorsal aortae. However, angioblasts detected by flk1 expression are accumulated in these regions, suggesting that defects in vascular tubulogenesis are not caused by a
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deficiency in angioblast formation. Furthermore, shh expression is present in the normal developed endoderm in smo/ embryos. Thus, a failure to form artery tube is a direct eVect of deficient smo activity rather than a secondary consequence of altered endodermal development. It is worth pointing out that targeted inactivation of shh in mice does not reveal defects in aorta formation and assembly (Chiang et al., 1996). The reason for the diVerence between mouse and zebrafish is unknown. It is conceivable that the zebrafish syut4 mutants might act in a dominant‐negative fashion to block activities of other hedgehog genes, including tiggy‐winkle hedgehog (twhh) and echidna hedgehog (ehh). It may also be that there is less functional redundancy with shh in the notochord in fish. Studies on vascular tubulogenesis in zebrafish have uncovered interesting cellular events for reshaping endothelial cords into tubes (Parker et al., 2004). At the 22‐somite stage, arterial and venous angioblasts coalesce into single cords at the midline, and thereafter, arterial and venous angioblasts gradually segregate from each other to occupy distinct domains and align in the form of rudimentary tubes. A secreted factor EGF‐like domain 7 (Egfl7) was identified and thought to regulate the cord‐to‐tube transition. Inactivating Egfl7 in zebrafish causes a failure for arterial angioblasts to separate from venous angioblasts, resulting in lack of lumen formation of the aorta and cardinal vein. As the secreted Egfl7 protein remains in the vicinity of endothelial cells and associates with the extracellular matrix, it is proposed that Egfl7 provides a permissive substrate around angioblasts to facilitate the local movement and segregation of arterial angioblasts from venous angioblasts.
V. Patterning of the Intersegmental Vessels Once the dorsal aorta and cardinal vein form in the midline in zebrafish embryos, intersegmental vessels grow and sprout from the dorsal aorta and the posterior cardinal vein in order to form the trunk vascular system. These networks are believed to form via a two‐step mechanism, which has been documented in zebrafish and might also happen in other vertebrates (Isogai et al., 2003). In the first step, primary sprouts emerge bilaterally from the dorsal aorta, then elongate dorsally, branching cranially and caudally along the dorsolateral roof of the neural tube to form paired dorsal longitudinal anastomotic vessels (DLAVs). In the second step, secondary sprouts grow from the posterior cardinal vein, and some of them connect to the base of primary segments. The primary segments with patent connections to secondary segments become intersegmental veins, while the primary segments that remain connected to the dorsal aorta become intersegmental arteries. These intersegmental vessels develop via migration and growth directly from axial
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Figure 4 Model of the construction of an intersegmental vessel in zebrafish embryos. An intersegmental vessel is composed of three types of endothelial cells: the dorsal connection to the DLAV is a T‐shaped endothelial cell; the ventral connection to the aorta is an inverted ‘‘T’’‐ shaped endothelial cell; and the middle connection is an elongated endothelial cell. Adapted with permission from Childs et al., 2002.
vessels. Each of the intersegmental vessels is composed of three types of endothelial cells (Fig. 4) (Childs et al., 2002; Isogai et al., 2003). The first one has its cell body in the dorsal aorta and branches dorsally, the second one connects the first one and the third one, and the third one has its cell body in the DLAV and branches ventrally. These endothelial cells derive from the lateral plate mesoderm, incorporate into the dorsal aorta, and move up to be part of intersegmental vessels. It is important to realize again that zebrafish oVer the genetic and experimental accessibility as well as the optical clarity of developing embryos. The ability to readily combine these components allows rapid imaging and analysis of vascular ‘‘wiring’’ and patterning of intersegmental vessels in developing transgenic embryos. These studies represent the first comprehensive investigation to our understanding of development of angiogenic networks. In zebrafish, many mutants that have been identified are defective in angiogenic growth and patterning of intersomitic and subintestinal vessels (Childs et al., 2002; Habeck et al., 2002; Lawson et al., 2003; MGH screen, unpublished data), including Schwentine, y10, and Out of bounds (obd). As previously discussed, Schwentine and y10 encode flk1 and phospholipase C gamma‐1 (plcg1), respectively (Habeck et al., 2002; Lawson et al., 2003). plcg1 is thought to transduce intracellular VEGF signaling. In addition to arterial endothelial defects in y10 mutants, both of these mutations, Schwentine and y10, disrupt growth and sprouting of intersegmental and
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subintestinal vessels. Defects in VEGF/Flk1 pathway impair sprouting and growth of intersegmental vessels, which is consistent with the function of VEGF for promoting blood vessel growth. Out of bounds (obd) mutants display mispatterning of intersegmental vessels but not growth defects of those sprouts. In obd mutant embryos, intersegmental vessels sprout precociously at irregular sites, and these misguided sprouts do not elongate along the intersomitic boundaries, instead of forming highly interconnecting branches along the trunk. Positional cloning identifies obd as plexinD1 (plxnD1), one of the receptors for semaphorin ligands. Furthermore, reducing plxnD1 function by microinjection of antisense morpholinos causes the same defects as obd mutants (Torres‐Vazquez et al., 2004). These studies clearly indicate that plxnD1 is required for proper blood vessel pathfinding. In the developing nervous system, the axonal pathway is shaped by repulsive cues provided by semaphorin ligands that are responded to by plexin receptors in migrating neuronal growth cones (Raper et al., 2000). Semaphorins could be repulsive signals in restricting navigation of intersegmental vessels to the intersomitic boundaries. Indeed, sema3a1 and sem3a2 are expressed in developing somites. sema3a1 expression is detected in the entire somites at 15 hpf, but is excluded from the horizontal myoseptum from 18–36 hpf (Yee et al., 1999). Reducing the function of either sem3a1 or sem3a2 using antisense morpholinos causes similar but not nearly as severe phenotypes as the plxnD1 morphants or obd mutants. Such eVects appear to be mediated by direct interactions of the PlxnD1 receptor and the Sem3a1 ligand. Addition of Sema3a to the cultured HUVECs, which express plxnD1, results in loss of acting fibers and decreases endothelial cell migration. Furthermore, overexpression of sem3a2 under myogenin promoter in somitic muscle in zebrafish embryos inhibits growth of intersegmental vessels, and such inhibition is not observed in embryos lacking plxnD1 activity. These works provide a molecular mechanism for patterning of blood vessels in the trunk in which intersegmental vessels expressing plexinD1 extend to the semaphorin‐free intersomitic boundaries but cannot grow into the somitic region where semaphorins are expressed. Consistent with the role of plxnD1 in zebrafish, targeted inactivation of plxnD1 in mice causes mispatterning of intersegmental vessels. In addition to vascular defects, plxnD1/ mutant mice exhibit phenotypes similar to congenital heart diseases in humans, including outflow tract defects, enlarged atria, and persistent truncus arteriosus (Gitler et al., 2004). The observation of cardiac defects in mice lacking plexinD1 suggests that the plxnD1 receptor has acquired new functions during evolution. More importantly, these cardiac defects that have previously been attributed to abnormalities of neural crest cells can now be caused by cell‐autonomous endothelial defects.
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In addition to semorphrins/plexinD1, the netrin receptor unc5B and its ligand netrin‐1a have been demonstrated to play an important role in regulating endothelial tip cell filopodia and branching (Lu et al., 2004). unc5B is selectively expressed in arterial endothelial cells and a subset of venous capillary endothelial cells. Notably, unc5B is expressed in endothelial tip cells of both arterial and venous sprouts. Loss of function of unc5B in mice causes defects in vascular branching and navigation, including excessive vessel branching in the internal carotid artery, intersegmental vessels, and vessels in the nervous system. In zebrafish embryos lacking activity of unc5B or netrin‐1a, intersegmental vessels exhibit ectopic filopodial extensions and form extra vessel branches at variable positions, resulting in developing aberrant anastomosis. This phenotype is similar to embryos reducing activity of plxnD1or sem3a1. In cultured HUAECs, netrin‐1 induces filopodial retraction, which reduces migration of endothelial cells. Such netrin‐1‐mediated repulsive eVects require unc5B for patterning vascular system. Blood vessels are guided through repulsive mechanisms using the same cues and receptors that guide axons (for reviews, see Carmeliet, 2003). However, it is not clear how netrin‐1/unc5B signaling interacts with semorphrins/plexinD1 signaling for navigating and patterning intersegmental vessels.
VI. Perspectives This overview highlights some of the signals and mechanisms involved in embryonic vessel formation. Although much has been determined about pathways and mechanisms in vascular development, many questions remain to be addressed. In particular, it is not clear which signaling mechanisms are required for angioblast specification. To understand the earliest vascular development, it will be important to identify mesodermal cells that have potential to be angioblasts, such as hemangioblast‐like stem cells. Cell‐ lineage analysis in living embryos will help to find these cells, and genetic screens for identifying cloche‐like mutations will aid to reveal signaling pathways for angioblast specification. To delve deeper into the regulation of arterial endothelial development, it will be critical to determine multiple cellular events and mechanisms in this process. For example, does arterial‐ venous endothelial diVerentiation contain multiple steps like those in neural cells? Does Delta‐Notch signaling determine arterial fate at the stage of angioblasts via lateral inhibition? Finding the answers to these questions should be facilitated by studies using loss‐of‐function and gain‐of‐function analysis in diVerent vascular developmental stages. Furthermore, dissecting the Delta‐Notch signal transduction pathway in vascular development will require fine control of gene expression in endothelial progenitor cells and diVerentiated endothelial cells. Thus, isolation of endothelial‐specific
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promoters marking early and late stages of arterial–venous diVerentiation, as well as generation of transgenic embryos will be critical for conducting these functional studies. Progress for blood vessel navigation indicates that members of four axon‐guidance families, including netrins, semaphorins, ephrins, and slits (Dickson et al., 2002), play important roles in mediating vessel branching and patterning. The extent to which their guidance functions are conserved and these signaling pathways are interacted in the vascular system is still incompletely understood. Further experiments will be required for characterizing and comparing gain‐of‐function and loss‐of‐ function phenotypes in diVerent model systems. Clearly, understanding of blood vessel formation will be continually benefited from studies of several model systems. The experimental and genetic tools currently available in zebrafish make this a very eYcient system for studying vascular development. Importantly, the power of forward genetics will set the pace for discovering novel endothelial genes. The capability to reduce gene function by using antisense morpholinos makes it possible to conduct reverse genetics in zebrafish. The synergistic eVects of combining loss‐of‐function and gain‐of‐function analysis in the same organism will enhance the ability of discovering genetic pathways and mechanisms in vascular development. As the basic mechanisms and signaling pathways are well conserved between fish and mammals, the results of zebrafish studies will contribute greatly to understanding normal vessel development and vascular lesions in humans.
Acknowledgments Members of the Zhong laboratory contributed to some of the ideas and work summarized here. I thank Brant Weinstein, Len Zon, Joey Barnnett, Scott Baldwin, and David Bader for helpful advice and critical reading of the manuscript, and oVer my apologies to many authors whose work I have not cited in this review. Work in the laboratory has been supported by grants from the NIH, AHA, March of Dimes Foundation, and Vanderbilt Academic Venture Capital Fund.
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Lawson, N. D., Scheer, N., Pham, V. N., Kim, C. H., Chitnis, A. B., Campos‐Ortega, J. A., and Weinstein, B. M. (2001). Notch signaling is required for arterial‐venous diVerentiation during embryonic vascular development. Development 128, 3675–3683. Lawson, N. D., Vogel, A. M., and Weinstein, B. M. (2002). sonic hedgehog and vascular endothelial growth factor act upstream of the Notch pathway during arterial endothelial diVerentiation. Dev. Cell 3, 127–136. Leimeister, C., Dale, K., Fischer, A., Klamt, B., Hrabe de Angelis, M., Radtke, F., McGrew, M. J., Pourquie, O., and Gessler, M. (2000). Oscillating expression of c‐Hey2 in the presomitic mesoderm suggests that the segmentation clock may use combinatorial signaling through multiple interacting bHLH factors. Dev. Biol. 227, 91–103. Leimeister, C., Externbrink, A., Klamt, B., and Gessler, M. (1999). Hey genes: A novel subfamily of hairy‐ and Enhancer of split related genes specifically expressed during mouse embryogenesis. Mech. Dev. 85, 173–177. Lewis, J. (1998). Notch signaling and the control of cell fate choices in vertebrates. Semin. Cell Dev. Biol. 9, 583–589. Liang, D., Chang, J. R., Chin, A. J., Smith, A., Kelly, C., Weinberg, E. S., and Ge, R. (2001). The role of vascular endothelial growth factor (VEGF) in vasculogenesis, angiogenesis, and hematopoiesis in zebrafish development. Mech. Dev. 108, 29–43. Liao, E. C., Paw, B. H., Oates, A. C., Pratt, S. J., Postlethwait, J. H., and Zon, L. I. (1998). SCL/Tal‐1 transcription factor acts downstream of cloche to specify hematopoietic and vascular progenitors in zebrafish. Genes Dev. 12, 621–626. Liao, H. J., Kume, T., McKay, C., Xu, M. J., Ihle, J. N., and Carpenter, G. (2002). Absence of erythrogenesis and vasculogenesis in Plcg1‐deficient mice. J. Biol. Chem. 277, 9335–9341. Liao, W., Bisgrove, B. W., Sawyer, H., Hug, B., Bell, B., Peters, K., Grunwald, D. J., and Stainier, D. Y. (1997). The zebrafish gene cloche acts upstream of a flk‐1 homologue to regulate endothelial cell diVerentiation. Development 124, 381–389. Lu, X., Le Noble, F., Yuan, L., Jiang, Q., De Lafarge, B., Sugiyama, D., Breant, C., Claes, F., De Smet, F., Thomas, J. L., Autiero, M., Carmeliet, P., Tessier‐Lavigne, M., and Eichmann, A. (2004). The netrin receptor UNC5B mediates guidance events controlling morphogenesis of the vascular system. Nature 432, 179–186. Maier, M. M., and Gessler, M. (2000). Comparative analysis of the human and mouse Hey1 promoter: Hey genes are new Notch target genes. Biochem. Biophys. Res. Commun. 275, 652–660. Moyon, D., Pardanaud, L., Yuan, L., Breant, C., and Eichmann, A. (2001). Plasticity of endothelial cells during arterial‐venous diVerentiation in the avian embryo. Development 128, 3359–3370. Mukouyama, Y. S., Shin, D., Britsch, S., Taniguchi, M., and Anderson, D. J. (2002). Sensory nerves determine the pattern of arterial diVerentiation and blood vessel branching in the skin. Cell 109, 693–705. Mumm, J. S., and Kopan, R. (2000). Notch signaling: From the outside in. Dev. Biol. 228, 151–165. Murray, P. D. F. (1932). The development in vitro of the blood of the early chick embryo. Proc. Roy. Soc. 11, 497–521. Myat, A., Henrique, D., and Ish‐Horowicz, D. J. L. (1996). A chick homologue of Serrate and its relationship with Notch and Delta homologues during central neurogenesis. Dev. Biol. 174, 233. Nakagawa, O., McFadden, D. G., Nakagawa, M., Yanagisawa, H., Hu, T., Srivastava, D., and Olson, E. N. (2000). Members of the HRT family of basic helix‐loop‐helix proteins act as transcriptional repressors downstream of Notch signaling. Proc. Natl. Acad. Sci. USA 97, 13655–13660.
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Shutter, J. R., Scully, S., Fan, W., Richards, W. G., Kitajewski, J., Deblandre, G. A., Kintner, C. R., and Stark, K. L. (2000). Dll4, a novel Notch ligand expressed in arterial endothelium. Genes Dev. 14, 1313–1318. Stalmans, I., Ng, Y. S., Rohan, R., Fruttiger, M., Bouche, A., Yuce, A., Fujisawa, H., Hermans, B., Shani, M., Jansen, S., Hicklin, D., Anderson, D. J., Gardiner, T., Hammes, H. P., Moons, L., Dewerchin, M., Collen, D., Carmeliet, P., and D’Amore, P. A. (2002). Arteriolar and venular patterning in retinas of mice selectively expressing VEGF isoforms. J. Clin. Invest. 109, 327–336. Talbot, W. S., and Hopkins, N. (2000). Zebrafish mutations and functional analysis of the vertebrate genome. Genes Dev. 14, 755–762. Talbot, W. S., Trevarrow, B., Halpern, M. E., Melby, A. E., Farr, G., Postlethwait, J. H., Jowett, T., Kimmel, C. B., and Kimelman, D. (1995). A homeobox gene essential for zebrafish notochord development. Nature 378, 150–157. Thompson, M. A., Ransom, D. G., Pratt, S. J., 3rd, MacLennan, H., Kieran, M. W., Detrich, H. W., 3rd, Vail, B., Huber, T. L., Paw, B., Brownlie, A. J., Oates, A. C., Fritz, A., Gates, M. A., Amores, A., Bahary, N., Talbot, W. S., Her, H., Beier, D. R., Postlethwait, J. H., and Zon, L. I. (1998). The cloche and spadetail genes diVerentially aVect hematopoiesis and vasculogenesis. Dev. Biol. 197, 248–269. Torres‐Vazquez, J., Gitler, A. D., Fraser, S. D., Berk, J. D., Van, N. P., Fishman, M. C., Childs, S., Epstein, J. A., and Weinstein, B. M. (2004). Semaphorin‐plexin signaling guides patterning of the developing vasculature. Dev. Cell 7, 117–123. Torres‐Vazquez, J., Kamei, M., and Weinstein, B. M. (2003). Molecular distinction between arteries and veins. Cell Tissue Res. 314, 43–59. Uyttendaele, H., Marazzi, G., Wu, G., Yan, Q., Sassoon, D., and Kitajewski, J. (1996). Notch4/ int‐3, a mammary proto‐oncogene, is an endothelial cell‐specific mammalian Notch gene. Development 122, 2251–2259. Varga, Z. M., Amores, A., Lewis, K. E., Yan, Y. L., Postlethwait, J. H., Eisen, J. S., and Westerfield, M. (2001). Zebrafish smoothened functions in ventral neural tube specification and axon tract formation. Development 128, 3497–3509. Villa, N., Walker, L., Lindsell, C. E., Gasson, J., Iruela‐Arispe, M. L., and Weinmaster, G. (2001). Vascular expression of Notch pathway receptors and ligands is restricted to arterial vessels. Mech. Dev. 108, 161–164. Visvader, J. E., Fujiwara, Y., and Orkin, S. H. (1998). Unsuspected role for the T‐cell leukemia protein SCL/tal‐1 in vascular development. Genes Dev. 12, 473–479. Vokes, S. A., and Krieg, P. A. (2002). Endoderm is required for vascular endothelial tube formation, but not for angioblast specification. Development 129, 775–785. Vokes, S. A., Yatskievych, T. A., Heimark, R. L., McMahon, J., McMahon, A. P., Antin, P. B., and Krieg, P. A. (2004). Hedgehog signaling is essential for endothelial tube formation during vasculogenesis. Development 131, 4371–4380. Wagner, R. C. (1980). Endothelial cell embryology Dan growth. Adv. Microcirc. 9, 21–35. Wang, H. U., Chen, Z. F., and Anderson, D. J. (1998). Molecular distinction and angiogenic interaction between embryonic arteries and veins revealed by ephrin‐B2 and its receptor Eph‐B4. Cell 93, 741–753. Warren, A. J., Colledge, W. H., Carlton, M. B., Evans, M. J., Smith, A. J., and Rabbitts, T. H. (1994). The oncogenic cysteine‐rich LIM domain protein rbtn2 is essential for erythroid development. Cell 78, 45–57. Weinstein, B. M., Schier, A. F., Abdelilah, S., Malicki, J., Solnica‐Krezel, L., Stemple, D. L., Stainier, D. Y., Zwartkruis, F., Driever, W., and Fishman, M. C. (1996). Hematopoietic mutations in the zebrafish. Development 123, 303–309. Weinstein, B. M., Stemple, D. L., Driever, W., and Fishman, M. C. (1995). Gridlock, a localized heritable vascular patterning defect in the zebrafish. Nat. Med. 1, 1143–1147.
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Westin, J., and Lardelli, M. (1997). Three novel Notch genes in zebrafish: Implications for vertebrate Notch gene evolution and function. Dev. Genes Evol. 207, 51–63. Wilde, J. I., and Watson, S. P. (2001). Regulation of phospholipase C gamma isoforms in hematopoietic cells: Why one, not the other? Cell. Signal. 13, 691–701. Wilt, F. (1965). Erythropoiesis in the chick embryo: The role of endoderm. Science 147, 1588–1590. Winkler, C., Elmasri, H., Klamt, B., VolV, J. N., and Gessler, M. (2003). Characterization of hey bHLH genes in teleost fish. Dev. Genes Evol. 213, 541–553. Xue, Y., Gao, X., Lindsell, C., Norton, C. R., Chang, B., Hicks, C., Gendron‐Maguire, M., Rand, E. B., Weinmaster, G., and Gridley, T. (1999). Embryonic lethality and vascular defects in mice lacking the Notch ligand Jagged1. Hum. Mol. Genet. 8, 723. Yamada, Y., Pannell, R., Forster, A., and Rabbitts, T. H. (2000). The oncogenic LIM‐only transcription factor Lmo2 regulates angiogenesis but not vasculogenesis in mice. Proc. Nat. Acad. Sci. USA 97, 320–324. Yee, C. S., Chandrasekhar, A., Halloran, M. C., Shoji, W., Warren, J. T., and Kuwada, J. Y. (1999). Molecular cloning, expression, and activity of zebrafish semaphorin Z1a. Brain Res. Bull. 48, 581–593. Zhong, T. P., Childs, S., Leu, J. P., and Fishman, M. C. (2001). Gridlock signaling pathway fashions the first embryonic artery. Nature 414, 216–220. Zhong, T. P., Rosenberg, M., Mohideen, M. A., Weinstein, B., and Fishman, M. C. (2000). gridlock, an HLH gene required for assembly of the aorta in zebrafish. Science 287, 1820–1824.
Further Reading CoYn, D. J., and Poole, T. J. (1998). Embryonic vascular development: Immunohistochemical identification of the origin and subsequent morphogenesis of the major vessel primordia in quail embryos. Development 102, 735–748. Leimeister, C., Schumacher, N., Steidl, C., and Gessler, M. (2000). Analysis of HeyL expression in wild‐type and Notch pathway mutant mouse embryos. Mech. Dev. 98, 175–178. Parker, L., and Stainier, D. Y. (1999). Cell‐autonomous and non‐autonomous requirements for the zebrafish gene cloche in hematopoiesis. Development 126, 2643–2651. Risau, W. (1997). Mechanisms of angiogenesis. Nature 386, 671–674. Stainier, D. Y., Fouquet, B., Chen, J. N., Warren, K. S., Weinstein, B. M., Meiler, S. E., Mohideen, M. A., Neuhauss, S. C., Solnica‐Krezel, L., Schier, A. F., Zwartkruis, F., Stemple, D. L., Malicki, J., Driever, W., and Fishman, M. C. (1996). Mutations aVecting the formation and function of the cardiovascular system in the zebrafish embryo. Development 123, 285–292. Song, H. D., Sun, X. J., Deng, M., Zhang, G. W., Zhou, Y., Wu, X. Y., Sheng, Y., Chen, Y., Ruan, Z., Jiang, C. L., Fan, H. Y., Zon, L. I., Kanki, J. P., Liu, T. X., Look, T. A., and Chen, Z. (2004). Hematopoietic gene expression profile in zebrafish kidney marrow. PNAS 101, 16240–16245. Vogel, A. M., and Weinstein, B. M. (2000). Studying vascular development in the zebrafish. Trends Cardiovasc. Med. 10, 352–360.
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Leaf Senescence: Signals, Execution, and Regulation Yongfeng Guo and Susheng Gan Cornell Genomics Initiative and Department of Horticulture, Cornell University, Ithaca, New York 14853‐5904
I. Introduction II. Leaf Senescence‐Regulating Signals A. Age B. Reproductive Growth C. Sugars D. Stresses E. Ethylene F. Jasmonic Acid (JA) G. Salicylic Acid (SA) H. Brassinosteroids (BRs) I. Abscisic Acid (ABA) J. Cytokinins K. Polyamines (PAs) L. Nitric Oxide (NO) III. The Cell Death Execution Process A. Chlorophyll Degradation B. Protein Degradation C. Lipid Degradation D. Nutrient Recycling IV. Gene Expression and Regulation During Leaf Senescence A. Gene Expression During Leaf Senescence B. Perception and Transduction of Senescence Signals C. Transcription Regulation of Gene Expression V. Closing Remarks Acknowledgments References
Leaf senescence is a type of postmitotic senescence. The onset and progression of leaf senescence are controlled by an array of external and internal factors including age, levels of plant hormones/growth regulators, and reproductive growth. Many environmental stresses and biological insults such as extreme temperature, drought, nutrient deficiency, insuYcient light/shadow/darkness, and pathogen infection can induce senescence. Perception of signals often leads to changes in gene expression, and the upregulation of thousands of senescence‐associated genes (SAGs) causes Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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the senescence syndrome: decline in photosynthesis, degradation of macromolecules, mobilization of nutrients, and ultimate cell death. Identification and analysis of SAGs, especially genome‐scale investigations on gene expression during leaf senescence, make it possible to decipher the molecular mechanisms of signal perception, execution, and regulation of the leaf senescence process. Biochemical and metabolic changes during senescence have been elucidated, and potential components in signal transduction such as receptor‐like kinases and MAP kinase cascade have been identified. Studies on some master regulators such as WRKY transcription factors and the senescence‐responsive cis element of the senescence‐specific SAG12 have shed some light on transcriptional regulation of leaf senescence. ß 2005, Elsevier Inc.
I. Introduction Senescence is a nearly universal biological phenomenon (Hughes and Reynolds, 2005). In plants, there exist two types of senescence: mitotic and postmitotic senescence. Mitotic senescence occurs in germ‐like shoot apical meristem, which is similar to replicative senescence or replicative aging, for example, in mammalian cell cultures and yeast. In contrast, postmitotic senescence occurs in somatic cells of organs such as leaves and flowers. This type of senescence is similar to that which occurs in somatic tissues of the animal adult body (Gan, 2003). Leaf senescence, a type of postmitotic senescence, is the final phase of leaf development, from maturation to attrition in the life history of a leaf. It is a unique process that is characterized by diVerential gene expression, active degeneration of cellular structures, and recycling of nutrients (Buchanan‐Wollaston, 1997; Feild et al., 2001; Gan, 2003; Gepstein, 2004; Lim et al., 2003; Quirino et al., 2000; Smart, 1994). The visible sign of leaf senescence is yellowing, resulting from the preferential degradation of the green pigment chlorophyll but not the yellow‐red pigment carotenoids. In some plant species, accompanying the senescence process is the synthesis of anthocyanins and other pigments, which contributes to the various colors in autumn leaves (Feild et al., 2001). During leaf senescence, macromolecules such as proteins, lipids, and nucleic acids are degraded, resulting in the sharp decrease in the photosynthetic activity. Nutrients released from catabolism of these macromolecules are translocated to active growing regions such as new buds, young leaves, developing fruits and seeds, or to be stored in trunks for next growing season. The massive operation of catabolism and nutrient mining leads to the ultimate cell death (Buchanan‐Wollaston, 1997; Feild et al., 2001; Gan, 2003; Gepstein, 2004; Quirino et al., 2000; Smart, 1994). As an evolutionary
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fitness strategy, stress‐induced leaf senescence contributes to plant survival under unfavorable environmental conditions (Munne‐Bosch and Alegre, 2004). Loss of assimilatory capacity as photosynthetic organs undergo senescence, on the other hand, contributes to yield limitation of cereal crops (Egli, 2004). Occurrence of senescence after harvest devaluates vegetable crops and ornamental plants during postharvest storage, transportation, and on shelves. This review summarizes our current understanding of the complex cell‐death process of leaf senescence, including signals or factors that induce or inhibit senescence, the cell‐death execution process and the underlying molecular regulatory mechanisms.
II. Leaf Senescence‐Regulating Signals The onset and progress of leaf senescence are controlled by a complement of external and internal factors. Internal factors influencing leaf senescence include age, levels of plant hormones/growth regulators, and developmental processes such as reproductive growth. Many environmental stresses and biological insults such as extreme temperature, drought, nutrient deficiency, insuYcient light/shadow/darkness, and pathogen infection can induce senescence (Gan, 2005; He et al., 2001; Smart, 1994). A. Age In the absence of external stressors, leaf senescence occurs in an age‐ dependent manner (Hensel et al., 1993; Jiang et al., 1993; Nooden and Penney, 2001). How age initiates leaf senescence is not well understood. It has been speculated that decline of photosynthesis with age is a possible mechanism (Hensel et al., 1993). Several lines of evidence, however, indicated an antagonistic role of a decline in photosynthetic capability on leaf senescence. The Arabidopsis mutant ore4 with a lesion in a plastid ribosomal small subunit protein has a reduced photosynthetic activity and its age‐ dependent leaf senescence is delayed (Woo et al., 2002). Leaves in the transgenic tobacco plants with suppressed expression of the Rubisco small subunit gene have a lower photosynthetic activity and a longer life span than those of wild‐type (Miller et al., 2000). Accumulation of reactive oxygen species (ROS) has also been suggested as a possible mechanism through which age induces senescence (Munne‐Bosch and Alegre, 2002). Degradation of stromal proteins such as Rubisco can be initiated by ROS when chloroplasts are incubated in photo‐oxidative stress conditions (Roulin and Feller, 1998). However, ROS accumulation occurs after protein and lipid degradation is initiated, that is, an increased level of
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ROS is the result of macromolecule degradation processes. A highly senescence‐specific Arabidopsis gene SAG12 appears to be regulated by developmental age but not by other environmental and endogenous factors (Noh and Amasino, 1999a). SAG12 encodes a cysteine proteinase (Gan, 1995). Analysis of the regulatory mechanism of the SAG12 expression may provide insights into the age‐dependent mechanisms of leaf senescence. B. Reproductive Growth Reproductive development may trigger leaf senescence in many plant species, especially in monocarpic plants. Monocarpic plants have single reproductive growth in their life history. Removal of flowers or fruits extends leaf longevity in many monocarpic plant species such as soybean (Craftsbrandner and Egli, 1987), pea (Pic et al., 2002), rice (Khan and Choudhuri, 1992), and sunflower (Sadras et al., 2000). The leaf life span was extended by 50% in pea plants when flowers were removed (Pic et al., 2002). The removal of flowers delayed the onset of leaf senescence and slowed down the senescence progression (Pic et al., 2002). However, not all experiments involving removal of flowers and/or fruits show the delay senescence phenotype. Ear removal in maize plants can lead to either rapid or delayed leaf senescence, depending on genotype (Craftsbrandner and Poneleit, 1987). Leaf senescence of Arabidopsis appears to be unaVected by the reproductive growth (Hensel et al., 1993; Nooden and Penney, 2001). C. Sugars Sugars are known to act as signaling molecules at various stages of plant development (Rolland et al., 2002). Although expression of several senescence‐associated genes (SAGs) is induced by sugar starvation (Fujiki et al., 2000, 2001), as mentioned previously, a reduced sugar level resulting from a decline in photosynthesis is unlikely a major senescence‐inducing factor. More and more studies suggest that increased sugar levels or sugar signals induce leaf senescence. First, sugar levels are found higher in senescing leaves of Arabidopsis and tobacco plants than nonsenescing leaves (Masclaux et al., 2000). Second, when yeast invertase is expressed in the extracellular space of Arabidopsis, tobacco, and tomato plants, sugars are accumulated, and the leaves of the transgenic plants undergo premature senescence (Dickinson et al., 1991; Ding et al., 1993). Third, tomato plants overexpressing hexokinase (HXK), which is believed to serve as the sugar sensor (Rolland et al., 2002), have an increased sensitivity to sugars; leaves of these plants become precociously senescent (Dai et al., 1999). In contrast, the Arabidopsis hxk1 knockout/null mutant plants display a delayed leaf
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senescence phenotype even in the presence of high levels of sugars (Moore et al., 2003). A subsequent study on invertase, however, challenged the idea that high levels of sugar induce leaf senescence. An extracellular invertase under the control of the senescence‐specific SAG12 promoter resulted in a delay of leaf senescence in tobacco plants (Lara et al., 2004). Because of the senescence specificity of the SAG12 promoter, invertase gene expression in this study was targeted to senescing leaves only and the invertase activity in young leaves was unchanged (Lara et al., 2004), which is in contrast to the previously discussed studies in which the invertase was expressed in early stages of leaf development in addition to the senescing leaves. However, it should be noted that the accumulation of sugars in these SAG12‐invertase plants might not have reached the threshold for inducing senescence. In other words, this study is unnecessarily against the role of high sugar levels in the induction of leaf senescence. Because sugars can serve as signal molecules and an important energy resource, their role in triggering leaf senescence may be complex. DiVerent types or diVerent levels of sugars, for example, may have diVerent impacts on signaling pathways and/or sink‐source relation, both of which are important modulating factors of leaf senescence.
D. Stresses Leaf senescence can be induced by a number of diVerent environmental stresses such as extreme temperature, nutrient deficiency, water stress, pathogen infection, and oxidative stresses induced by ozone or UV‐B (Buchanan‐ Wollaston et al., 2003; Smart, 1994). Expression of SAGs can be induced in leaves exposed to many types of stress such as darkness (KleberJanke and Krupinska, 1997; Lin and Wu, 2004; Weaver et al., 1998), drought (Pic et al., 2002; Weaver et al., 1998), pathogen infection (Butt et al., 1998; Pontier et al., 1999), ozone treatment (Miller et al., 1999), UV‐B exposure (John et al., 2001), and oxidative stress (Navabpour et al., 2003). Many defense‐ related (DR) genes, on the other hand, are induced in leaves during natural senescence (Quirino et al., 2000). Pathogenesis‐related (PR) genes, for example, are found to be expressed during the senescence of healthy leaves (Hanfrey et al., 1996; Quirino et al., 1999). Large‐scale analyses of gene expression in senescing leaves of Arabidopsis thaliana (Gepstein et al., 2003; Guo et al., 2004; Lin and Wu, 2004) and Populus tremula (Andersson et al., 2004; Bhalerao et al., 2003) revealed that DR genes make a significant portion of leaf senescence transcriptome. An expression profiling of Arabidopsis transcription factors revealed that 28 of the 43 transcription factor genes that are induced during senescence are also induced by various
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stressors, suggesting extensive overlap between senescence and stress responses (Chen et al., 2002). A common factor involved in diVerent stress responses and senescence is increased levels of ROS (Jimenez et al., 1998; Lin and Kao, 1998; Munne‐ Bosch and Alegre, 2002), which may mediate the upregulation of the same group of genes during senescence as well as in response to stress treatments. Senescence is a very diVerent process from stress responses, however. For many of the DR genes identified as SAGs, their expression patterns in senescence are diVerent from that in stress responses (Buchanan‐Wollaston et al., 2003; Quirino et al., 2000). It is still not clear whether the DR genes are causes or just consequences of senescence. In many cases, stress‐response pathways appear to be involved after the onset of senescence. Products of the DR genes could function in cellular detoxification and the maintenance of cell viability during the senescing process in leaves. E. Ethylene The correlation between ethylene production and leaf senescence in climacteric species supports a regulatory role of ethylene in senescence. Symptoms of leaf senescence in many plant species can be induced (Horton and Bourguoin, 1992; Reyes‐Arribas et al., 2001; Zacarias and Reid, 1990) or retarded (Ella et al., 2003; Philosophhadas et al., 1994; Podwyszynska and Goszczynska, 1998) by application of exogenous ethylene or its antagonists, respectively. When ethylene biosynthesis is suppressed by using antisense technology, leaf senescence in transgenic tomato plants is delayed (John et al., 1995). Leaf senescence is also delayed in ethylene‐insensitive mutants such as etr1–1 (Grbic and Bleecker, 1995; Zacarias and Reid, 1990) and ein2/ora3 (Oh et al., 1997). However, constitutive overproduction of ethylene in Arabidopsis and tomato plants did not cause precocious senescence, suggesting that ethylene alone is not suYcient to initiate leaf senescence (Grbic and Bleecker, 1995). It has been postulated that age‐dependent factors are required for ethylene‐regulated leaf senescence (He and Gan, 2003). F. Jasmonic Acid (JA) Methyl jasmonate (MeJA) and its precursor JA were first identified as bioactive substances that promote senescence in detached oat leaves (Ueda and Kato, 1980). It has been shown that JA levels in senescing leaves are fourfold higher than those in nonsenescing ones in Arabidopsis, and genes encoding enzymes that catalyze most of the reactions of the JA biosynthetic pathway are diVerentially activated during leaf senescence (He et al., 2002).
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Exogenous application of JA at a physiological level of 30 M causes premature senescence in attached and detached leaves in wild‐type Arabidopsis but fails to induce precocious senescence in the JA‐insensitive mutant coi1 plants (He et al., 2002). JA application was also shown to enhance expression of a subset of SAGs in Arabidopsis (Park et al., 1998; Schenk et al., 2000). Several Arabidopsis mutants that are deficient in JA production or JA signal transduction do not exhibit a delayed leaf senescence phenotype (He et al., 2002). This may challenge the JA role in senescence. It is, however, possible that other factors may induce leaf senescence in the absence of JA. G. Salicylic Acid (SA) The levels of endogenous SA are fourfold higher in senescing leaves than that in nonsenescing leaves (Morris et al., 2000). SA treatment induces premature leaf senescence in Arabidopsis. The Arabidopsis plants that produce much less SA due to overexpression of a SA‐degrading gene called NahG and the Arabidopsis mutants defective in SA‐signaling such as npr1 and pad4 display a retarded leaf senescence phenotype with altered expression of some SAGs (Morris et al., 2000). It has been reported that treatment with SA upregulates the expression of several SAGs (NIT2, AtOSM34, SAG25, SAG26, and SAG29) (Quirino et al., 1999). However, SA accumulation is not essential for the induction of these SAGs during natural leaf senescence in Arabidopsis (Quirino et al., 1999). The role of SA in leaf senescence remains uncertain. H. Brassinosteroids (BRs) Two lines of evidence support a leaf senescence‐promoting role of BRs. First, external application of 24‐epi‐brassinolide (eBR) induces senescence in mung bean leaves (He et al., 1996) and cucumber cotyledons (Zhao et al., 1990), and eBR induces a subset of potential SAGs in Arabidopsis (He et al., 2001). Second, several Arabidopsis mutants that are deficient either in BR biosynthesis such as det2 or in the BR signaling pathway such as bri1 display a delayed leaf senescence phenotype (Clouse and Sasse, 1998), and the bri1 suppressor mutant exhibits an accelerated leaf senescence phenotype (Yin et al., 2002). I. Abscisic Acid (ABA) Abscisic acid ABA promotes senescence of detached leaves of various plant species, but it is less eVective in in planta leaves (Hung and Kao, 2003; Smart, 1994; Weaver et al., 1998). Although levels of ABA increase
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when a plant is under environmental stress conditions such as drought, high levels of salt, and low temperatures and these conditions often induce leaf senescence, various Arabidopsis mutants defective in either ABA biosynthesis or signaling have no measurable eVects on the timing or progression of leaf senescence in Arabidopsis (Hensel et al., 1993). ABA is considered an enhancer rather than a triggering factor of leaf senescence (He and Gan, 2003).
J. Cytokinins In contrast to the previously mentioned senescence inducers, some plant growth substances such as cytokinins, polyamines, and nitric oxide suppress leaf senescence. Three approaches have been employed to investigate the inhibitory role of cytokinins in leaf senescence: external application of cytokinins, measurement of endogenous cytokinin levels before and during senescence, and manipulation of endogenous cytokinin production in transgenic plants (Gan and Amasino, 1996). Exogenous application of cytokinins (e.g., zeatin, and benzyladenine) or their analogs (e.g., 1‐phenyl‐ 3‐[1,2,3‐thiadiazol‐5‐yl]‐urea) delays leaf senescence and even causes regreening of yellowing leaves in a variety of plant species such as soybean, tobacco, and Arabidopsis (Gan and Amasino, 1996; Nooden and Letham, 1993). In tobacco and many other plant species, levels of cytokinins in leaves decrease with the progression of senescence (Singh et al., 1992). In a transgenic study, the leaf senescence‐specific promoter of SAG12 in Arabidopsis was used to direct the expression of isopentenyl transferase (IPT ). IPT catalyzes the first, and rate‐limiting, step in the biosynthesis of cytokinins. When senescence onsets in a somatic leaf cell, the SAG12 promoter is activated to direct IPT expression, resulting in the biosynthesis of cytokinins. The increase in endogenous cytokinins will in turn inhibit senescence. Because senescence is inhibited, the senescence‐specific promoter becomes attenuated or inactivated, which prevents overproduction of cytokinins. This forms an autoregulatory senescence inhibition system. Tobacco plants harboring this system display a significantly delayed leaf senescence phenotype (Gan and Amasino, 1995). A similar phenotype has been observed in other transgenic plants harboring the SAG12‐IPT transgene, including lettuce (McCabe et al., 2001), bok choy (Yuan et al., 2002), broccoli (Chen et al., 2001), and petunia (Chang et al., 2003). Overexpression of components of the cytokinin signal transduction pathway also delays leaf senescence in Arabidopsis (Hwang and Sheen, 2001), which further confirms the inhibitory role of cytokinins in leaf senescence.
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K. Polyamines (PAs) Putrescine, spermidine, and spermine are the major members of PAs in plants. Although levels of PAs are higher in actively dividing cells than those in cells that do not grow and divide (Kaur Sawhney et al., 1985), PAs have been shown to inhibit leaf senescence. In many plant species and experimental systems, exogenous application of PAs can retard leaf senescence by preventing chlorophyll loss and membrane peroxidation and by inhibiting RNase and protease activities (Borrell et al., 1997; Evans and Malmberg, 1989; Sood and Nagar, 2003). S‐adenosylmethionine is the common substrate for the biosynthesis of PAs and ethylene in plants. When ethylene production is blocked by antisense technology, S‐adenosylmethionine is channeled to the PA biosynthesis pathway and PA levels in transgenic tobacco plants are significantly elevated and stress‐induced leaf senescence is prevented (Wi and Park, 2002). However, because ethylene promotes leaf senescence, the blocking of ethylene biosynthesis may at least in part contribute to the delayed leaf senescence phenotype in the transgenic tobacco plants.
L. Nitric Oxide (NO) Studies suggest that NO may play a role in retarding leaf senescence. In senescent pea leaves, NO production was reduced and NO synthase activity in peroxisomes was downregulated by 72% (Corpas et al., 2004). ABA and MeJA induced leaf senescence in rice, and the induction was oVset by exogenous application of NO. Furthermore, this antagonist eVect of NO was eliminated by an NO‐specific scavenger, 2‐(4‐carboxy‐2‐phenyl)‐ 4,4,5,5‐tetramethylimidazoline‐1‐oxyl‐3‐oxide (Hung and Kao, 2004). Because NO can remove ROS including H2O2 in peroxisomes (del Rio et al., 2003) and the ROSs are senescence modulators as previously mentioned, it is not surprising that NO inhibits senescence.
III. The Cell Death Execution Process At the cellular level, the senescence program unfolds in an orderly manner. Chloroplasts, which contain up to 70% of the proteins in a leaf cell, are one of the first organelles to be targeted for breakdown. Other organelles, such as the peroxisome, also undergo biochemical changes as senescence proceeds. The nucleus, which is needed for gene transcription, and the mitochondria, which are essential for providing energy, remain intact until the
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last stages of the senescence (Inada et al., 1998; Nooden et al., 1997). Also associated with leaf senescence is the decline in the structural and functional integrity of the cellular membrane (e.g., lipid phase separations and membrane leakage) (Thompson et al., 1998, 2000). Accompanying the cytological changes, the leaf ’s contribution of fixed carbon to the plant diminishes with a sharp decline in photosynthesis. During senescence, nutrients such as nitrogen, phosphorus, and sugars released from the degradation of macromolecules in leaf cells are reallocated to growing organs or storage tissues (Quirino et al., 2000; Smart, 1994). As revealed by a transcriptome analysis (Guo et al., 2004), degradation processes appear to be predominant over biosynthesis during leaf senescence. Although for the entire Arabidopsis genome, genes annotated to be involved in catabolism are half as many as in anabolism, the number of ‘‘catabolism’’ genes expressed in senescing leaves doubles that of ‘‘anabolism’’ genes (Guo et al., 2004). A. Chlorophyll Degradation As indicated by the visible sign of yellowing, a leaf loses most of its chlorophyll during senescence. The pathway of chlorophyll degradation has been elucidated and several genes in the pathway have been cloned (Berghold et al., 2002; Takamiya et al., 2000). The cleavage of the tetrapyrrole ring to produce red chlorophyll catabolite (RCC) by Pheide a oxygenase (PaO) is the decisive step for chlorophyll catabolism (Hortensteiner et al., 1998; Matile et al., 1999), so the pathway is often referred to as the PaO pathway. PaO is a nuclear‐encoded enzyme and its activity increases dramatically during senescence (Matile et al., 1999). The expression of five predicted PaO genes in Arabidopsis was shown to be upregulated during dark‐induced leaf senescence (Lin and Wu, 2004). sid, a Festuca pratensis PaO‐defective mutant, displays a stay‐green phenotype (Vicentini et al., 1995). Other genes in the PaO pathway do not show enhanced expression during senescence (Lin and Wu, 2004; Takamiya et al., 2000), but the absence of one of the catabolic enzymes, RCC reductase, causes a rapid cell death in Arabidopsis acd2 mutant (Mach et al., 2001); the accelerated cell death is likely caused by the accumulation of free chlorophyll molecules that are released from the pigment‐protein complexes; the protein‐free chlorophyll molecules are highly reactive and toxic to the cells. It is believed that chlorophyll degradation is carried out to detoxify the phytotoxic compound when senescence is taking place, but not to recycle the nutrients. In fact, the final products of chlorophyll catabolism, nonfluorescent chlorophyll catabolites (NCCs), are deposited in the vacuole and none of the nitrogens in NCCs is recycled (Hinder et al., 1996; Tommasini et al.,1998).
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B. Protein Degradation Plant cells lose about two third of their soluble proteins during senescence (Inada et al., 1998). Up to 70% of leaf proteins are in the chloroplasts. How chloroplast proteins are degraded and mobilized is one of the research focuses in leaf senescence. It has been demonstrated that at least the initial steps in the degradation of chloroplast proteins occur in the intact organelles. Chloroplasts isolated from mature pea leaves are able to degrade stromal proteins including the predominant enzyme Rubisco (Mitsuhashi and Feller, 1992; Roulin and Feller, 1998). As supported by global gene expression analyses, chloroplast localized protein‐degrading enzymes such as Clp protease (Andersson et al., 2004; Guo et al., 2004; Lin and Wu, 2004) and FtsH protease families (Andersson et al., 2004; Guo et al., 2004) could be involved in protein degradation in chloroplasts. ERD1 (Nakashima et al., 1997) and several other Clp family genes (Nakabayashi et al., 1999) were previously isolated as SAGs. These chloroplast proteases also play a role during leaf development (Majeran et al., 2000), although expression of genes encoding subunits of these proteases is not upregulated (but constitutive or downregulated) during leaf senescence (Craftsbrandner et al., 1996; Humbeck and Krupinska, 1996). It is most unlikely that these families of proteases are responsible for the rapid reduction of chloroplast proteins. Several reports indicate that in senescing leaves Rubisco could be degraded by vacuolar proteases (Minamikawa et al., 2001; Yoshida and Minamikawa, 1996). Substantial upregulation of vacuolar cysteine proteases during leaf senescence has been well documented (Buchanan‐Wollaston et al., 2003; He and Gan, 2003) and supported by global transcriptome analyses (Bhalerao et al., 2003; Gepstein et al., 2003; Guo et al., 2004) (Table I). Vacuolar proteases may play an important role in degradation of chloroplast proteins, at least at the final lytic stages after membrane is disrupted. At earlier stages of leaf senescence, when chloroplast membranes are still intact, chloroplast protein degradation by vacuolar proteases may take place through the association of chloroplasts with the central vacuole, which is supported by electron microscopic study on senescing leaves of French bean (Minamikawa et al., 2001). One possible mechanism of the chloroplast–vacuole association is the autophagic pathway. Autophagy is a ubiquitous eukaryotic process responsible for sequestration of substrate proteins, bulk cytosolic constituents, and organelles into the vacuoles for degradation. The number and the size of chloroplasts are remarkably reduced during senescence (Inada et al., 1998), which may result from the autophagic sequestration of part and/or whole chloroplasts into vacuoles. Transcripts of six Autophagy genes (APGs) were detected in Arabidopsis senescent leaves (Guo et al., 2004). Expression of APG7 (Doelling
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Table I
Senescence‐Associated Cysteine Proteases
Gene Name
Accession No.
Plant Species
Predicted Subcellular Localization
SAG2 (At5g60360)
AF360273
Arabidopsis
Secretory pathway
RD21 (At4g11310) SAG12 (At5g45890)
AY093066
Arabidopsis
Cytoplasm
AF370131
Arabidopsis
Secretory pathway
X99936 M21444 Z48736 U31094
Maize Tomato Tomato Petunia
Secretory pathway Secretory pathway Secretory pathway
Brassica napus
Cytoplasm
See1 SENU2 SENU3 PeTh3 LSC7 LSC790
U68221
Brassica napus
Cytoplasm
SmCP TPE4A Cysteine protease NTCP‐23 SPG31 TDI‐65 PsELSA Pcyprot1 Pcyprot2 Pcyprot3 Pcyprot4 Pcyprot5 Pcyprot6 Pcyprot7 Pcyprot8 Pcyprot9 HvSF42
AF082181 AJ004958 AJ249847
Secretory pathway Secretory pathway Secretory pathway
AJ496571
Eggplant Pea Lolium multiflorum Tomato Sweet potato Tomato Pea Populus tremula Populus tremula Populus tremula Populus tremula Populus tremula Populus tremula Populus tremula Populus tremula Populus tremula Barley
At4g01610
AF370193
Arabidopsis
Secretory pathway
At4g16190 At1g47128
AY136316 AY072130
Arabidopsis Arabidopsis
Secretory pathway Cytoplasm
At4g39090
AY080598
Arabidopsis
Secretory pathway
At2g27395 At1g02300
NM_128300 AY039887
Arabidopsis Arabidopsis
Cytoplasm Cytoplasm
AB032168 AF242185 AF172856 AJ278699
Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory Secretory
pathway pathway pathway pathway pathway pathway pathway pathway pathway pathway pathway pathway pathway
Reference Gepstein et al., 2003; Guo et al., 2004; Hensel et al., 1993 Koizumi et al., 1993 Guo et al., 2004; Lohman et al., 1994 Smart et al., 1995 Drake et al., 1996 Drake et al., 1996 Tournaire et al., 1996 Buchanan‐ Wollaston, 1997 Buchanan‐ Wollaston and Ainsworth, 1997 Xu and Chye, 1999 Cercos et al., 1999 Li et al., 2000 Ueda et al., 2000 Huang et al., 2001 Harrak et al., 2001 Pic et al., 2002 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Bhalerao et al., 2003 Scharrenberg et al., 2003 Gepstein et al., 2003; Guo et al., 2004 Guo et al., 2004 Gepstein et al., 2003; Guo et al., 2004 Gepstein et al., 2003; Guo et al., 2004 Guo et al., 2004 Guo et al., 2004
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et al., 2002) and APG8 (Doelling et al., 2002; Gepstein et al., 2003) are upregulated during senescence. All these data support the existence of the autophagic pathway in senescing leaves. The role of the autophagic pathway in leaf senescence is, however, puzzling because Arabidopsis plants with disrupted APG7 (Doelling et al., 2002) or APG9 (Hanaoka et al., 2002) exhibit an accelerated, not delayed, leaf senescence phenotype. It is possible that the autophagy may not be the only chloroplast protein degradation pathway. It is also possible that the autophagy may be involved in keeping the cells viable. Plastoglobuli (lipid‐protein globules) secreted from chloroplasts of senescing soybean leaf could be an alternative pathway of vacuolar or cytoplastic degradation of chloroplast protein (Guiamet et al., 1999). Proteins in the cytoplasm of the cells are likely degraded through the ubiquitin pathway. Expression of a large portion of genes in the ubiquitin‐ 26S proteasome pathway is associated with leaf senescence in Arabidopsis (Gepstein et al., 2003; Guo et al., 2004; Lin and Wu, 2004) and Populus tremula (Bhalerao et al., 2003). Mutation of ORE9, an F‐box protein that interacts with the plant SCF complex (component of the ubiquitin E3 ligase complex), causes a delay of leaf senescence in Arabidopsis (Woo et al., 2001). Another Arabidopsis mutant named dsl1 that is defective in arginyl tRNA: protein transferase (R‐transferase) also exhibits a delay in leaf senescence (Yoshida et al., 2002). R‐transferase is a component of the N‐end‐rule proteolytic pathway, which transfers arginine to the N‐terminus of proteins with N‐terminal glutamyl or aspartyl residues, thereby targeting the proteins for ubiquitin‐dependent proteolysis (Varshavsky, 1997). The ubiquitin‐mediated degradation of specific proteins appears to have an important role in the control of senescence. It has been postulated that ORE9 and DSL1 may play regulatory roles in leaf senescence through degrading proteins that negatively regulate leaf senescence.
C. Lipid Degradation During senescence there is a decline in the structural and functional integrity of cellular membranes as a result of enhanced catabolism of membrane lipids (Thompson et al., 1998). Putative lipid‐degrading enzymes include phospholipase D, phosphatidic acid phosphatase, lytic acyl hydrolase, lipoxygenase, ‐galactosidase, ‐galactosidase, and galactolipase. Genes encoding these enzymes have been shown to be well represented in senescence EST database (Guo et al., 2004). Their transcript levels are increased as revealed by microarray analysis (Lin and Wu, 2004) or suppression substrative hybridization analysis (Gepstein et al., 2003). A rice gene encoding alkaline ‐galactosidase, Osh69, has been studied. The Osh69 protein is specifically localized in the chloroplast of senescing
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leaves and is able to hydrolyze galactolipids, the major component of thylakoid membranes (Lee et al., 2004). Phospholipase D has been reported to be involved in the regulation of ABA or ethylene‐induced leaf senescence (Fan et al., 1997). Transgenic Arabidopsis plants with reduced levels of a senescence‐enhanced lipase showed a delayed leaf senescence phenotype (Thompson et al., 2000). The Arabidopsis SAG101, which encodes an acyl hydrolase, is induced at the early stages of leaf senescence and its expression increases with the progression of leaf senescence (He and Gan, 2002). Antisense suppression of SAG101 retards the progression of leaf senescence, whereas inducible overexpression of the gene promotes precocious senescence in young leaves (He and Gan, 2002). These results suggest that the degradation of lipids may promote leaf senescence.
D. Nutrient Recycling The nutrients released from the degradative processes just mentioned are translocated to actively growing regions such as young leaves, floral buds, and developing fruits and seeds. In Arabidopsis, levels of C, Cr, Cu, Fe, K, Mo, N, P, S, and Zn drop by greater than 40% during leaf senescence, but the nutrient recycled to the greatest extent is N (90%) (Himelblau and Amasino, 2001). In a senescing leaf, N is released primarily from protein degradation and nucleic acid catabolism (Hortensteiner and Feller, 2002) (Fig. 1). The released N is assimilated, generally in the form of ammonium, via the GS/GOGAT cycle that is carried out by the concerted action of glutamine synthetase (GS) and glutamate synthase (GOGAT). The Arabidopsis genome encodes eight cytosolic and one chloroplastic GSs. The chloroplastic GS is reported to be downregulated during leaf senescence (Bernhard and Matile, 1994) while several cytosolic GSs are upregulated during leaf senescence in several plant species (Andersson et al., 2004; Bernhard and Matile, 1994; Buchanan‐Wollaston and Ainsworth, 1997; Gepstein et al., 2003; Kamachi et al., 1992a; Masclaux et al., 2000). Glutamine (Gln) is the major mobile amino acid involved in the long distance transport of N. Consistent with this, expression of the cytosolic GSs is reported to be restricted to the phloem (Carvalho et al., 1992; Edwards et al., 1990; Kamachi et al., 1992b; Oliveira et al., 2002). In contrast to the chloroplastic GS that functions in concert with ferredoxin (FD)‐dependent GOGAT (Coschigano et al., 1998), the cytosolic GSs are often coupled with the NADH‐dependent GOGAT that is low in leaves but abundant in nonphotosynthetic tissues such as roots (Lancien et al., 2002). Fd‐GOGAT activity is reported to decrease in senescing leaves of grapevine (Loulakakis et al., 2002) and wheat plants (Cincerova et al., 1991; Peeters and Vanlaere, 1992) while NADP‐GOGAT
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Figure 1 An outline of nutrient‐recycling pathways during leaf senescence. AS, Asparagine synthetase; ASPA, aspartate aminotransferase; GDH, glutamate dehydrogenase; GS, glutamine synthetase; ICL, isocitrate lyase; MS, malate synthase; NADP‐GOGAT, NADP‐dependent glutamate synthase; PPDK, pyruvate orthophosphate dikinase.
activity increases (Peeters and Vanlaere, 1992). A gene encoding NADP‐ GOGAT is expressed together with four cytosolic GS in senescent leaves of Arabidopsis (Guo et al., 2004). A senescing leaf loses its photosynthetic capability and may behave like a nonphotosynthetic organ. Genes encoding enzymes related to the GS/GOGAT cycle such as glutamate dehydrogenase (GDH) (Masclaux et al., 2000; Masclaux‐Daubresse et al., 2002), asparagine
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synthetase (AS) (Fujiki et al., 2001; King et al., 1995; Nozawa et al., 1999; Winichayakul et al., 2004), and aspartate aminotransferase (ASPA) (Gepstein et al., 2003; Yoshida et al., 2001) have been shown to be upregulated during leaf senescence of diVerent plant species. In senescing leaves, lipid catabolites are also believed to be remobilized. Fatty acids released can be converted to sugars through gluconeogenesis, or to organic acids (e.g., ‐ketoglutarate) via the glyoxylate cycle (Chen et al., 2000b) (Fig. 1). Expression of genes encoding key enzymes in these processes such as malate synthase (MS) (Graham et al., 1992), isocitrate lyase (ICL) (Chen et al., 2000a; Corpas et al., 1999), and pyruvate orthophosphate dikinase (PPDK) (Lin and Wu, 2004; Smart et al., 1995) has been shown to be enhanced during leaf senescence. Expression of genes encoding the vegetable storage proteins (VSPs) (Gepstein et al., 2003; Mira et al., 2002) and various nutrient transporters (Guo et al., 2004; Quirino et al., 2001) is associated with leaf senescence. EYcient transportation of nutrients between subcellular compartments may facilitate the recycling processes in senescing mesophyll cells.
IV. Gene Expression and Regulation During Leaf Senescence A. Gene Expression During Leaf Senescence Leaf senescence is under direct nuclear control and involves dramatic alteration in gene expression. In a senescing leaf, the majority of the genes that are expressed in green leaves, including those photosynthesis‐related genes, are downregulated, while a subset of genes, generally referred to as SAGs, are upregulated. SAG expression is required for senescence because inhibitors of both transcription and translation prevent leaves from senescing. For the past decade much eVort has been made to isolate SAGs, and hundreds of SAGs have been cloned from various plant species including Arabidopsis, barley, Brassica, maize, cucumber, rice, tobacco, radish, asparagus, and soybean (Buchanan‐Wollaston et al., 2003; Gepstein et al., 2003; He and Gan, 2003). Until recently, application of genomics approaches has led to the identification of thousands of potential SAGs (Andersson et al., 2004; Bhalerao et al., 2003; Buchanan‐Wollaston et al., 2003; Guo et al., 2004; Lin and Wu, 2004; Zentgraf et al., 2004). Microarray analyses of Arabidopsis cDNAs revealed that approximately 20% of the studied genes change their expression during leaf senescence (Buchanan‐Wollaston et al., 2003; Zentgraf et al., 2004). According to the result of large‐scale single‐pass sequencing of a senescent Arabidopsis leaf cDNA library, approximately 2500 unique genes are expressed in senescing leaves (Guo et al., 2004). As discussed previously, diVerential gene expression in leaf senescence could be triggered by many
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external and internal factors. The expression of thousands of SAGs leads to the execution of senescence including the degradation of various macromolecules and remobilization of diVerent nutrients. DiVerent signals often induce diVerent sets of genes (He et al., 2001; Park et al., 1998; Weaver et al., 1998), which may in turn initiate diVerent biochemical/physiological processes (Fig. 2 ). It has been postulated that multiple pathways are interconnected to form a regulatory network that controls leaf senescence (Gan and Amasino, 1997). A simplified version of the network has been revealed by using Arabidopsis leaf senescence enhancer trap lines (He et al., 2001).
Figure 2 A simple model to illustrate the molecular regulatory mechanisms by which various internal or external signals promote or retard leaf senescence. The senescence promoting signals are most likely to exert their eVect by activating a subset of senescence‐associated genes (SAGs) and by suppressing the expression of senescence downregulated genes (SDGs). In contrast, the retardation of leaf senescence by senescence‐inhibiting signals is most likely achieved by activating SDGs and possibly by suppressing SAGs. Expression of SAGs and SDGs eventually leads to the execution of leaf senescence. ABA, Abscisic acid; BR, brassinosteroid; JA, jasmonic acid; NO, nitric oxide; PA, polyamines; RG, reproductive growth; SA, salicylic acid.
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B. Perception and Transduction of Senescence Signals The perception and signal transduction of several senescence‐inducing factors such as sugars, environmental stresses, and phytohormones have been investigated and light has been shed on some of the signals. The perception and signal transduction of senescence‐inducing hormone ethylene are perhaps best characterized thus far (Klee, 2004), while the mechanism by which age is sensed to initiate senescence is least understood. Receptor kinases may serve as receivers and/or transducers of internal or external signals. Genes encoding receptor‐like kinase (RLK) such as SARK in tomato (Hajouj et al., 2000), lecRK‐a1 (Riou et al., 2002), SIRK (Robatzek and Somssich, 2002), and At5g48380 (Gepstein et al., 2003) in Arabidopsis, and Paul27 in Populus tremula (Bhalerao et al., 2003) are induced by leaf senescence. The Arabidopsis genome has a large gene family of RLKs consisting of more than 610 genes (Shiu and Bleecker, 2003). Transcripts of 44 RLK genes are found in senescent leaves (Guo et al., 2004). Functional study of these RLK genes will help understand signal perception of leaf senescence. The mitogen‐activated protein kinase (MAPK) signal cascades, MAPKKK‐MAPKK‐MAPK, are ubiquitously used by eukaryotic cells to link extracellular stimuli to a wide range of cellular responses (Ichimura et al., 2002). This type of signal cascade via protein phosphorylation may play an important role in signal transduction of leaf senescence. Arabidopsis genes encoding for 3 MAPKs, 3 MAPKKs, 9 MAPKKKs, and 1 MAPKKKK are represented in the aforementioned senescent leaf EST database (Guo et al., 2004). Populus tremula MAPK gene Paul31 is among the most strongly expressed genes unique to autumn senescing Populus leaves (Bhalerao et al., 2003). A maize MAPK, zmMPK5, has been reported to accumulate in senescent leaves. A MAPKKK‐encoding Arabidopsis gene, EDR1, was cloned as a negative regulator of disease resistance (Frye and Innes, 1998). The kinase activity of EDR1 has been confirmed and the kinase domain has been shown to possess autophosphorylation activity (Tang and Innes, 2002). The edr1 mutant as well as the Arabidopsis plants overexpressing a kinase‐ deficient EDR1 gene showed an enhanced ethylene‐induced senescence phenotype (Tang and Innes, 2002). These data suggest an overlap between the genetic control of senescence and the SA‐induced defense responses. Other signal transduction pathways such as calcium signaling, G‐protein‐ mediated signaling, phosphatases, and 14‐3‐3 proteins may also be involved in leaf senescence. Possible associations of calcium‐dependent protein kinases, calcium‐binding proteins, G‐proteins, protein phosphatases, and 14‐3‐3 proteins with leaf senescence have been reported in a variety of plant species (Gepstein et al., 2003; Guo et al., 2004; Guterman et al., 2003; Yang and Poovaiah, 2000).
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C. Transcription Regulation of Gene Expression Signal transduction often leads to changes in gene expression. DiVerential gene expression is generally mediated by transcription factors that bind to specific cis elements of target gene promoters, resulting in activation and/or suppression of the target genes. There are approximately 1500 transcription factor genes in the Arabidopsis genome that belong to more than 30 gene families based on their DNA‐binding domains (Riechmann et al., 2000), and more than 130 of them are represented in the Arabidopsis leaf senescence EST collection (Guo et al., 2004). These senescence transcription factors are in the families of NAC, WRKY, AP2/EREBP, C2H2, C3H, and C2C2 zinc finger proteins, MYB, HB, bZIP, and bHLH, among others (Guo et al., 2004). Of the 402 transcription factor genes on a microarray, 43 genes are induced by senescence (Chen et al., 2002). Although gene families in this study have been preselected, predominant representation of gene families such as MYB, WRKY, AP2/EREBP, and zinc finger proteins is consistent with the senescence EST analysis. Similarly, analysis of an Arabidopsis whole‐genome array revealed that 303 and 81 transcription factor genes were upregulated and downregulated, respectively, during dark‐induced leaf senescence. These upregulated transcription factor genes belong to such gene families as AP2/EREBP (44 genes), bZIP (26), C3H (23), NAC (22), bHLH (22), WRKY (21), C2H2 (21), MYB (13), HB (11), GRAS (11), C2C2 (10), and others (Lin and Wu, 2004). It should be noted that many of these genes are also diurnal genes (estimated 24.7%) or induced by carbon starvation (estimated 19–26%) (Lin and Wu, 2004). WRKYs are perhaps the best studied among the leaf senescence‐ associated transcription factors. WRKY6 (Robatzek and Somssich, 2001) and SIRK (Robatzek and Somssich, 2002) are expressed during leaf senescence in Arabidopsis. SIRK encodes a receptor kinase. It is likely that WRKY6 binds to the promoter region of SIRK to regulate the expression of this gene. WRKY6 has also been shown to positively regulate PR1 (a defense‐related gene that is also expressed during leaf senescence), most likely through regulating the expression of the transcription factor gene NPR1 (Zhou et al., 2000). Another WRKY gene, WRKY53, was shown to be induced at early stage of leaf senescence (Hinderhofer and Zentgraf, 2001). Arabidopsis plants overexpressing this gene displayed an accelerated senescence phenotype. In contrast, when the gene was suppressed by using RNAi or insertional mutation, the onset of leaf senescence was delayed (Miao et al., 2004). These data suggest an important role of WRKY53 in regulating leaf senescence. Possible downstream target genes of WRKY53 include stress‐related genes, defense‐related genes, and some SAGs (Miao et al., 2004). Because there exists a highly conserved cis element called W‐box to which WRKY domain binds on the promoters of many WRKYs,
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it has been suggested that WRKYs form a regulatory transcription factor network to regulate such developmental processes as senescence and defense responses (Ulker and Somssich, 2004). Both WRKY6 and WRKY53 have other WRKY genes as putative downstream targets and can negatively regulate their own expression (Miao et al., 2004; Robatzek and Somssich, 2002). As to the upstream regulatory components of the senescence WRKYs, the MAP kinase cascade may be the right candidates. In fact, some WRKY transcription factors have been shown to function downstream of the MAP kinase cascade in defense responses in Arabidopsis (Asai et al., 2002; Wan et al., 2004) and tobacco (Kim and Zhang, 2004). To fully understand mechanisms by which SAGs are regulated, eVorts have been made to identify regulatory cis elements on the promoter regions of various SAGs. SAG12 encodes an apparent cysteine protease and is highly senescence specific in Arabidopsis (Gan, 1995). A 33‐base pair (bp) sequence in the promoter region of SAG12 is responsible for its senescence‐specific expression. This sequence seems to interact with diVerent proteins extracted from young or old leaves (Noh and Amasino, 1999a), suggesting that there is a transcription repressor to suppress the SAG12 expression in young leaves; this repressor is replaced by a newly transcribed or modified transcription factor to activate the SAG12 expression at the onset of senescence. This cis element sequence has been found on the promoter regions of two SAG12 homologues in Brassica napus (Noh and Amasino, 1999b). More recently, analysis of the promoter of a senescence‐induced asparagine synthetase (AS) gene of asparagus revealed that the 640 to 523 region of the AS promoter was responsible for the dark‐induced senescence response in detached asparagus leaves (Winichayakul et al., 2004). A 7‐bp core sequence TTGCACG was identified by comparing the cis element sequences of SAG12 and AS (Winichayakul et al., 2004). Whether this 7 bp is suYcient for the senescence specificity needs to be further investigated. Analyses of several other SAGs have also led to the identification of other potential senescence‐responsive sequences. For example, two cis elements, JASE1 and JASE2, have been identified from the promoter of OPR1 in Arabidopsis. These two elements are required in order for OPR1 to be expressed during leaf senescence and in response to JA (He and Gan, 2001). Although only several senescence cis elements have been identified so far, it is most likely that individual SAGs are regulated via unique mechanisms.
V. Closing Remarks Significant progress has been made in leaf senescence research. It has been demonstrated that initiation and/or progression of leaf senescence can be aVected by age, sugar levels, phytohormones, and other plant growth
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substances, and environmental cues. Perception of signals often leads to changes in gene expression, and the upregulation of thousands of SAGs causes the senescence syndrome: decline in photosynthesis, degradation of macromolecule, mobilization of nutrients, and ultimate cell death. Potential components in signal transduction such as receptor‐like kinases and MAP kinase cascade have been identified. Studies on the WRKY transcription factor gene family and the SAG12 senescence‐responsive cis element have shed some light on the regulatory mechanisms underlying leaf senescence. Because transcription factors are master gene regulators, immediate future research should be directed to the construction of the senescence transcription factor network by using integrated approaches involving DNA chip, chromosome immunoprecipitation (ChIP), and other molecular genetics tools.
Acknowledgments We thank Cara Winter for critical reading of the manuscript. Our senescence research has been supported by grants to S.G. from NSF (MCB‐0445596), DOE (DE‐FG02‐02ER15341, and DE‐ FG02‐99ER20330), USDA NRI (2001‐35304‐09994 and 2001‐35301‐10565) and Cornell Genomics Initiative.
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Muscle Stem Cells and Regenerative Myogenesis Iain W. McKinnell, Gianni Parise, and Michael A. Rudnicki Ottawa Health Research Institute, Ottawa, Ontario, K1H 8L6 Canada
I. Developmental Origin of Muscle II. Postnatal Muscle Repair III. Adult Stem Cells in Skeletal Muscle A. What Are the Contributing Sources of Adult Muscle Stem Cells? B. Is Pax7 a Master Regulator of Muscle Stem Cells? C. How Do CD45þ/Sca1þ Cells Become Directed Toward a Muscle Stem Cell Fate? IV. Interpretation of Findings from Models of Extreme Muscle Damage A. Is the Regenerated Muscle Functional? B. Would This Response Ever Occur Physiologically? References
A population of myogenic progenitors termed satellite cells undertakes postnatal development and repair of skeletal muscle. Studies have indicated that atypical myogenic precursors can also participate in muscle regeneration. The source of this regenerative capacity has been attributed to ‘‘adult stem cells’’ that represent poorly understood multipotent cell lineages, believed to reside in all adult tissue populations. Here we review the origin and location of muscle satellite cells and stem cells, as well as the mechanisms by which they may be specified. We discuss how the experimental models utilized raise important questions regarding the validity of extrapolating these findings. ß 2005, Elsevier Inc.
I. Developmental Origin of Muscle All skeletal muscle of the body originates from the somites. These structures form on either side of the midline as a consequence of a mesenchymal to epithelial transformation of the anterior portion of the segmental plate. The newly formed somite has a mesenchymal core surrounded by an epithelial sphere of cells. The somite undergoes a stereotypic program of diVerentiation. The onset of diVerentiation is marked by the ventromedial portion responding to signals from the notochord and neural tube by enacting an epithelial to mesenchymal transition to form the sclerotome, which together with the cells of the somitocoel gives rise to the axial skeleton. The dorsal Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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region remains as an epithelial layer, termed the dermomyotome, which goes on to form both the skeletal muscle and the dermis of the back and responds to signals emanating from the ectoderm, neural tube, and lateral plate mesoderm. The medial and lateral thirds of the dermomyotome (which form the epaxial and hypaxial musculature, respectively) represent the region of proliferating myogenic precursors and express genes such as Pax3 and to a lesser extent, Myf5. These precursors ingress beneath the dermomyotome to form the myotome, a sheet of postmitotic diVerentiating myoblasts that express members of the muscle regulatory factor family (e.g., MyoD). These mononucleate cells subsequently express markers of terminal diVerentiation (e.g., myosin heavy chain‐MHC) before fusing to form syncytiums that mature into contractile muscle fibers. During the late stages of embryonic myogenesis, a subset of cells termed satellite cells are observed; their role is discussed later, but it is worth noting that their embryonic origin remains somewhat unclear. Traditional embryological studies appeared to demonstrate that these cells were also of a somitic origin (Armand et al., 1983); however more recent evidence appears to indicate that they may be derived from the embryonic vasculature, in particular the dorsal aorta (De Angelis et al., 1999; Minasi et al., 2002). More extensive reviews of embryonic muscle development and the origins of embryonic stem cells have been done (Buckingham et al., 2003; Christ and Ordahl, 1995; Patel et al., 2002).
II. Postnatal Muscle Repair ‘‘. . .the presence of certain cells, intimately associated with the muscle fiber, have been observed which we have chosen to call satellite cells. Since these cells have not been reported previously and. . .as we shall suggest, might be pertinent to the vexing problem of skeletal muscle regeneration, a brief communication describing this finding is warranted. . .’’ Mauro, 1961
With admirable restraint and foresight, Alexander Mauro began the first anatomical definition of a muscle stem cell in 1961. Adult skeletal muscle is composed of multinucleated myofibers. Located between the sarcolemma and the basal lamina of each fiber are small mononucleated cells called satellite cells, which represent a self‐replenishing pool of muscle‐specific stem cells responsible for the vast majority of postnatal muscle growth and repair. It is worth revisiting Mauro’s initial observations, if only to make the point that his characterization of these cells via anatomical location and observations regarding the significant skew in the nucleus–cytoplasm ratio (manifest as an almost complete lack of cytoplasm) remains the only
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unequivocal criteria by which satellite cells are defined. Satellite cells are mitotically quiescent and do not express markers typically associated with myogenic potential or terminal diVerentiation. However, in response to injury and muscle degeneration, satellite cells forgo their anatomical properties, and in becoming activated, re‐enter the cell cycle and express muscle regulatory factors (Myf5 and MyoD). Subsequently they undergo a number of cycles of replication to form a pool of muscle precursor cells that can migrate to the site of injury, terminally diVerentiate, and fuse with existing myofibers, thus eVecting muscle repair. Strikingly, satellite cells persist in muscle that is subjected to repeated cycles of degeneration and regeneration, suggesting that self‐renewal or recruitment maintains the pool of satellite cells (reviewed in Seale and Rudnicki, 2000). The belief that satellite cells are responsible for all postnatal muscle repair was challenged by the finding that pluripotent stem cells purified from both bone marrow and adult skeletal muscle are able to participate in muscle repair (Ferrari et al., 1998; Gussoni et al., 1999). Significantly, this ability was shown to be restricted to a subpopulation of stem cells referred to as the side population (SP), which is readily isolated from the main population (MP) by fluorescence‐activated cell sorting (FACS) on the basis of Hoechst 33342 dye eZux (Fig. 1). Although both marrow SP (maSP) and muscle SP (mSP) cells could participate in the regeneration process, only mSP cells (and not maSP) appeared to give rise to satellite cells (Asakura et al., 2002; Gussoni et al., 1999), leading to speculation that mSP cells may be the progenitors of satellite cells.
Figure 1 Postnatal muscle regeneration. Mitotically quiescent satellite cells are activated in response to injury, at which time they eschew their definitive morphological properties and emerge from the satellite cell compartment by upregulating muscle regulatory factors, proliferating, and migrating to the site of injury. In addition, evidence suggests that alternative cell populations, isolated from muscle on the basis of Hoechst dye exclusion, are also able to contribute to muscle repair.
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The Myf5‐LacZ knock‐in allele recapitulates the expression of native Myf5 mRNA in myogenesis. However, whereas Myf5 mRNA or protein cannot be detected in satellite cells, the expression of Myf5‐LacZ is readily detected in the nuclei of satellite cells, and so can be used to localize their presence (Beauchamp, 2000). By using the Myf5‐LacZ mouse, researchers (Asakura et al., 2002) were able to identify in which population of adult stem cells the satellite cells reside. Crucially, after cell sorting by FACS and subsequent plating, all cells positive for LacZ were found to reside in the MP of Hoechst stained cells, with no LacZ‐positive cells detected in the SP fraction. Furthermore, culturing of SP and satellite cells allowed some investigation into their developmental potential and demonstrated that when cultured in the presence of cytokines, SP cells diVerentiated into hematopoietic cells but not muscle, whereas satellite cells were unable to exhibit hematopoietic potential, indicating that SP and satellite cells are distinct populations. However, SP cells do have the potential to diVerentiate into satellite cells after injection into regenerating muscle, and a percentage of muscle SP cells commit to muscle following coculture with satellite cell‐derived myoblasts. These observations suggested that muscle‐derived SP cells participate in the regeneration of skeletal muscle and that some portion of satellite cells may be derived from adult stem cells resident in skeletal muscle. There are several studies available for more in‐depth review of adult muscle repair (BischoV, 1994; Charge and Rudnicki, 2004; Goldring et al., 2002; Parker et al., 2003; Seale and Rudnicki, 2002; Seale et al., 2001).
III. Adult Stem Cells in Skeletal Muscle Skeletal muscle regeneration has long been considered to be mediated solely by satellite cells that function as the monopotential stem cell of skeletal muscle (Asakura et al., 2002). However, as previously discussed, novel populations of adult stem cells have been demonstrated to participate in regeneration of skeletal muscle. Work that has utilized various cell surface markers to purify adult stem cell populations from skeletal muscle, including c‐kit, Sca1, CD34, and CD45 (Dell’Agnola et al., 2002; Mahmud et al., 2002; McKinney‐Freeman et al., 2002; Romero‐Ramos et al., 2002; Tamaki et al., 2002; Torrente et al., 2001), would appear to have now established the significance of this process. The cell surface antigen Sca1, used routinely in purification protocols for hematopoietic stem cells, is expressed on over 80% of muscle SP cells. By contrast, the hematopoietic‐restricted lineage marker CD45 is expressed on about 16% of SP cells. Importantly, Sca1 and CD45 are not expressed on muscle satellite cells, and whereas almost all muscle‐ derived hematopoietic progenitor and in vivo blood reconstitution activity is
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þ
derived from CD45 cells (Asakura et al., 2002; McKinney‐Freeman et al., 2002), muscle‐derived CD45þ cells purified from uninjured muscle are uniformly nonmyogenic in vitro and do not form muscle in vivo (Asakura et al., 2002; McKinney‐Freeman et al., 2002). Coculture studies demonstrated that a large proportion of CD45þ:Sca1þ cells isolated from regenerating muscle acquire myogenic potential (Polesskaya et al., 2003) and in vivo injection experiments indicate that CD45þ SP as well as CD45 SP cells possess myogenic potential (Asakura et al., 2002; McKinney‐Freeman et al., 2002), representing a significant source of myogenic progenitors during muscle repair. Finally, the potential of bone marrow‐derived cells to give rise to satellite cells has been reported in transplant recipients (LaBarge et al., 2002). As such it is hypothesized that adult stem cells have a developmental relationship with the committed tier of muscle satellite cell progenitors and are fully capable of giving rise to satellite cells following muscle trauma. The relationship between CD45 :Sca1þ cells from SP preparations and CD45þ:Sca1þ preparations from whole muscle remains to be resolved. Developmental studies have suggested that satellite cells may originate from ‘‘mesangioblasts’’ that have the potential to contribute to many mesodermal cell lineages (Minasi et al., 2002). The ontological origin of such a cell would presumably be upstream of both Pax7þ myogenic progenitors and CD45þ hematopoietic stem cells. It can be postulated therefore that Sca1þ cells within the SP preparation may be upstream of a CD45þ stem cell, which is possibly transitional in nature and therefore more akin to an activated adult stem cell. Ongoing studies in many laboratories are addressing these emerging issues and the preceding developmental schema will continue to be revised as new knowledge is acquired.
A. What Are the Contributing Sources of Adult Muscle Stem Cells? The ability of circulating cells to participate in muscle repair and regeneration has been the topic of spirited debate for decades, and during this time of uncertainty concerning the fate of human embryonic stem‐cell research, perhaps never more pertinent. Since the turn of the century several reports have demonstrated that circulating cells integrate into various adult tissues including brain (Mezey et al., 2000), liver (Lagasse et al., 2000), heart (Bittner, 1999), and skeletal muscle (Blau et al., 2001). Early studies utilizing direct injection of unfractionated bone marrow‐derived cells (BMDCs) into muscle demonstrated that BMDCs could undergo myogenic diVerentiation and participate in muscle regeneration, albeit to very low levels. This suggested that only a small number of BMDCs were actually contributing to muscle regeneration (Ferrari et al., 1998). Attempts to enrich
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this ‘‘myogenic’’ population by fractionation into adherent and nonadherent populations failed (Ferrari et al., 1998), suggesting that subpopulations of each fraction were equally eVective in contributing to muscle regeneration. Attempts to enrich bone marrow‐derived myogenic progenitors have focused on specific cell surface proteins associated with ‘‘stem‐like’’ cells. Researchers (Corbel et al., 2003) demonstrated that a single hematopoietic stem cell (HSC) possessed the ability to fully repopulate the hematopoietic lineage, as well as contribute nuclei to regenerating muscle. These cells were characterized by the expression of c‐kit and Sca1, and did not express any Lin markers. In a similar fashion, it was demonstrated that a single hematopoietic cell characterized on the basis of Sca1 and CD45 expression also had the ability to fully repopulate the hematopoietic lineage and contribute to regenerating skeletal muscle (Camargo et al., 2003). Studies have demonstrated that cells expressing Mac1low increased in number in regenerating muscles following injury and demonstrated significant potential for myogenic activity, whereas cells expressing Mac1high demonstrated negligible myogenic activity (Ojima et al., 2004). These findings suggest that cells possessing myogenic potential residing in the SP are derived from the hematopoietic lineage. Further evidence has demonstrated that only those cell fractions containing the tyrosine kinase receptor c‐kit possess the ability to contribute to muscle fibers following intramuscular injection. Albeit interesting, most of these investigations suggest that marrow‐derived myogenic progenitors actually make minimal contributions to myogenesis during regeneration. In fact, the frequency of marrow‐derived contribution to skeletal muscle is reported to be between 0.01–0.1% in resting muscle and increasing to approximately 5% and as high as 12% of total fibers after injury, demonstrating that muscle injury plays an integral role in BMDC homing to skeletal muscle (Abedi et al., 2004; Dreyfus et al., 2004). Despite significant advances, many of the findings in these studies have not yet been reconciled, and the identification of the optimal HSC for muscle regeneration remains elusive. Continued exhaustive experiments aimed at identifying unique cell surface proteins and establishing a genetic fingerprint of HSCs with myogenic potential are necessary to fully take advantage of the potential for BMDCs in therapeutic environments. However, it is important to acknowledge that significant headway must be achieved before these cells become a viable option for therapy. A second hotly debated topic is whether the observed myogenic contribution from circulating cells is a directed process in response to environmental cues, or whether myogenic contribution simply represents a cell‐fusion event between myeloid cells and existing myofibers. As previously discussed, it has been reported that cells expressing Mac1low increased in number in regenerating muscles following injury and expressed significant myogenic activity,
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whereas cells expressing Mac1 demonstrated negligible myogenic activity (Ojima et al., 2004). In addition, others (Doyonnas et al., 2004) have reported that immature myelomonocytic cells, expressing c‐kit, contributed significantly to muscle regeneration following intramuscular injection. Conversely, mature myelomonocytic cells made no significant contribution to regenerating muscle fibers, ruling out the possibility that vascular contribution to muscle fiber regeneration is simply a fusion event involving mature macrophages, and other hematopoietic cells associated with the inflammatory response (Ojima et al., 2004). Data from other work challenged this notion (Camargo et al., 2003). They used a LysM‐Cre transgene, which is highly expressed in immune cell types, linked to a LacZ reporter. Following muscle regeneration the investigators described rare LacZ‐positive muscle fibers, suggesting that cells of the myeloid lineage can fuse with regenerating muscle fibers and provide support for the synthesis of new myotubes. Although an attractive and perhaps logical hypothesis given the substantial inflammatory response following muscle damage and the necessity of the inflammatory response for normal muscle regeneration, there is good reason to question these findings. First, evidence suggests that LysM does not mark myeloid commitment and that a population of HSCs express LysM and are fully pluripotent (Ye et al., 2003). Thus, it is entirely possible that what was observed was a legitimate contribution to muscle fibers via HSC commitment to the myogenic lineage, as opposed to a myeloid fusion event (Camargo et al., 2003). Certainly the most compelling evidence of directed commitment of HSCs to muscle is the observation that BMDCs follow a biological progression from the vasculature, as an immature myelomonocytic cell, into skeletal muscle, giving rise to muscle satellite cells and finally, fusing to form mature myofibers (Dreyfus et al., 2004; LaBarge and Blau, 2002). Researchers (Dreyfus et al., 2004) demonstrated that following GFPþ bone marrow transplantation into irradiated muscle, the proportion of GFPþ cells in the satellite cell position increased from 4% to 14% between 1 and 6 months after transplantation. Furthermore, this was associated with the observation of scattered GFPþ muscle fibers (1.3%) between 3 and 6 months after transplantation. Thus, the contribution to muscle regeneration via myeloid cell fusion, if in fact it does occur, is an exceedingly rare event, and it begs the question as to whether there is any physiological significance for such a phenomenon. The more likely scenario is that myeloid cell fusion events, with respect to muscle regeneration, do not play a directed role or contribute significant numbers of cells in muscle regeneration. Despite an abundance of scientific data supporting the involvement of BMDCs in muscle repair and regeneration, several questions remain regarding the physiological significance of their contribution to muscle regeneration (Fig. 2). We have shown that cells expressing CD45 and Sca1, residing in muscle, do not demonstrate any intrinsic myogenic potential when taken
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from uninjured muscle. However, following cardiotoxin injection, CD45þ cells expand about 30‐fold in number, acquire myogenic potential, and can readily diVerentiate into myotubes, suggesting that CD45þ cells play a physiological role in muscle regeneration. A working hypothesis is that adult stem cells are a resident population in skeletal muscle and do not originate from cells that transit from the marrow in response to muscle injury. There are several lines of evidence that suggest that CD45þ stem cells are a resident and stable population within muscle. First, numerous investigations have demonstrated that high‐dose local irradiation of limbs results in a prolonged period of attenuated growth and regeneration, suggesting that circulating stem cells may not play a significant role in muscle repair (Heslop et al., 2000; Pagel and Partridge 1999; Wakeford et al., 1991). Second, transplanted muscles do not incorporate host nuclei following injury and regeneration (Schultz et al., 1986). Third, we observed a 30‐fold reduction in the total number of satellite cells 18 hours after cardiotoxin, suggesting that the satellite cells did not survive the insult. By 4 days post‐ctx, the total number of myogenic progenitors increased to about 6.0 106 Myf5‐LacZ expressing
Figure 2 Possible contributions of BMDCs to muscle repair. Model based on findings regarding the potential of BMDCs as determined by their level of Mac1 expression. Mac1high cells represent mature macrophages that may partake in random fusion events into existing muscle fibers. Mac1low cells represent immature myelomonocytic cells with the ability for directed myogenic diVerentiation.
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cells per gram of muscle tissue. Together, these experiments suggest that circulating stem cells do not significantly contribute to stem‐cell pools within regenerating muscle tissue. Expansion of myogenic progenitors to this level appears to exceed the capacity of the TA muscle, given the starting number of satellite cells and the fact that satellite cells do not migrate between muscle groups (BischoV and Heintz, 1994; Schultz et al., 1986). Finally, findings concerning the origin of CD45þ cells resident in skeletal muscle suggest that less than 20% of the CD45þ population in muscle is derived from bone marrow, and that the multipotent muscle SP is entirely derived from tissue other than bone marrow (Rivier et al., 2004). This finding, however, is confounded by the possibility that the muscle SP and other muscle HSCs may be resistant to irradiation, and therefore simply did not turn over during the course of the experiments. Taken together, these results suggest that CD45þ/Sca1þ cells may be a resident and stable muscle population.
B. Is Pax7 a Master Regulator of Muscle Stem Cells? A question of some significance arises from the preceding sections; given that there appear to be various mechanisms for muscle repair, the potential cellular origins of which are numerous, are there multiple independent signaling pathways that recruit, activate, and maintain the cells responsible, or do all pathways begin or converge upon the same molecule(s) upstream of the myogenic diVerentiation program? Utilizing representational diVerence analysis, it was demonstrated that the paired box transcription factor Pax7 is specifically expressed in satellite cells in vivo and in areas of regenerating muscle in the mdx mouse model of muscular dystrophy (Seale et al., 2000). Interestingly, the closely related Pax3 was not expressed in these cells. This was an intriguing result given the widely held assumption that regeneration of adult muscle would recapitulate the same mechanisms observed in embryonic muscle development. During embryonic development, Pax3 and Pax7 share overlapping patterns of expression, and whilst it had been proposed that they might in turn share overlapping function, Pax7 cannot rescue the splotch mutant Pax3 KO phenotype, suggesting this is not the case. Therefore, it was significant that Pax7 / mutant mice showed grossly normal musculature, meaning Pax7 was dispensable for muscle development, but a reported absence of satellite cells, with accompanying failure of postnatal muscle growth and regeneration and accordingly, primary muscle cell cultures from these mice were devoid of satellite cell‐derived myoblasts. This data was interpreted as meaning that Pax7 is necessary for the specification of satellite cells and demonstrated a crucial role for Pax7 as a
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key player in the signaling pathways involved in at least one muscle stem cell niche. Interestingly, mSP cells were isolated from Pax7 / mice at a level comparable to wild‐type, suggesting again that SP and satellite cells represent distinct populations. In vitro analysis of Pax7 / mSP cells revealed that these cells were incapable of diVerentiating into muscle but showed an almost 10‐fold increase in the ability to diVerentiate into hematopoietic cells (Seale et al., 2000). This suggested that Pax7 might induce satellite cell diVerentiation in mSP cells by restricting alternate fates. Furthermore, mSP cells extracted from the Pax7 / mouse, when cultured in the presence of primary myoblasts, were induced to diVerentiate into muscle, suggesting that some portion of satellite cells is derived from muscle stem cells, though whether this was dependent upon Pax7 was not known. Building on this work and that of others (Polesskaya et al., 2003), researchers (Seale et al., 2004) used FACS analysis to extract the myogenic precursor pool present in SP cells and first observed that a proportion of wild‐type CD45þ:Sca1þ cells from regenerating muscle upregulated Pax7. In addition, unlike cells isolated from heterozygous animals, in the Pax7 / mouse CD45þ:Sca1þ cells from regenerating muscle were never observed to give rise to MyoD‐ or Desmin‐expressing cells, seemingly supporting a central role for Pax7 in myogenic specification of CD45þ:Sca1þ stem cells. Furthermore, the authors then took the same cells from uninjured wild‐type muscle, infected them with Pax7 retrovirus, and observed expression of Myf5 and MyoD, indicating that these cells were undergoing myogenesis, and following addition of low‐serum media were able to fully diVerentiate, expressing such markers of terminal diVerentiation as MHC. Finally, and most strikingly, when CD45þ:Sca1þ cells infected with Pax7 were injected into the tibialis anterior muscle of the mdx mouse, they were observed to integrate and diVerentiate in vivo, thus eVecting muscle repair by forming dystrophin‐expressing myofibers in a dystrophin‐null muscle. The satellite cell compartment of muscle stem cells is responsible for the vast majority of muscle repair and appears to be controlled by Pax7; in addition, the HSCs described previously as partaking in muscle repair are observed to upregulate Pax7 in regenerating muscle, and the same cells taken from nonregenerating muscle and exposed to exogenous Pax7 become myogenic. Taken together, this data strongly supported the idea of Pax7 as a common denominator in the specification of muscle stem cells. More recently, it has been suggested that Pax7 / mice possess satellite cells, albeit at a greatly reduced number compared to wild‐type animals, at least in the immediate postnatal period and into adulthood (Oustanina et al., 2004). These data describing satellite cells in adult Pax7 mutant muscle have not been supported in our laboratory; nonetheless, the presence of satellite cells as identified via electron microscopy, at least immediately postnatally,
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does raise the possibility that Pax7 does not directly specify the ontogeny of satellite cells. Two studies essentially propose an alternative role that may reconcile all the data currently available regarding Pax7 and muscle stem cells. Researchers (Zammit et al., 2004) neatly showed that satellite cells are capable of adopting divergent fates. Of greatest significance was the demonstration that after synchronous activation and coexpression of Pax7 and MyoD, most satellite cells proliferated, downregulated Pax7, and proceeded onto diVerentiation; however, a proportion maintained their expression of Pax7, downregulated MyoD, and withdrew from diVerentiation. Subsequently a proportion of the cells from these clusters withdrew from the cell cycle, in eVect returning to a state of quiescence and replenishing the satellite cell compartment. This interpretation was further supported (Olguin and Olwin, 2004) by work that also detected heterogeneity within individual clones of activated satellite cells, whereby cells that upregulated Pax7 were observed to escape diVerentiation and exit the cell cycle. Furthermore, the overexpression of Pax7 was observed to downregulate MyoD and block myogenic conversion of the fibroblastic cell line 10T1/2. The obvious interpretation in summarizing this data is that Pax7 is actually responsible not for the specification of satellite cells, but is vital for their self‐renewal or other aspects of their function. However, both questions remain open to debate. Some studies (Olguin and Olwin, 2004; Zammit et al., 2004) may hint at a generalized mechanism for muscle stem cells applicable to both satellite cells and HSCs. That is, both satellite cells and HSCs would require the presence of Pax7 to propagate and maintain a steady‐state cell population able to aVect muscle repair. This would explain the absence of alternative repair mechanisms in the Pax7 / mouse. There is a significant increase in CD45þ:Sca1þ cells in response to muscle damage; as stated earlier, up to 10% of CD45þ:Sca1þ cells isolated from injured muscle upregulate Pax7. The CD45þ:Sca1þ cell lineage is somewhat heterogeneous; hence, the low percentage of these cells that do upregulate Pax7 may do so in order to first replenish/maintain the muscle stem cell compartment before the repair process begins. Alternatively, if Pax7 is not required for the specification of satellite cells per se, induction of damage in Pax7 mutant muscle may result in the generation of new satellite cells that again become depleted because they cannot undergo appropriate self‐renewal and myogenic commitment in the absence of Pax7. A final possible interpretation is that Pax7 acts directly as a survival signal, allowing certain cell populations (satellite cells and HSC) to survive that in the absence of Pax7 would otherwise initiate an apoptotic cascade or follow an alternate lineage. Cells that are maintained alive in this population would then be present to respond to cues, which lead them to respond to muscle damage.
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C. How Do CD45 /Sca1 Cells Become Directed Toward a Muscle Stem Cell Fate? UndiVerentiated adult stem cells require environmental cues to initiate commitment to a specific cell type. It is reported that CD45þ/Sca1þ cells express no intrinsic commitment to the myogenic lineage when extracted from resting muscle tissue. However, when these cells were extracted at various time points following damage, they readily expressed myogenic markers suggestive of their commitment to the myogenic lineage. These findings suggest that the CD45þ/Sca1þ population requires a signaling cascade, cytokine, or growth factor in the events following muscle damage for the commitment of these cells to the myogenic lineage. Logical candidates for the commitment of CD45þ/Sca1þ cells to muscle include embryonic myogenesis pathways. In the developing embryo, sonic hedgehog (Shh), secreted by the notochord, and various Wnt family proteins, secreted by the dorsal neural tube, are required for the induction of embryonic myogenesis (Munsterberg et al., 1995; Tajbakhsh et al., 1998). In addition, adult myogenesis is thought to recapitulate the process of embryonic myogenesis (Parker et al., 2003). As discussed previously, proliferating and diVerentiating myogenic progenitor cells express myogenic regulatory factors expressed during embryonic muscle development (MyoD, Myf5, etc.), and newly formed myotubes are known to express embryonic myofibrillar genes, such as embryonic myosin heavy chain (Cornelison and Wold, 1997; Grounds et al., 1992). Here are provided several lines of evidence implicating Wnt signaling as playing an analogous pathway in adult muscle regeneration, as to that of embryonic myogenesis: (1) The addition of lithium, a Wnt‐signaling agonist, immediately induced muscle specification of CD45þ/Sca1þ cells in vitro; (2) coculture of CD45þ/Sca1þ cells with cells ectopically expressing Wnt proteins induced the expression of Pax7 and myogenic commitment; (3) regenerating muscle treated with sFRP, a known Wnt‐pathway antagonist, resulted in a significant reduction in the number of proliferating CD45þ/Sca1þ cells; (4) sFRPs were upregulated late in muscle regeneration, suggesting that sFRPs inhibit Wnt‐dependent recruitment of myogenic progenitors when regeneration is complete; and (5) proliferating myoblasts express high levels of Wnt proteins, suggesting that myoblasts may ultimately determine the fate of CD45þ/Sca1þ cells through a Wnt‐mediated mechanism. Taken together, these in vivo and in vitro results suggest that Wnt‐signaling mediates the induction of CD45þ adult stem cells to undergo myogenic specification. In contrast to these findings, another study has reported that, using an in vivo experimental design, none of the Wnt‐related proteins were upregulated
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during muscle regeneration, and in fact several sFRP family proteins were significantly upregulated suggesting that Wnt signaling was inhibited (Zhao and HoVman, 2004). Real‐time polymerase chain reaction (PCR) analysis was performed and confirmed initial observations that several Wnts are induced to significant levels during the first 5 days of muscle regeneration at the time when the myoblasts are present in greatest numbers. Therefore, Wnts are significantly upregulated during muscle regeneration, consistent with the postulated role for Wnt signaling during regenerative myogenesis (Polesskaya et al., 2003).
IV. Interpretation of Findings from Models of Extreme Muscle Damage Given that the contributions of BMDCs to the repair of muscle could be fairly described as low (at best 5%), it is understandable that researchers question the validity of cells other than those of the satellite cell lineage to repair muscle. An obvious question may be, simply because these cells can contribute to muscle repair under experimental conditions, does this mean they do so under ‘‘normal’’ circumstances? Furthermore, can these events be considered meaningful, or given the millions of cellular events occurring each second in a living organism, do they simply represent biological noise or experimental phenomenon? Indeed a study appears to confirm previously mentioned observations, that muscle repair is almost entirely attributable to satellite cells, but that a small population of BMDCs can attain myogenic qualities if presented with a regenerative environment (Sherwood et al., 2004). It is worth reiterating that the reason for the significant focus on BMDCs is that they still retain the allure of a potential therapeutic tool. They represent an accessible and seemingly inexhaustible supply of cells that have been shown to contribute (however small) to muscle repair. Therefore, the potential remains that further analysis and dissection of those mechanisms that are responsible for directing these cells toward a muscle stem cell fate will furnish researchers with the ability to commit more of these cells to this function in pathological conditions. Furthermore, satellite cells are almost obliterated by cardiotoxin, yet 4 days postinjury, there are an immense number of myogenic progenitors that far outstrip the potential of any remaining satellite cells and it is noteworthy that the study did not examine this time point (Sherwood et al., 2004). Additionally, 21 days later muscle repair is complete—taken together these data do suggest that there exists a significant, non‐satellite cell‐derived mechanism for muscle repair. As such, two questions must be addressed.
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A. Is the Regenerated Muscle Functional? To the best of our knowledge, no one has extensively examined how complete the muscle repair process is with regard to the physiological properties of the muscle. Whereas some authors have demonstrated that cardiotoxin‐ injured muscle can go through multiple rounds of injury and regeneration, this would only indicate that the repair mechanism is maintained. The electrophysiological properties of these muscles, such as the twitch and torque properties, have not been established. This could be of some significance, given that exercise‐induced damage results in significant hypertrophy and adaptation, resulting in stronger muscles. It would be of interest to know if these same eVects were present in the cardiotoxin‐injured muscle or if the repair process results in a grossly structurally sound muscle, but one that alters in its physiological properties as compared to the uninjured leg.
B. Would This Response Ever Occur Physiologically? The devastating eVect of cardiotoxin and other similarly extreme models of damage on muscle architecture could never be emulated by either disease or physiological condition. Therefore, is the response observed purely an experimental phenomenon? One simple but direct way to approach this question is to test the contributions of these cell types in physiological models of damage, such as that seen in exercise (Fig. 3). Some believe that such experiments would considerably reduce the confounding presence of the multiple other cell types and nonspecific eVects caused by such a traumatic injury that result in considerable diYculty of interpretation. For example, any analysis of gene upregulation/ downregulation from cardiotoxin‐injured muscle is rendered problematic given that there is little or no consensus as to which ‘‘housekeeping’’ genes should even be used to normalize against, given the wild fluctuation observed both within and between studies. In a more physiological model these variables are more controlled and in the absence of such noise allow the researcher to examine the specific eVects of muscle damage and its repair mechanisms. Finally, it is always tempting to make arguments from a design perspective, suggesting that because a mechanism does not fit the most desirable model, it is therefore unlikely. The possibility still remains that all the models and mechanisms discussed are valid in that they all are a directed response to muscle damage and as such all represent muscle stem cells. They may represent only an extreme response situation (such as the CD45þ:Sca1þ influx of cells in response to cardiotoxin), which ordinarily is never activated, or they may represent evolutionary conserved mechanisms of repair that
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Figure 3 Cardiotoxin versus exercise‐induced damage. (A) The extreme nature of the damage caused by cardiotoxin can be easily observed by H&E staining, with an almost complete loss of muscle fibers in the aVected area at 4 days postinjection. This can be quantified via myoblast preps using the Myf5Lacz mouse. Contrast with the damage caused via exercise observed in B. The EM images show a clear disruption of Z‐bands, indicating damage, which is also clear from the H&E.
under normal modern day conditions would be unlikely to ever play a significant role. Nevertheless, the fact that multiple cell types appear capable of fulfilling the role of a muscle stem cell should remain a cause for optimism and a continued driving force in the search for mechanisms that direct their action and would allow manipulating their potential.
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Seale, P., Ishibashi, J., Scime, A., and Rudnicki, M. A. (2004). Pax7 is necessary and suYcient for the myogenic specification of CD45(þ):Sca1(þ) stem cells from injured muscle. PLoS. Biol. 2, E130. Seale, P., Sabourin, L. A., Girgis‐Gabardo, A., Mansouri, A., Gruss, P., and Rudnicki, M. A. (2000). Pax7 is required for the specification of myogenic satellite cells. Cell 102, 777–786. Sherwood, R. I., Christensen, J. L., Conboy, I. M., Conboy, M. J., Rando, T. A., Weissman, I. L., and Wagers, A. J. (2004). Isolation of adult mouse myogenic progenitors: Functional heterogeneity of cells within and engrafting skeletal muscle. Cell 119, 543–554. Tajbakhsh, S., Borello, U., Vivarelli, E., Kelly, R., PapkoV, J., Duprez, D., Buckingham, M., and Cossu, G. (1998). DiVerential activation of Myf5 and MyoD by diVerent Wnts in explants of mouse paraxial mesoderm and the later activation of myogenesis in the absence of Myf5. Development 125, 4155–4162. Tamaki, T., Akatsuka, A., Ando, K., Nakamura, Y., Matsuzawa, H., Hotta, T., Roy, R. R., and Edgerton, V. R. (2002). Identification of myogenic‐endothelial progenitor cells in the interstitial spaces of skeletal muscle. J. Cell Biol. 157, 571–577. Torrente, Y., Tremblay, J. P., Pisati, F., Belicchi, M., Rossi, B., Sironi, M., Fortunato, F., El Fahime, M., D’Angelo, M. G., Caron, N. J., Constantin, G., Paulin, D., Scarlato, G., and Bresolin, N. (2001). Intraarterial injection of muscle‐derived CD34(þ)Sca‐1(þ) stem cells restores dystrophin in mdx mice. J. Cell Biol. 152, 335–348. Wakeford, S., Watt, D. J., and Partridge, T. A. (1991). X‐irradiation improves mdx mouse muscle as a model of myofiber loss in DMD. Muscle Nerve 14, 42–50. Ye, M., Iwasaki, H., Laiosa, C. V., Stadtfeld, M., Xie, H., Heck, S., Clausen, B., Akashi, K., and Graf, T. (2003). Hematopoietic stem cells expressing the myeloid lysozyme gene retain long‐term, multilineage repopulation potential. Immunity 19, 689–699. Zammit, P. S., Golding, J. P., Nagata, Y., Hudon, V., Partridge, T. A., and Beauchamp, J. R. (2004). Muscle satellite cells adopt divergent fates: A mechanism for self‐renewal? J. Cell Biol. 166, 347–357. Zhao, P., and HoVman, E. P. (2004). Embryonic myogenesis pathways in muscle regeneration. Dev. Dyn. 229, 380–392.
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Gene Regulation in Spermatogenesis James A. MacLean II and Miles F. Wilkinson Department of Immunology, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030
I. Introduction II. Hormonal Regulation of Transcription in the Testis A. Mechanisms of Androgen Action B. The Androgen Receptor C. Estrogen and the Testis D. Perspective III. Transcription Factors with Testicular Functions A. CREM and CREB B. Nonsteroid Hormone Nuclear Receptors C. Heat‐Shock Factors D. Sox E. Plzf F. Dmrt1 G. CAF1 H. Homeobox Genes I. Perspective IV. Testis‐Specific Gene Expression and DNA Methylation A. Male Germ Cell–Specific Genes B. Somatic‐Cell Testis Genes C. Tissue‐Specific Hypomethylation: Cause or Consequence of Transcription? D. Perspective Acknowledgments References
Mammalian spermatogenesis is a complex hormone‐dependent developmental program in which a myriad of events must take place to ensure that germ cells reach their proper stage of development at the proper time. Many of these events are controlled by cell type– and stage‐specific transcription factors. The regulatory mechanisms involved provide an intriguing paradigm for the field of developmental biology and may lead to the development of new contraceptives an and innovative routs to treat male infertility. In this review, we address three aspects of the genetic regulatory mechanism that drive spermatogenesis. First, we detail what is known about how steroid hormones (both androgens and estrogens) and their cognate receptors initiate and maintain mammalian spermatogenesis. Steroids act through three mechanistic routes: (i) direct activation of genes through Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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hormone‐dependent promoter elements, (ii) secondary transcriptional responses through activation of hormone‐dependent transcription factors, and (iii) rapid, transcription‐independent (nonclassical) events induced by steroid hormones. Second, we provide a survey of transcription factors that function in mammalian spermatogenesis, including homeobox, zinc‐ finger, heat‐shock, and cAMP‐response family members. Our survey is not intended to cover all examples but to give a flavor for the gamut of biological roles conferred by transcription factors in the testis, particularly those defined in knockout mice. Third, we address how testis‐specific transcription is achieved. In particular, we cover the evidence for and against the idea that some testis‐specific genes are transcriptionally silent in somatic tissues as a result of DNA methylation. ß 2005, Elsevier Inc.
I. Introduction Mammalian spermatogenesis is a complex program of diVerentiation that presents many unique challenges. Specialized mechanisms have evolved to ensure proper control of gene expression and a smooth transition from primordial germ cells to spermatogonial stem cells to terminal spermatozoa. Understanding the mechanisms by which genes and gene products are expressed or repressed in the testes is germane to understanding normal spermatogenesis, can aid the development of strategies to treat infertility, and may provide inroads to developing new methods of male contraception. In this review, transcriptional regulatory mechanisms that govern spermatogenesis will be covered. Post‐transcriptional regulatory controls—including mechanisms regulating RNA splicing, polyadenylation, translation, and RNA stability—are also important for spermatogenesis, as discussed in other reviews (Braun, 1998; Kleene, 2003; Venables, 2002), but this topic will not be included here. In addition, this review will not cover the transcriptional networks that drive the formation of the male gonads during embryogenesis (sex determination), an area that has already been dealt with in many excellent reviews (Graves, 2002; Schepers et al., 2002; Tohonen et al., 2003). Another area of active interest that will not be discussed is the transcriptional networks controlling the development and function of male accessory organs. It is becoming apparent that the epididymis has a more important role in guiding fertility than simply storing sperm. The molecules that participate in the epididymis’ other functions, including maturation of sperm, protection from oxidation, and acquisition of sperm motility, are under active study; some candidate transcription factors that may control these genes are beginning to be identified (Lye and Hinton, 2004; Rodriguez et al., 2001; Suzuki et al., 2004). Another well‐characterized male accessory organ, the prostate, has received a great
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deal of attention due to the high incidence of malignant prostate tumors. Studies examining the transcription factors that participate in prostate gene regulation, including the extensive literature on androgen‐mediated gene regulation in the prostate, have been covered in other reviews (Heinlein and Chang, 2004; Kim and Coetzee, 2004). Section II of this review focuses on the molecular roles of steroid hormones in the initiation and maintenance of spermatogenesis. We will describe the three mechanistic routes used by steroid hormones to elicit changes in gene expression: (1) direct activation of genes with hormone receptor‐ binding sites; (2) indirect regulation of genes by hormone‐activated transcription factors; and (3) rapid activation of signal transduction events by membrane‐bound hormone receptors (Fig. 1). Most of Section II will focus
Figure 1 Three modes of androgen action. In the primary mode, testosterone (T) directly activates transcription by promoting androgen receptor (AR) dimerization, increasing AR’s aYnity for androgen response elements (ARE) in the promoter of target genes. In the secondary mode, androgen‐induced transcription factors (TF) alter the transcription of secondary response genes. (These typically lack AREs and thus cannot respond directly to androgen.) In nonclassical androgen action, rapid signal transduction events are induced in the absence of new transcription by androgen. Such rapid cell membrane‐triggered events, including Caþþ uptake and protein phosphorylation (P), sometimes require AR (pictured). In other circumstances, they occur independently of AR (not pictured). See Section II.A for further details.
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on the actions of androgens and androgen receptors (ARs; the transcription factor androgens bind and activate) as these have long been considered the key players in male fertility. However, we will also discuss molecular mechanisms of ‘‘female’’ hormones such as estrogen and their cognate receptors, as they are also essential for spermatogenesis. The section concludes by drawing attention to other issues in the field—including the need to define more primary and secondary hormone‐response genes and their downstream targets—that will need to be addressed before the hormone‐dependent regulatory pathways directing spermatogenesis will be completely understood. Section III delineates the spermatogenic role of transcription factors other than steroid receptors. In addition to individual transcription factors, several transcription factor families will be covered, including the zinc‐finger, Sox, and homeobox families (Table I). Attention will focus on transcription factors implicated in spermatogenesis based on transgenic and knockout mouse models. Phenotypic analysis of these mutant mice has identified transcription factors with roles in germ‐cell development and testicular somatic‐ cell function. The focus will be on transcription factors active in the testis after birth, with only limited discussion of any function they might have before birth. Future research directions will be outlined, including the need to use tissue‐specific knockout and knockdown approaches to address the roles of many transcription factors, particularly those whose loss results in early death or crippling deformities that prevent productive mating. Section IV concerns testis‐specific transcription. In particular, this section addresses an interesting property of many testis‐specific genes: Their promoters are rarely methylated in the testis but are highly methylated in other tissues. This has led many investigators to hypothesize that DNA methylation might dictate testis‐specific expression by selectively repressing
Table I
Transcription Factors Essential for Spermatogenesis
Gene Product
Cell Type
Reference
CREB CREM‐ TR4 Hsf1 Hsf2 Sox3 Plzf Dmrt1 CAF1 Sperm1 Pem (Rhox5)
Sertoli cells Male germ cells Male germ cells Male germ cells Sertoli and germ cells Sertoli cells Spermatogonia Sertoli and germ cells Sertoli and germ cells Male germ cells Sertoli cells
Hummler et al., 1994 Blendy et al., 1996; Nantel et al., 1996 Mu et al., 2004 Nakai et al., 2000 Wang et al., 2003 Weiss et al., 2003 Costoya et al., 2004 Raymond et al., 2000 Berthet et al., 2004 Pearse et al., 1997 MacLean et al., 2005
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Genes Selectively Expressed and Hypomethylated in Testis
Gene Product
Cell Type
Reference
Pgk2 Pdha‐2 LDH‐c ALF (TFIIA) TH2B H1t Tnp1 MAGE‐A1 Tact1 (Act17b) Sry FSHR SGP2 Rhox5 (Pem) 5‐reductase
Male germ cells Male germ cells Male germ cells Male germ cells Male germ cells Male germ cells Male germ cells Male germ cells Male germ cells Fetal Sertoli cells Sertoli cells Sertoli cells Sertoli cells Leydig cells
Zhang et al., 1998 Iannello et al., 1997 Bonny and Goldberg, 1995 Xie et al., 2002 Choi and Chae, 1991 Singal et al., 2001 Trasler et al., 1990 De Smet et al., 1996 Hisano et al., 2003 Nishino et al., 2004 Griswold and Kim, 2001 Rosemblit and Chen, 1994 S. Shanker and M.F.W., unpublished Reyes et al., 1997
transcription in nontesticular tissues. DNA methylation was proposed as a silencing mechanism more than 30 years ago and has since been shown to be important for transcription repression in some instances (including X‐chromosome inactivation, imprinting, and defense against transposable elements), but its role in tissue‐specific expression has been controversial. The possibility that several testis‐specific genes are good candidates for regulation by DNA methylation will be discussed. Many of these are male germ‐cell genes, whereas others are selectively expressed in testicular somatic cells (Table II). This review will describe their expression pattern and detail the evidence that their selective expression in the testis is regulated by DNA methylation. The various methods that have been used to examine this issue will be discussed, including their positive and negative attributes. Finally, we will discuss additional approaches that, along with methods that have already been used, may definitively identify genes whose testis‐specific transcription is controlled by DNA methylation.
II. Hormonal Regulation of Transcription in the Testis The two primary functions of the testis are spermatogenesis and the synthesis of steroid hormones under the control of gonadotropins. Spermatogenesis is the process by which germ‐cell precursors mature into functional spermatids. The local secretion of androgens by Leydig cells ensures that seminiferous tubules are exposed to suYcient levels of androgen to drive the expression of spermatogenesis‐promoting genes. The crucial role of
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testosterone in regulating spermatogenesis is well known, but evidence indicates that estrogens are also important. Androgens and other steroid hormones exert their actions through a specific class of transcription factors termed nuclear hormone receptors (NRs). NRs are allosteric transcription factors that form multi‐subunit complexes with their cognate ligands (such as steroid hormones), their DNA targets, and regulatory proteins (such as coactivators and corepressors) that help direct the nature of the transcriptional response. The type 1 NRs, which form the steroid receptor subgroup, are the receptors for the sex hormones (androgens, estrogens, and progestins), as well as for glucocorticoids, vitamins, and mineralocorticoids (Pettersson and Gustafsson, 2001). In the absence of ligands, NRs shuttle between the nucleus and the cytoplasm. The progestin and glucocorticoid receptors are primarily cytoplasmic at steady state, whereas unbound estrogen receptor (ER) is predominantly nuclear (Dauvois et al., 1993). The localization of unbound AR is somewhat controversial; diVerent groups have reported the receptor as being either predominantly cytoplasmic or nuclear (Georget et al., 1997; Jenster et al., 1993; Kemppainen et al., 1992; Tyagi et al., 2000; Waller et al., 2000). AR is part of a multiprotein complex that also includes heat‐shock factors (Hsfs) that dissociate in the presence of androgen. Androgen‐dependent stripping of these factors allows AR to dimerize and undergo a conformational change, such that the receptor‐steroid complex gains increased aYnity for specific DNA regulatory elements and is then able to promote the transcription of target genes (Bagchi et al., 1992). Androgens, like other steroid hormones, exert their influence through three modes of action: 1. Primary hormone responses. These responses are the result of direct interaction of hormone‐steroid receptor complexes with cis‐regulatory DNA elements in promoters and enhancers of primary target genes. As little as 30 min of hormone treatment is suYcient to alter the transcription of primary‐ response genes (Shang et al., 2000, 2002). Transcriptional activation is usually the result, but steroids can also downregulate their target genes. 2. Secondary hormone responses. Nascent protein synthesis is required after steroid hormone stimulation to elicit some transcriptional responses. These are termed secondary hormone responses, as the target genes do not respond directly to NRs but instead are controlled by steroid‐dependent transcription factors (Dean and Sanders, 1996). Secondary‐response genes typically require hours to days to maximally respond. Secondary‐response genes typically lack binding elements for NRs and thus cannot function as primary‐response genes. 3. Nonclassical responses. In addition to eliciting primary and secondary transcriptional responses, steroid hormones can trigger rapid responses that do not require new transcription (Castoria et al., 2004; Falkenstein
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et al., 2000; Walker, 2003). These include increased levels of secondary messengers such as Caþþ and activated protein kinases. In some cases, nonclassical responses require steroid receptors, including progestin receptor (Bramley, 2003), ER (Hess et al., 1997; Kelly et al., 2002), and AR (Heinlein and Chang, 2002). In other cases, the responses appear to occur independently of NRs and instead involve as‐of‐yet unidentified alternative hormone‐binding receptors (Benten et al., 1999a,b; Leung et al., 2001; Lieberherr and Grosse, 1994; Lieberherr et al., 1993). Nonclassical responses are often called ‘‘nongenomic’’ responses, a name that accurately reflects steroid‐induced responses triggered independently of de novo transcription. However, the name nongenomic does not describe downstream transcriptional events that can occur in response to rapid steroid hormone activation of secondary messages and protein kinases (Fix et al., 2004; Kwon et al., 2000). Thus, we suggest that nonclassical is a better general term, as it describes both rapid hormone‐induced responses and the consequent downstream transcriptional events (the latter of which are diVerent than either primary or secondary responses). A fair amount is known about all three modes of steroid hormone action, including AR‐mediated events in the pituitary gland and skeletal muscle (Compston, 2001; Pelletier, 2000; Tohonen et al., 2003); AR action in sexual development and cancer (Bagchi et al., 1992; Wang and Chang, 2004); and ER‐mediated events in bone and female reproductive tissues, such as during ovulation and uterine cyclicity (Matzuk and Lamb, 2002; Zhu et al., 1999). Surprisingly, less is known about AR action in the postnatal and adult testis. Few primary and secondary androgen‐responsive genes have been identified in the testis, and we are only beginning to understand the nonclassical actions of AR in the testis. Some AR‐responsive putative transcription factors expressed in the testis have been identified, including the cAMP‐ response element (CRE) binding protein (CREB) and some members of a newly described reproductive homeobox gene subfamily on the X chromosome (De Cesare et al., 1999; MacLean et al., 2005), but their direct targets in the testis are not known. A summary of the current state of the field follows. A. Mechanisms of Androgen Action 1. Primary Androgen Response Genes Although several genes are directly regulated by AR in the prostate, including kallikrein‐2, probasin, cystatin‐related protein‐2, and prostate‐specific antigen (Devos et al., 1997; Kim and Coetzee, 2004), only a few genes have been identified that are directly regulated by AR in the testis. One of
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these is the Pem/Rhox5 homeobox gene, whose proximal promoter is expressed in an androgen‐dependent manner exclusively in Sertoli and caput epididymal cells in vivo (De Gendt et al., 2004; Lindsey and Wilkinson, 1996b; Maiti et al., 1996; Sadate‐Ngatchou et al., 2004). Transfection experiments have shown that its androgen‐dependent transcription depends on androgen‐response elements (AREs) within a 0.3‐kb region immediately upstream of the Pem/Rhox5 transcription start site (Barbulescu et al., 2001; Geserick et al., 2003). This region is suYcient to confer androgen‐ dependent expression specifically in Sertoli and epididymal cells in transgenic mice in vivo (Rao et al., 2002b, 2003). Another gene suggested to be directly responsive to androgen is the proto‐ oncogene MYC, as it is induced by testosterone in a Sertoli cell line even in the presence of the protein synthesis inhibitor cycloheximide (Lim et al., 1994). As further evidence, the MYC promoter region is bound by AR in LNCaP prostate cancer cells, as shown by chromatin immunoprecipitation experiments (Amir et al., 2003). 2. Secondary Androgen‐Response Genes Among the first secondary androgen‐response genes identified were those encoding arginase, connexin‐43, and glutathione S‐transferase (Dean and Sanders, 1996). These genes are regulated by androgen in the testis and prostate (Benbrahim‐Tallaa et al., 2002; Gotoh et al., 1994), as well as in tissues that express little or no AR, including the liver and kidney (Benbrahim‐ Tallaa et al., 2002; Fan et al., 1992; Kumar and Kalyankar, 1984; Piersanti and Lye, 1995); the latter is evidence that they are bona fide secondary‐ response genes. Further evidence is that none of the genes have obvious AREs in their promoters. The regulation of the arginase gene was re‐examined in the mouse kidney (Levillain et al., 2004). Both physiological and pharmacological doses of testosterone resulted in strong enhancement of arginase expression. Surprisingly, the response was greater in the kidneys of female mice than those of males, possibly because basal transcription of arginase (which is not aVected by androgens) is initially higher in female kidneys. In addition, more free androgen may be available in the female circulation because females have overall lower levels of sequestering androgen‐binding proteins. Arginase upregulation requires several hours of treatment with androgen, which is consistent with its being a secondary androgen response, but it is still unclear what transcription factor serves as the bridge for arginase transcriptional activation. Numerous putative response elements for NF‐B, SP1, and AP1 are present in the arginase 50 flanking sequence, but these transcription factors are more likely to have a role in basal arginase transcription
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than in androgen‐dependent upregulation, as they do not have known androgen‐dependent actions. Other possible secondary androgen‐response genes are the many other genes that are induced by androgen but lack obvious AREs in their promoters (Sadate‐Ngatchou et al., 2004). Testis‐specific examples of this category include the genes encoding carbonic anhydrases and angiotensin‐converting enzyme in Leydig cells and androgen‐binding protein and claudin‐1 in Sertoli cells (Carter et al., 2001; Douglas et al., 2004; Gye, 2003). To date, no secondary androgen‐response genes repressed by androgen in the testis have been identified. A gene of this type in the kidney is the ornithine aminotransferase gene (Levillain et al., 2004). The identities of androgen‐regulated transcription factors that regulate secondary androgen‐response genes in the testis remain obscure. One transcription factor that may have this role in Sertoli cells is the androgen‐ regulated homeodomain protein Pem/Rhox5 (Barbulescu et al., 2001; Geserick et al., 2003; Lindsey and Wilkinson, 1996b; Rao et al., 2002b, 2003). Microarray analysis has identified genes whose expression is altered in response to loss of Pem/Rhox5 in the postnatal [day 12 post partum ( p.p.)] testis (MacLean et al., 2005). Many of these genes encode proteins involved in energy metabolism, including the nuclear receptor PPAR and the transcriptional coactivator PGC1. Other genes exhibiting altered expression in response to loss of Pem/Rhox5 encode insulin II, adiponectin, and resistin, all of which are secreted proteins that regulate metabolism. Most of these gene products were not previously known to be expressed in the testis; their identification suggests the existence of complex metabolic pathways in the testis regulated by communication between diVerent cell types. Interestingly, resistin was shown to be expressed in an androgen‐ and PPAR‐dependent manner in the testis during the early stages of rat spermatogenesis (Nogueiras et al., 2004). Resistin’s stage‐specific expression pattern suggests it may be negatively regulated by Pem/Rhox5. However, whether resistin or any of the other energy metabolism genes are direct targets of Pem/Rhox5 remains to be determined. If instead Pem/Rhox5 indirectly regulates these genes (by boosting the metabolism of Sertoli cells), it will be important to identify the direct targets of Pem/Rhox5 that ultimately lead to metabolic enhancement. 3. Nonclassical Androgen Responses Although primary and secondary androgen responses contribute to spermatogenesis, they may not fully account for androgen action in the testis, as suggested by two lines of evidence. First, AR‐induced gene transcription is maximal in response to a much lower level of testosterone (1 nM) than
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required in vivo to fully support spermatogenesis (70 nM) (Walker, 2003). Second, very few testis genes have been identified that are directly responsive to AR and androgen (Barbulescu et al., 2001; Geserick et al., 2003; Lim et al., 1994; Rommerts, 1988; Veldscholte et al., 1992). This ‘‘testosterone paradox’’ brings up the possibility that nonclassical androgen responses, which often require higher levels of testosterone than classical responses, may contribute substantially to spermatogenesis. Nonclassical androgen responses can be triggered either independently of AR or in an AR‐dependent manner. Evidence for AR‐independent activation comes from studies in rat osteoblasts and human granulosa cells, both of which rapidly respond to androgen treatment by increasing their concentrations of Caþþ and other secondary messengers (Kousteni et al., 2002; Lieberherr and Grosse, 1994). It is unlikely that this Caþþ response in these cells is mediated by AR, as Caþþ levels increase within 10 seconds. Similarly, cultured eVerent ducts and epididymides rapidly respond to 5‐dihydroxytestosterone treatment by reducing their chloride secretion within 10–20 seconds (Leung et al., 2001). This response may be physiologically relevant, as chloride secretion is required for fluid reabsorption in these organs. Fibroblasts also appear to respond in an AR‐independent manner, as the proliferation of NIH‐3T3 fibroblasts is triggered by the nonhydrolyzable androgen agonist R1881 without movement of AR into the nucleus or measurable alterations in AR‐mediated transcription (Castoria et al., 2004). Perhaps the best evidence for AR‐independent androgen responses comes from studies in the IC‐21 macrophage cell line and splenic T cells, neither of which have active AR. Both respond to androgen by increasing intracellular Caþþ within seconds (Benten et al., 1999a,b). Interestingly, the increase in Caþþ in the macrophage line appears to come from intracellular stores, whereas in splenic T cells it probably derives from the extracellular milieu through non‐voltage‐gated Caþþ channels in the plasma membrane. Nonclassical responses that depend on AR have been reported to occur in Sertoli and prostate cells. The rapid androgen‐induced increase in Caþþ in both of these cell types is inhibited by the AR antagonist hydroxyflutamide (Gorczynska and Handelsman, 1995; Steinsapir et al., 1991). Further evidence that AR is required comes from studies of AR‐negative prostate and human genital skin fibroblast cell lines transfected with AR expression constructs (Peterziel et al., 1999). What signal transduction pathways are used to initiate nonclassical androgen responses? The initial studies to address this were conducted in breast and prostate cancer cells. The breast cancer cell line PMC42 responds to R1881 by rapidly activating extracellular‐regulated kinase (ERK), leading to activation of the mitogen‐activated protein (MAP) kinase pathway and
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its downstream genes (Zhu et al., 1999). 5‐Dihydroxytestosterone also rapidly and transiently (within 2–60 min) induces the phosphorylation of ERK in primary prostate cells (Peterziel et al., 1999). Similar results have been obtained with the AR‐positive prostate cell line LNCaP (Lyng et al., 2000). In the case of the AR‐negative human prostate cell line PC3, ERK activation is observed only when an AR‐expression plasmid is cotransfected, indicating that activation of the MAP kinase pathway is an AR‐dependent response (Lyng et al., 2000). Other studies have shown that androgens rapidly induce many other factors upstream of the MAP kinase pathway, including protein kinase A, protein kinase C, calmodulin, phospholipase C, and guanine nucleotide exchange factors (Finkbeiner and Greenberg, 1996; Peterziel et al., 1999). Sertoli cells respond to androgen by rapidly elevating Caþþ levels as a result of Caþþ influx through L‐type channels (Gorczynska and Handelsman, 1995; Lyng et al., 2000). This response appears to be relatively specific to androgens, as 17‐estradiol, progesterone, and other steroids have a weaker or no eVect (Gorczynska and Handelsman, 1995). Sertoli cells also respond to androgens by activating the MAP kinase pathway, as R1881 rapidly induces the phosphorylation of ERK1 and ERK2 (Fix et al., 2004). Interestingly, R1881 also rapidly induces the phosphorylation of the transcription factor CREB. Measurable increases in both ERK and CREB phosphorylation are observed within 1 min of androgen stimulation, and this hyperphosphorylated state is maintained for at least 12 hr. Several lines of evidence suggest that this nonclassical pathway depends on AR, including the finding that CREB hyperphosphorylation is largely prevented when AR is depleted, knocked out, or inhibited. CREB phosphorylation has been implicated in the upregulation of several Sertoli‐cell genes that may contribute to germ‐cell proliferation and development (Scobey et al., 2001). Two putative targets of rapidly activated CREB are the lactate dehydrogenase A and early growth response‐1 genes, both of which contain consensus CREB‐binding sites, lack AREs, and are rapidly induced (within 6 hr) by androgen stimulation following the same kinetics as CREB (Fix et al., 2004) and the previously published CREB‐ responsive genes proenkephalin and c‐Fos (Sassone‐Corsi et al., 1988). There are many other potential targets of CREB‐mediated signaling, including the genes encoding transferrin, IGF‐1, AR, NF‐B, Elk‐1, c‐Jun, C/EBP, and some members of the STAT transcription factor family. Many of these genes have promoters with putative CREB‐binding sites and are upregulated when MAP kinase is activated in response to hormones and growth factors (Fix et al., 2004; Kwon et al., 2000; Walker, 2003). CREB also binds and activates its own promoter, suggesting that its expression is maintained by a positive feedback loop (Walker et al., 1995).
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B. The Androgen Receptor 1. AR Domains The AR gene is located on the X chromosome (q11–12 in humans, C3 in mice) and encodes a multidomain receptor that responds to ligand binding by promoting the transcription of androgen‐responsive genes (see Section II.A). AR shares the same modular structure found in other NRs: a DNA‐binding domain in the middle is flanked by a ligand‐binding domain at the C‐terminus and an activation domain at the N‐terminus. Remarkably, more than 70 protein cofactors have been identified that interact with specific domains of AR (Gottlieb et al., 2004). Some of these cofactors collaborate with AR to activate transcription of target genes, while others lead to transcriptional repression. The binding sites for both the positive and negative regulating factors are distributed evenly across AR’s three domains, with a few factors interacting with multiple domains. This, along with the even distribution of gain‐of‐function and loss‐of‐function mutations (more than 600 characterized in humans) along the entire length of the AR gene, suggests that AR has evolved so that virtually none of its sequences are dispensable (Gottlieb et al., 2004). AR’s transactivating domain contains glutamine‐rich and a glycine‐rich motifs important for fine‐tuning AR’s activity (Gao et al., 1996; Sleddens et al., 1992). Both of these motifs have evolved into docking sites for cofactors that are crucial for normal AR function (Gottlieb et al., 2004). Expanded lengths of the trinucleotide repeats encoding these motifs have been linked to defects in fertility, neuromuscular development, and cancer, including testicular tumors (Giwercman et al., 2004; Gottlieb et al., 2004; Yeap et al., 2004). The polyglutamine tract may represent an emerging paradigm for testicular transcription‐factor complexes, as it is also found in at least two testis‐ expressed factors besides AR. One of these is the germ cell–specific transcription factor CREM (CRE modulator protein; detailed in Section III), which contains a polyglutamine repeat in the activation domain that interacts with transcriptional cofactors (Radhakrishnan et al., 1997). The other is Rhox8 (also called Tox), an X‐linked homeobox gene encoding a putative transcription factor containing two large polyglutamine stretches (Kang et al., 2004; MacLean et al., 2005). The Rhox8 gene is developmentally regulated in the testis; it is expressed at least as early as day 7 p.p., is maximally expressed at day 12 p.p., and its expression persists in the adult testes (MacLean et al., 2005). Real‐time RT‐PCR analysis of adult testicular cell subsets indicated that Rhox8 is expressed primarily in Sertoli cells (MacLean et al., 2005). Rhox8 has also been reported to be expressed in mature germ cells, based on in situ hybridization analysis (Kang et al., 2004), which is surprising given that the X chromosome is transcriptionally inactivated in postmeiotic
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germ cells. Interestingly, the exon encoding the Rhox8 polyglutamine region is skipped in a proportion of Rhox8 transcripts in the testis (J. A. M. and M. F. W., unpublished observations). Further studies are required to determine the functional consequences of this alternative splicing event and whether it is developmentally regulated. 2. Role of AR in Spermatogenesis Disruption of AR by a variety of approaches (surgical, chemical, and genetic) prevents the generation of sperm, feminizes the external genitalia, and prevents complete development of the male accessory glands (Collins et al., 2003; Roberts and Zirkin, 1991; Sharpe and Skakkebaek, 1993; Yeh et al., 2002). That AR is absolutely required for spermatogenesis is further supported by the finding that testosterone alone, in the absence of follicle‐ stimulating hormone (FSH) or luteinizing hormone (LH), is suYcient to support spermatogenesis (Cunningham and Huckins, 1979). While the role of AR in spermatogenesis is undisputed, it has not been clear which specific cell type(s) in the testes require AR signaling to drive spermatogenesis. A popular candidate is the Sertoli cell, as it expresses high levels of AR in a stage‐dependent manner and is in intimate contact with germ cells at all phases of their development. To begin to address this issue, three groups have generated conditional knockout mice that selectively ablate AR in Sertoli cells (Chang et al., 2004; De Gendt et al., 2004; Holdcraft and Braun, 2004). As described in the following, these studies clearly show that AR functions in Sertoli cells to direct spermatogenesis by a mechanism independent of AR’s other function of promoting testicular descent. One of these groups also generated mice with a hypomorphic (subfunctional) allele of AR in all tissues (also described later) that possesses a unique phenotype with interesting implications (Holdcraft and Braun, 2004). All three groups knocked out AR in Sertoli cells using the Cre recombinase/loxP system. They generated mice possessing a floxed Ar gene (containing loxP sites) and a Cre transgene driven by the anti‐Mu¨ llerian hormone (Mu¨ llerian‐inhibiting substance) promoter, which is expressed in fetal and prepubertal Sertoli cells. Expression of Cre causes deletion of the DNA between the loxP sites, thereby generating a disrupted Ar gene only in Sertoli cells. All three groups found that the completion of spermatogenesis was blocked in these mutant mice, as evidenced by the virtual absence of mature spermatids in the testis and spermatozoa in the epididymis (Chang et al., 2004; De Gendt et al., 2004; Holdcraft and Braun, 2004). All three groups also found that the mutant testes were smaller and contained fewer germ cells, which more frequently underwent apoptosis. These results strongly suggest that AR expression in Sertoli cells is necessary for these
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nurse cells to promote the diVerentiation and/or survival of the adjacent germ cells. Spermatogenesis was blocked despite the fact that the mice exhibited normal testicular descent, indicating that the spermatogenesis blockade was not caused by cryptorchidism. This was consistent with previous results from other groups, all of which found that spermatogenesis could be inhibited in animals with descended testes by drastically lowering intratesticular testosterone levels—for example, by ablation of Leydig cells in rats using ethane dimethane sulfonate (Sharpe et al., 1990). The AR‐mutant mice also exhibited relatively normal development of the male genital tract (although one group reported a twofold reduction in the size of the epididymis [De Gendt et al., 2004]), indicating that AR expression in perinatal and postnatal Sertoli cells is not essential for the development of other organs in the urogenital tract. The data from the three groups do not agree on the specific stages of germ cells aVected by loss of AR expression in Sertoli cells. Two groups found that loss of AR expression in Sertoli cells specifically aVects germ cells progressing through meiosis (early spermatocytes) (Chang et al., 2004; De Gendt et al., 2004), whereas the third group found that the defect occurs at a later stage, during the transition of spermatids from round to elongated (Holdcraft and Braun, 2004). The first group (Chang et al., 2004) found that spermatocytes were arrested at the diplotene stage, based on flow cytometric DNA analysis, visual inspection of cell subsets in the seminiferous epithelium, and reduced expression of germ‐cell stage‐specific markers (Cap3, cyclin A1, and sperm1). The second group (De Gendt et al., 2004) used diVerent methods to come to a similar conclusion. They found that the mutant mice had fewer spermatocytes than normal (twofold reduction) and almost no round spermatids. They observed a progressive loss of pachytene spermatocytes between stages VI and XII, suggesting a defect in progression through this stage of meiosis (because of either a meiotic blockade or an inability of pachytene spermatocytes to survive). Both studies indicate that AR expression is not necessary for germ‐cell proliferation or the initiation of meiosis (the preleptotene stage and the beginning of the pachytene stage). Instead, AR expression in Sertoli cells is necessary for germ cells to fully progress through the initial stages of meiosis. What is still uncertain, based on diVerent results from the two groups, is whether AR expression is necessary to complete the pachytene stage or to progress to the diplotene stage. However, because the two groups used diVerent approaches to examine the stages of spermatogenesis, their diVerences may be more apparent than real. Also, the eVects on germ‐cell development were not absolute. For example, some round spermatids were present in the mutant mice studied (De Gendt et al., 2004).
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In contrast to the other two studies, a third group (Holdcraft and Braun, 2004) found that ablation of AR expression in Sertoli cells did not prevent germ cells from going through meiosis. Instead, they found that the postmeiotic germ cells formed round spermatids but were unable to progress to the elongating spermatid stage, indicating that this is an AR‐sensitive step. This observation suggests that AR‐positive Sertoli cells are required for either the diVerentiation or the survival of late‐round spermatids. Either explanation is also supported by testosterone withdrawal and replacement studies performed years earlier (O’Donnell et al., 1994, 1996). The mouse phenotype obtained may have been markedly diVerent than the one obtained by the other two groups because of diVerences in the mutated Ar allele created after Cre‐mediated deletion (Holdcraft and Braun, 2004). The floxed Ar gene in the mice made by Holdcraft and Braun loses exon 1, whereas the gene in the mice made by the other two groups loses exon 2. Perhaps the loss of exon 1 does not completely disrupt AR function, allowing germ cells to progress through the meiotic block observed by the other two groups. The study also used diVerent anti‐Mu¨ llerian hormone/Cre mice than the other groups, which leads to the possibility that Cre expression was lower in their mice and AR expression was therefore less eYciently ablated (Holdcraft and Braun, 2004). Another possible explanation is that the floxed Ar gene in the mice created by this group is a hypomorphic allele in the absence of Cre. There is a weak splice acceptor in the neocassette that diverts some splicing to produce a cryptically spliced mRNA that does not encode normal AR. This hypomorphic allele reduces (but does not completely ablate) the level of AR in both Sertoli and non‐Sertoli cells (such as Leydig cells, myoid cells, and nontesticular AR‐positive cells). This reduction in AR might somehow allow germ cells to progress through meiosis, perhaps because of the massive increase in serum testosterone levels caused by the hypomorphic allele (Holdcraft and Braun, 2004). Regardless of the exact mechanism, it is likely that Sertoli‐cell AR‐ mediated signaling is less dramatically aVected in the mice in this study (Holdcraft and Braun, 2004), as expression of the androgen‐responsive Sertoli cell–specific gene Pem/Rhox5 was less reduced in those mice (20% of the control) than in the mice from the second group (De Gendt et al., 2004) (<1% of the control). The simplest interpretation of the combined results of these three studies is that AR is required for two distinct steps in spermatogenesis: (1) the progression of male germ cells through the initial stages of meiosis (which requires only minimal AR signaling); and (2) the progression of late‐round spermatids to elongating spermatids (which requires high levels of AR signaling). A third step that appears to require AR signaling was revealed by examination of the mutant mice that contain the hypomorphic Ar allele but do not express Cre (Holdcraft and Braun, 2004). These hypomorphic
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Ar mice, which have reduced (but not completely extinguished) Ar function in all AR‐positive cells, have a selective defect in the terminal diVerentiation of spermatids and their release from the seminiferous epithelium. They have less elongating spermatids in the seminiferous epithelium, more degenerating elongating spermatids (which are rarely seen in wild‐type controls), and reduced levels of Prm‐1, a marker of mature germ cells. Because these mutant mice still expressed some AR (as judged by immunohistochemistry) and had only a modest defect in AR signaling (as judged by a twofold reduction in Pem/Rhox5 mRNA), this suggests that the final maturation of spermatids is acutely sensitive to the loss of AR signaling. The identity of the cell type that requires AR signaling for this late germ‐cell maturation step remains to be determined. 3. Role of AR in Regulating Testosterone Production In addition to spermatogenesis, AR regulates other testicular events. An age‐old theory is that AR regulates the production of its own ligand, testosterone. This is supported by the finding that although androgen is not required for fetal Leydig cells to develop, it is required for the generation of adult Leydig cells fully competent to produce androgens (Quigley et al., 1995). This may be the result of an autoregulatory loop, as Leydig cells express AR. However, it is also possible that other AR‐positive cells, such as Sertoli cells, mediate this regulation. This latter hypothesis was addressed by the three groups that generated Sertoli cell–specific Ar‐knockout mice. Remarkably, each group obtained a diVerent result. One group (De Gendt et al., 2004) found that ablation of AR in Sertoli cells did not have a significant eVect on either serum testosterone or LH levels. In contrast, another group (Chang et al., 2004) found that mice with ablated Sertoli‐cell Ar had decreased levels of testosterone and increased levels of LH. They attributed the decrease in testosterone to increased production of anti‐Mu¨ llerian hormone, which is a known inhibitor of androgen production. The last group (Holdcraft and Braun, 2004) obtained the opposite result; they found that testosterone levels were dramatically increased (40‐fold) in their mice containing the hypomorphic Ar allele (either with or without an intact Ar gene in Sertoli cells). This apparent increase in testosterone production was matched by an almost equally dramatic increase in serum LH levels (25‐fold). The divergent results obtained by these three groups suggest that the regulation of testosterone production in response to AR is quite complex, involving intricate and sophisticated feedback signaling along the hypothalamic‐pituitary‐gonadal axis, and is aVected in diVerent ways when AR signaling is perturbed to diVerent degrees in diVerent cell types. Sertoli cells clearly regulate steroidogenesis in Leydig cells, but how they achieve this under diVerent circumstances is unclear.
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C. Estrogen and the Testis 1. Requirement of Estrogen Signaling for Male Fertility Disruption of estrogen signaling, either through disruption of estrogen synthesis or knockout of its cognate receptor, has adverse eVects on male fertility, indicating that ER signaling is important for some aspect of testis physiology (reviewed in O’Donnell et al., 2001). This goes against the conventional wisdom that estrogen is a ‘‘female’’ hormone but is now generally accepted. The initial evidence for a role of estrogens in testis function came from the discovery that male ER knockout (ERKO) mice are infertile (Hess et al., 1997; Honda et al., 1998; Lubahn et al., 1993; Robertson et al., 1999). The testes of these mutant mice appear histologically normal until puberty, at which time they begin to degenerate and become atrophic. Very few mature sperm are produced in ERKO testes, and those that are made exhibit motility defects and are not competent to undergo the acrosome reaction (Eddy et al., 1996). The primary cause of ERKO testis pathology has been attributed to abnormalities in fluid movement through the rete testes and eVerent ducts, as artificial ligation of testis tubules in normal mice recapitulates some ERKO testis defects (Hess et al., 1997). Additional support for the importance of estrogen in male fertility came from the discovery that male mice lacking aromatase, the enzyme responsible for conversion of androgens to estrogens (Simpson et al., 1994), also have reproductive defects. Aromatase‐deficient mice (independently generated by three diVerent groups) all had normal fertility through 14 weeks of age, but when older exhibited a pronounced loss of germ cells and alterations in reproductive behavior (Honda et al., 1998; Robertson et al., 1999; Toda et al., 2001). There was some variability in the timing and manifestation of these defects, which has been attributed to diVerences in exposure to maternal estrogens (in utero or during lactation), dietary estrogens (phytoestrogens can abrogate or delay the onset), and uncharacterized ligand‐independent ER actions. Interestingly, unlike ERKO mice, aromatase‐deficient mice do not exhibit defects in eVerent ductules, perhaps because maternal estrogen exposure allows normal development of the eVerent ductules. The testes of aromatase‐deficient mice have spermatids with poor acrosome development and that tend to undergo apoptosis (O’Donnell et al., 2001). This suggests that estrogen signaling is crucial for the survival and/or diVerentiation of round spermatids. Estrogen‐dependent germ‐cell survival may be through a nonclassical mechanism not involving ER, as low concentrations of estradiol (much less than testosterone) are suYcient to reduce apoptosis of meiotic spermatocytes and elongating spermatids in cultured seminiferous epithelium (Pentikainen et al., 2000).
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2. Testicular Cell Types Synthesizing and Responding to Estrogen What cell types in the testis synthesize estrogen? Leydig cells express relatively high levels of aromatase and thus are capable of generating estrogen. Aromatase is also expressed by germ cells from the pachytene stage to the elongated‐spermatid stage, and in some species, aromatase is expressed in ejaculated sperm (Carreau et al., 1999; Robertson et al., 1999). There is considerable interest in the latter, as it has been reported that there is a strong correlation between seminal estrogen levels and fertility in humans (Bujan et al., 1993). The ability of estradiol injections to increase sperm count in patients with congenital defects in the aromatase gene is being investigated (Herrmann et al., 2002). With regard to estrogen responsiveness, the consensus is that ER is predominantly localized to eVerent duct cells in many mammalian species and Leydig cells in some nonprimate mammals (Nie et al., 2002; O’Donnell et al., 2001; Zhou et al., 2002). The second ER receptor, ER, is also expressed in the testis (O’Donnell et al., 2001; Saunders et al., 2001; Zhou et al., 2002). Interestingly, ER is expressed by germ cells at many stages of development, suggesting that ER could have direct eVects on many aspects of germ‐cell function. Because germ cells also express aromatase, it is possible that germ cells respond in an autocrine fashion to estrogens to control their survival and capacitation (Carreau et al., 2002). Consistent with this, evidence suggests that germ cells respond to ER‐mediated signaling (Saunders et al., 1997). However, despite its ability to signal and its abundant expression in the male reproductive tract, the in vivo role of ER remains unclear, as targeted deletion of ER in mice does not cause obvious abnormalities in testicular physiology or epididymal function (Krege et al., 1998). And while targeted disruption of both ER and ER results in more severe defects in ovarian follicle development, the male reproductive defects appear to be no worse than when only ER is knocked out (Dupont et al., 2000; Krege et al., 1998). 3. Estrogen‐Responsive Genes in the Testis Like testosterone, estrogen stimulates cells at three levels; it induces primary hormone responses by directly activating genes, it induces secondary hormone responses by indirectly regulating genes, and it induces nonclassical responses by triggering rapid events that do not require new gene expression. We will briefly cover all three modes of action, with an emphasis on eVects in postnatal and adult testes. The testis has the necessary cofactors to directly activate genes in response to estrogen signaling, based on the demonstration that a luciferase construct under the control of a promoter containing three tandem ERE elements is highly active in testes of transgenic mice (Lemmen et al., 2004). One putative
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direct target of estrogen signaling in the testis is the gene Gpx4. In mice and rats, Gpx4 is upregulated in testis upon treatment with estradiol and downregulated by treatment with the estrogen antagonist tamoxifen (Nam et al., 2003). While not a proven estrogen primary‐response gene, Gpx4 is a reasonable candidate, as it contains estrogen‐responsive elements (EREs) in its promoter. To clarify its role, however, experiments must be done with ERE‐defective mutants of the Gpx4 promoter. A study identified seven putative secondary estrogen‐response genes whose expression in the testis vary in response to the estrogenic compound diethylstilbestrol (Matsuno et al., 2004). Among the upregulated genes were those encoding axonemal dynein heavy chain (whose loss is associated with motility defects), adaptin 1, a protein kinase C‐binding protein, and an aurora‐related kinase linked to formation of the blood‐testis barrier. All seven of these genes are likely to be bona fide estrogen secondary‐response genes, as none had obvious EREs in their promoters and all required several hours of diethylstilbestrol exposure for regulated expression. ER and ER have been shown to trigger nonclassical responses through secondary messenger signaling in transfected cells in vitro and in several tissues in vivo, including brain, liver, kidney, uterus, and ovary (reviewed in Sak and Everaus, 2004). To our knowledge, the only study reporting nonclassical ER signaling in the male mammalian reproductive tract examined human spermatozoa, which possess membrane‐bound receptors that can only function nongenomically, as sperm chromatin is transcriptionally incompetent (Luconi et al., 1999). This study found that 17‐estradiol rapidly induced calcium influx and tyrosine phosphorylation in human spermatozoa. Because the female reproductive tract has high levels of estradiol, this suggests that the ER signaling response could have profound consequences on capacitation and/or fertilization. Other reported eVects of 17‐estradiol on sperm include enhanced motility, improved oocyte penetration, and alterations in metabolism that increase sperm longevity (Idaomar et al., 1989). Another example of the nonclassical eVects of estrogen comes from some species of fish. Atlantic croaker fish respond to estrogen by reducing androgen synthesis; this eVect is not blocked by transcription and translation inhibitors, suggesting it is a nonclassical action of estrogen (Loomis and Thomas, 2000). This has potential commercial consequences, as environmental xenoestrogens and phytoestrogens can inhibit the fertility of male spawning fish.
D. Perspective This section has addressed some of the mechanisms underlying hormone action in the testis. Three main topics were covered: the molecular mechanisms of androgen action in the testis, the biological role of AR in the testis,
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and the role of estrogen signaling in the testis. What are the future challenges in these diVerent research areas? With regard to androgen action, the field is only just beginning to identify primary androgen‐response genes in the testis. It is crucial that more genes of this class are identified by genome‐wide searches for ARE‐containing genes expressed in the testis. Another approach is to screen for genes rapidly induced (or repressed) in response to androgens; relevant to this, a microarray analysis study recently identified several genes rapidly responsive to testosterone in Sertoli cells (Zhou et al., 2004). There is also a need to identify primary androgen‐response genes in the two other major AR‐positive cell types in the testis: Leydig cells and myoid (peritubular) cells. Another challenge for the field is the identification of bona fide secondary androgen‐response genes relevant to spermatogenesis. Secondary androgen‐ response genes are commonly defined using two criteria: (1) delayed induction after androgen treatment; and (2) lack of discernible AREs in their promoters. The first criterion can give false positives; as an example, the Pem/Rhox5 gene is induced with delayed kinetics (Zhou et al., 2004), yet it is probably a primary (not a secondary) response gene, as it contains AREs essential for transcription in cell lines (Barbulescu et al., 2001; Geserick et al., 2003) that are housed in a 0.3‐kb promoter region essential for androgen‐dependent expression in transgenic mice in vivo (Rao et al., 2002b, 2003). While the explanation for the delayed kinetics of Pem/Rhox5 is not known, it may result from delayed expression and/or activity of a limiting factor that participates with AR to activate Pem/Rhox5 transcription. The second criterion—lack of discernible AREs in a promoter—is useful as an indicator but is not a foolproof identifier of secondary androgen‐response genes. Functional AREs can diverge extensively from the consensus sequence, making them diYcult to identify, or they may be present in a distant enhancer element far from the promoter. Another diViculty in identifying secondary response genes is that vertebrate promoters are typically regulated by a constellation of transcription factors that form large multiprotein complexes, making it diYcult to discern which one of them might be androgen regulated. Another complicating factor is the crosstalk that can occur between AR and other nuclear receptors, particularly the glucocorticoid receptor, as they bind to identical DNA sequences, at least in vitro. New approaches for identifying secondary androgen response genes are clearly needed. In part, this search will be aided by the discovery of more primary androgen‐response genes encoding transcription factors, as their downstream targets will be, by definition, secondary androgen‐response genes. An understanding of androgen action in the testis will no doubt also benefit from the advances in our knowledge of AR function in other tissues, particularly the prostate (Heinlein and Chang, 2004). In addition, the
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tremendous strides that have been made in understanding how NRs other than AR work may, by analogy, help us understand how AR functions at the molecular level. In particular, there has been much progress in the identification of proteins that associate with the functional domains of NRs to provide bridges to the transcriptional machinery, modulate ligand binding, promote receptor dimerization, and regulate DNA binding (Gottlieb et al., 2004). The discovery that steroid hormones, including androgens, induce not only transcriptional responses but also transcription‐independent responses has opened up a new paradigm for hormone action. Because the discovery of nonclassical responses is relatively new, many unanswered questions remain. For example, the signal transduction mechanisms that carry the ligand‐mediated signal to the nucleus have not yet been fully elucidated. In the case of androgen action in Sertoli cells, the role of MAP kinase pathways has been clearly demonstrated (Fix et al., 2004), but the roles of other signaling pathways that have been established for AR and other NRs in nontesticular cells—including the phosphatidylinositol‐3 kinase, JAK/STAT, and Akt pathways (Papakonstanti et al., 2003; Weiss‐Messer et al., 2004)— are not yet known. It is also unclear whether other cell types in the postnatal and adult testis use the same signal transduction pathways as Sertoli cells. Another question concerns the nature of the androgen‐binding receptor that elicits nonclassical responses. Although it is clear that Sertoli cells use AR to mediate these rapid responses (Gorczynska and Handelsman, 1995; Peterziel et al., 1999; Steinsapir et al., 1991), this does not rule out the participation of other androgen‐binding proteins in Sertoli cells. Indeed, evidence suggests that other cell types respond in an AR‐independent fashion to androgen (Walker, 2003). It will be interesting to know what androgen‐binding protein(s) are responsible for AR‐independent responses; candidates include the sex hormone‐binding globulin receptor and GPR30 (Walker, 2003). The characterization of alternative membrane receptors, cofactors, and signal transduction pathways unique to nonclassical androgen responses is important, as this may facilitate the development of new tools, such as antagonists that specifically block nonclassical responses elicited by androgen. Such tools may help clarify the biological significance of nongenomic androgen responses, and they may also be useful as male contraceptives and for treating some cases of male infertility. Regardless of whether androgens act through AR, nonclassical receptors, or a combination of the two, another issue that requires further study is whether the receptors are located at the cell surface. The primary evidence that they are at the cell surface comes from studies using conjugates of testosterone and bovine serum albumin (BSA), which are not able to cross the plasma membrane by diVusion because of their large size. Several studies
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have shown that testosterone‐BSA conjugates are capable of inducing rapid responses in a variety of cell types, including Sertoli cells, T cells, macrophages, and osteoblasts (Benten et al., 1999a; Lyng et al., 2000; Wunderlich et al., 2002). For example, a study using LNCaP prostate cells showed that testosterone‐BSA is able to trigger several signal‐transduction events, including phosphorylation of focal adhesion kinase, the association of focal adhesion kinase with phosphatidylinositol‐3 kinase, and activation of Rac1 (Papakonstanti et al., 2003). While supportive, these studies have not proven that cell‐surface receptors are responsible for nonclassical responses. High concentrations of testosterone‐BSA are typically required to elicit a response, which brings up the possibility that suYcient free steroid might dissociate from the complexes and diVuse from the cell surface to bind putative intracellular receptors (Stevis et al., 1999). While our understanding of AR action in vivo is now clearer as a result of the three aforementioned studies examining the phenotypic consequences of AR loss in Sertoli cells in vivo (Chang et al., 2004; De Gendt et al., 2004; Holdcraft and Braun, 2004), many questions remain. For example, both the myoid cells that surround the seminiferous epithelial tubules and the Leydig cells in the interstitium express high levels of AR, but AR’s function in these cells is not known. Another crucial issue is the nature of the androgen‐dependent circuitry between the many cell types in the testis. This circuitry is likely to be quite complex, as multiple cell types in the testis respond to androgens by expressing secreted molecules that have the potential to interact with other testicular cell types. Particularly important is the androgen‐dependent communication between Sertoli and Leydig cells. The complexities of this circuitry are exemplified by the divergent eVects on testosterone synthesis observed by the three groups that knocked out AR specifically in Sertoli cells in mice in vivo (see Chang et al., 2004; De Gendt et al., 2004; Holdcraft and Braun, 2004). Further dissection of the mechanisms of AR action using cell type‐specific models may lead to the development of new, more eVective strategies for the treatment of male infertility. Another area requiring further research is the role of estrogens in male fertility. Although the controversy concerning the role of estrogens in spermatogenesis has died down—as the importance of estrogens is now clear—little is known about their mechanism of action in the testis. One area of future research is the elucidation of the precise roles of estrogens in the testis, including their direct and indirect roles in spermatogenesis. This will be a complicated task, as estradiol can induce the production of secreted factors that influence male fertility. For example, since estradiol can induce FSH production, its ability to rescue fertility in gonadotropin‐deficient hypogonadal mice is probably due to both direct and indirect eVects (Ebling et al., 2000). Another important area for future research is the identification of molecular targets of estrogen in the testis. As described in Section II.C.3, very
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few estrogen‐regulated genes have been identified in the testes, and none have yet been definitively shown to use classical EREs. Microarray analysis has allowed the field to identify a plethora of androgen‐regulated testis genes in diVerent developmental stages under diVerent hormonal conditions from both normal and hormone signaling‐deficient mice (Sadate‐Ngatchou et al., 2004; Small et al., 2004; Zhou et al., 2004). Analogous approaches should be employed to identify estrogen‐regulated genes. The practical consequences of ER action in the testis, including the eVect of environmental estrogen‐like compounds on male fertility, also need to be determined. It is also possible that existing or new estrogen‐signaling agonists could cure some forms of male infertility and that estrogen‐signaling antagonists could serve as male contraceptives. The latter might have several potential advantages over androgen‐signaling inhibitors, including the possibility that male‐specific features besides fertility (such as libido and physical phenotype) would not be aVected. Clearly, there is much left to do with regard to the role of estrogen action in the testis and its practical consequences.
III. Transcription Factors with Testicular Functions This section focuses on transcription factors that have essential roles in postnatal and adult testis function in vivo (Table I). Our chapter does not attempt to cover all transcription factors but highlights those whose ablation (typically by targeted gene disruption in mice) results in clear‐cut disruption of spermatogenesis. The role of transcription factors directly involved in hormone action, such as AR and ER, are covered in Section II.
A. CREM and CREB Transcriptional responses triggered by the secondary messenger cAMP are typically mediated through CREs, specific cis elements present in the promoters of cAMP‐responsive genes. This element has a palindromic consensus sequence (TGACGTCA) that interacts specifically with highly conserved transcription factors that have in common a basic leucine zipper DNA‐ binding domain (Sassone‐Corsi, 1995). In mammals, these transcription factors include CREB, CREM, and ATF‐1. CREB and CREM are phosphorylated by several kinases, including the cAMP‐dependent kinase protein kinase A, thus providing a molecular basis for how these transcription factors function in cAMP‐dependent signaling pathways (Sassone‐Corsi, 1995). Both CREM and CREB are considered master control genes that govern diverse physiological processes, including memory, circadian rhythm, growth, and spermatogenesis.
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1. CREM Many isoforms of CREM are generated by alternative splicing; some are transcriptional activators and others are repressors. The particular isoform of CREM that has a role in spermatogenesis is CREM‐ (Sassone‐Corsi, 1998). The initial indication that CREM‐ might be important for spermatogenesis came from the discovery that its expression is restricted to the testes in mice and that the level of CREM‐ transcripts is regulated during spermatogenesis (Foulkes et al., 1992). CREM‐ is not expressed in prepubertal mice testes, but its expression in testes dramatically increases as mice reach adulthood, achieving levels hundreds of times higher than in any other adult tissue (Foulkes et al., 1992). This developmental switch occurs as a result of the use of an alternative polyadenylation site that generates a more stable CREM mRNA encoding the CREM‐ isoform (Foulkes et al., 1993). CREM‐ is expressed from the pachytene spermatocyte stage onward; its mRNA accumulates to high levels in later‐stage germ cells, suggesting it might have a role in late stages of spermatogenesis. This suggestion proved to be correct. Targeted disruption of CREM causes male sterility as a result of spermatogenic arrest at the round‐spermatid stage (Blendy et al., 1996; Nantel et al., 1996). The loss of CREM results in downregulation of several key postmeiotic genes encoding structural proteins required for spermatid diVerentiation, including the transition proteins TP1 and TP2, the protamines PN1 and PN2, proacrosin, and calspermin (Sassone‐Corsi, 1998). All of these genes appear to be direct targets of CREM‐, as their promoters possess CREs capable of being bound and activated by CREM‐ (Sassone‐Corsi, 1998). Thus, CREM‐ appears to be a master controller of several key genes essential for postmeiotic diVerentiation events in germ cells. To activate its downstream targets, CREM‐ must associate with coactivator molecules. Because CREM‐ is unphosphorylated in male germ cells (De Cesare et al., 1999), it must be activated by a mechanism independent of phosphorylation and CREB‐binding protein, unlike other CREB and CREM isoforms. To identify other regulatory molecules that might work with CREM‐ to control transcription, yeast two‐hybrid screening was used to identify CREM‐‐binding proteins. This led to the identification of a protein called activator of CREM in testes (ACT), a LIM domain‐ containing protein that is abundantly and exclusively expressed in testes (Fimia et al., 1999). ACT colocalizes with CREM in spermatids, and both molecules share the same developmental expression pattern in testes (De Cesare et al., 1999). ACT is a CREM coactivator, as it has intrinsic transactivation potential and is able to convert CREM‐ into a potent transcriptional activator. Studies in yeast showed that ACT activates CREM‐ by a phosphorylation‐independent mechanism, as yeast lacks
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CBP and other molecules involved in the phosphorylation of CREM and CREB (De Cesare et al., 1999; Fimia et al., 1999). Thus, in postmeiotic male germ cells, ACT provides a tissue‐specific, phosphorylation‐independent means to transcriptionally activate CREM‐‐dependent genes. Rather than being a constitutively nuclear protein, ACT accumulates in the cytoplasm when the downstream targets it shares with CREM‐ are transcriptionally inactive. This suggests that regulation of ACT’s subcellular localization is crucial for controlling CREM‐/ACT activity. ACT’s subcellular localization was shown to be controlled by KIF17b, an ACT‐binding protein identified by yeast two‐hybrid analysis (Macho et al., 2002). KIF17b is a testis‐specific kinesin highly related to another kinesin, KIF17, that functions as a motor protein in the brain. ACT and KIF17b colocalize in the same cell types in a stage‐specific manner during spermatogenesis, and both occupy the same subcellular compartment at diVerent stages of germ‐ cell development. In round spermatids, both ACT and KIF17b are nuclear. When spermatids begin to elongate during stage IX, KIF17b localization shifts to the cytoplasm, which also coincides with a depletion of ACT from the nucleus. This coincidental localization pattern suggested that KIF17b is a transporter of ACT, removing it from the nucleus, thereby leading to transcriptional shut‐oV of genes dependent on ACT and CREM‐. This model was supported by the finding that overexpression of KIF17b in mammalian cell lines excludes ACT from the nucleus and results in decreased promoter activity of CREM‐dependent genes, including calspermin and angiotensin‐converting enzyme (Macho et al., 2002). Surprisingly, the export of ACT to the cytoplasm does not require a functioning motor domain in the KIF17b protein (Kimmins et al., 2004). This suggests a transport mechanism for KIF17b that contrasts with the motor‐driven mechanism of other known kinesins, all of which associate with microtubules to direct the movement of organelles, vesicles, and proteins. Given the evidence that ACT is a coactivator of CREM‐, one might expect that loss of ACT in mice would cause similar phenotypic eVects as loss of CREM, but this is not the case. Mice lacking ACT have some male reproductive defects, such as a twofold reduction in caudal sperm numbers, reduced sperm motility, and increased abnormalities in sperm heads and tails, but they are fertile and contain germ cells in all developmental stages, including the spermatid stages missing in CREM‐deficient mice (Kotaja et al., 2004). Furthermore, expression of CREM‐dependent target genes, including TP1, PN1, and PN2, is not aVected in ACT‐deficient mice. This clearly indicates that there must be additional coactivators for CREM‐ besides ACT. It remains to be seen whether related LIM‐domain‐only family members (Fimia et al., 2000) compensate for the loss of ACT, or whether novel coactivators exist that work with CREM‐ to regulate spermatid gene expression.
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2. CREB In comparison with CREM, CREB has a less well‐defined role in spermatogenesis. Three approaches have been used to examine its role: partial ablation of the CREB gene (the and isoforms) (Hummler et al., 1994), complete disruption of expression of all CREB isoforms ( and ) (Rudolph et al., 1998), and expression of a dominant‐negative form of CREB (Scobey et al., 2001). The partial knockout mice were originally reported to be viable and fertile (Hummler et al., 1994). However, subsequent studies showed that these mice were sometimes subfertile and produced fewer gametes than their wild‐type littermates, primarily because the mutation interferes with germ‐cell development at the spermatocyte stage (Scobey et al., 2001). The mild and variable nature of the partial knockout phenotype may be the result of compensatory upregulation of other CREB isoforms (and the related factor CREM and ATF‐1) when transcription of CREB and CREB is diminished. Complete knockout of all CREB isoforms did not provide any additional clues about the spermatogenic role of CREB, as these mice die shortly after birth (Rudolph et al., 1998). To selectively deplete CREB function under conditions that avoid perinatal lethality, an adenovirus construct was made to express a dominant‐ negative form of CREB containing mutations that prevented it from being phosphorylated (and thus activated). Injection of this construct into the rat testis resulted in increased germ‐cell apoptosis and a dramatic reduction of round spermatids in more than half of the exposed tubules (Scobey et al., 2001). The success of this approach may be partially attributed to the fact that adenovirus absorption by Sertoli cells is most eYcient during spermatogenic stages II–VI, when CREB is expressed at peak levels (Blanchard and Boekelheide, 1997). When the dominant‐negative CREB construct was expressed in primary Sertoli cells (Hall et al., 1988; Scobey et al., 2001), this resulted in complete inhibition of the expression of c‐Fos, a CREB‐dependent transcription factor induced by FSH in Sertoli cells (Hall et al., 1988; Scobey et al., 2001). The finding that c‐Fos is regulated by CREB is intriguing, but the functional role of c‐Fos in Sertoli cells remains to be determined. There is also a need to identify other CREB target genes in Sertoli cells; candidates include the many other cAMP‐regulated genes expressed in testis that have been discovered (for a list, see Hummler et al., 1994; Scobey et al., 2001). B. Nonsteroid Hormone Nuclear Receptors Orphan NRs are a large class of transcription factors that possess structural similarities to steroid hormone receptors but have no known ligands (Shiau et al., 2001). Some orphan NRs may function in a ligand‐independent
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manner and others may require a ligand that has yet to be identified. Two orphan NRs that have been implicated in spermatogenesis will be discussed. One of them, testicular receptor 4 (TR4), appears to have a role in the timing of the first wave of spermatogenesis. In the testis, TR4 is specifically expressed in primary spermatocytes during meiotic prophase. Peak TR4 expression occurs on day 21 p.p. (the end of meiotic prophase) during the initial wave of spermatogenesis, but expression persists in subsequent waves. TR4‐deficient mice exhibit reduced fertility and produce 50% less sperm than their wild‐type littermates (Mu et al., 2004). Interestingly, the first wave of spermatogenesis is markedly delayed in TR4‐deficient mice, and the timing to reach meiotic prophase and subsequent meiotic divisions is drastically lengthened. As evidence of this, the expression of specific markers of the terminal stages of meiosis (cyclin A1 and sperm1) is delayed during the first wave of spermatogenesis. Ultimately, this germ‐cell progression defect leads to prolonged or arrested spermatogenesis at stages XI–XII in adult mice. Like many other NRs, TR4 can either activate or repress transcription (Zhang and Dufau, 2004). For example, TR4 stimulates the promoters of the genes encoding thyroid hormone and ciliary neurotrophic factor, and it represses promoter of the gene encoding the steroid 21‐hydroxylase, all through well‐defined TR4‐binding sites (Lee et al., 1999; Young et al., 1998). Other genes, such as cyclin A1 and sperm1, lack consensus TR4‐ binding sites and may thus be indirectly regulated by TR4. Alternatively, TR4 may directly regulate these genes by collaborating with its cofactors, which have been largely uncharacterized. It is interesting to note that TR4 can form heterodimers with several hormone receptors of known function, including receptors for retinoic acid and other retinoids, vitamin D, PPAR, androgens, and estrogens (Lee et al., 1996; Shyr et al., 2002; Zhang and Dufau, 2004). This ability of TR4 to heterodimerize is functionally relevant; for example, the association of TR4 with AR mediates the transcriptional repression of genes that contain both ARE‐ and TR4‐binding elements (Lee et al., 1999). While gene network patterns are emerging for orphan NRs such as TR4, little is known about the molecular mechanism by which these orphan NRs regulate target genes. Ultimately, the transactivation of their target genes may require the formation of large complexes composed of orphan NRs, steroid hormone receptors, and as‐yet‐unidentified cofactors. For example, the corepressor TRA16 has been identified as a high‐aYnity binding partner for TR4 (Yang et al., 2003). TR4 and TRA16 colocalize in the nuclei of H1299, MCF‐7, and LNCaP cells, and TRA16 expression plasmids inhibit reporter constructs containing TR4‐response elements in a dose‐dependent manner. Electrophoretic mobility shift analysis has revealed that TRA16 blocks the action of TR4 by decreasing its aYnity for TR4‐binding elements
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(Yang et al., 2003). It remains to be seen whether TRA16 has a role in TR4‐mediated transcriptional responses in the testis. Another orphan NR expressed during spermatogenesis is TR2 (Chang and Kokontis, 1988; Chang et al., 1994). TR2 is expressed in postnatal testis on day 16 p.p., is massively upregulated by day 28 p.p., and remains high in adult testes, where its expression is confined to round and elongating spermatids (Lee et al., 1995). TR2 may be important for spermatogenesis, as two animal models with induced testicular defects leading to loss of germ cells (surgically induced cryptorchidism and vitamin A depletion) both exhibit reduced TR2 expression in the testis (Lee et al., 1996; Mu et al., 2000). However, phenotypic analysis of TR2‐deficient animals did not reveal any defects in testis development or adult fertility (Shyr et al., 2002). It remains to be determined whether TR2 has a minimal role in spermatogenesis, a context‐dependent role, or a role compensated by a related molecule such as TR4.
C. Heat‐Shock Factors Hsfs are best known for their ability to transcriptionally activate genes in response to heat shock, but in the absence of heat shock they also regulate genes involved in both embryonic development and spermatogenesis (Kallio et al., 2002; Morimoto, 1998; Wang et al., 2003, 2004; Wu, 1995). Hsf1, Hsf2, and Hsf4 are the primary vertebrate Hsf proteins. They all exist as monomers in the cytoplasm, but upon activation by a variety of diVerent environmental cues they become phosphorylated and translocate to the nucleus where they associate with other factors to form high‐molecular‐ weight transcription‐promoting complexes (Kline and Morimoto, 1997; Knauf et al., 1996). The first indication that Hsfs might have a role in spermatogenesis came from the discovery that Hsfs are expressed at high levels in the developing mouse testis (Loones et al., 1997). Their functional role was further suggested by the discovery that spermatogenesis is blocked in transgenic mice that overexpress Hsf1 (Nakai et al., 2000). However, the functional role of Hsfs was not made clear until Hsf‐knockout mice were generated. Ablation of Hsf2 was found to cause male subfertility associated with a marked reduction in testis weight, 70% reduction in sperm counts, and loss of spermatocytes as a result of increased apoptosis (Wang et al., 2003). In contrast, loss of Hsf1 resulted in no apparent defects in spermatogenesis, but mice lacking both Hsf1 and Hsf2 had a severe defect in spermatogenesis that resulted in complete sterility (Zhang et al., 2002). Together, these data indicate that Hsf1 and Hsf2 serve partially redundant but crucial roles in spermatogenesis (Kallio et al., 2002; Wang et al., 2003, 2004).
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The earliest testicular defect in mice lacking both Hsf1 and Hsf2 is at the juvenile stage (day 18 p.p.). Unlike control mice, these double‐mutant mice had virtually no late spermatocytes or spermatids, as the germ cells that entered meiotic prophase failed to progress beyond the pachytene stage. Microarray analysis revealed that the expression of over 100 genes was strongly reduced (more than fivefold) in the testes of these double‐mutant mice. Most were probably reduced simply because of the lack of mature germ cells, but some of the diVerentially expressed genes may be bona fide targets of Hsf1 and/or Hsf2, including meg1 (a cell‐cycle protein expressed in early meiotic germ cells), Xmr (a synaptonemal complex‐promoting factor), and the homeobox genes Hoxa4 (expressed during early meiosis) and Pem/Rhox5 (expressed in Sertoli cells). The latter may be functionally relevant, as Pem/Rhox5‐null mice and Hsf2‐null mice have a similar phenotype (see MacLean et al., 2005 and Section III.H.2). D. Sox Sox proteins are transcriptional regulators that have in common a 79‐amino‐ acid DNA‐binding domain known as the HMG box. Sox proteins have been shown to bend DNA by binding to it at two positions, which may be crucial for their ability to regulate transcription (Love et al., 1995). The founding member of the Sox family is the Y‐chromosome gene Sry, which is responsible for directing the bipotential embryonic gonad to develop down the male pathway (Koopman et al., 1991). Other members of the Sox family are also expressed in the testis, including Sox3, Sox8, Sox9, and Sox10. The X‐linked Sox3 gene has been suggested to be the ancestral precursor to Sry. Sox3‐null male mice have small testes (40% smaller than wild‐type controls) and have about threefold less epididymal sperm than control mice (Weiss et al., 2003). Rather than being aVected at a defined stage of germ‐cell development, these mutant mice have heterogeneous defects that vary among tubules. The least aVected tubules have round and elongated spermatids that have detached from the seminiferous epithelium and migrated to the tubule lumen; the most severely aVected tubules have the ‘‘Sertoli‐cell‐only’’ phenotype in which germ cells are entirely absent. The cause of the germ‐cell defects in Sox3‐null mice may be an indirect eVect of Sox3, as immunohistochemical analysis revealed that Sox3 is expressed at high levels in the nuclei of Sertoli cells (Weiss et al., 2003). However, Sox3 is also expressed in germ cells, raising the possibility that Sox3 directly regulates germ‐cell diVerentiation and/or survival (J. L. Jameson, personal communication). Does Sox3 also have an overt eVect on Sertoli cells themselves? Sox3 is expressed when Sertoli cells are undergoing proliferation, but its targeted
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disruption has no eVect on the number of Sertoli cells present after this proliferative phase, indicating that Sox3 does not promote Sertoli‐cell mitosis (Weiss et al., 2003). However, loss of Sox3 does aVect Sertoli‐ cell morphology, as the cytoplasm of Sox9‐deficient Sertoli cells acquires numerous large vacuoles. These large vacuoles have also been observed in Sertoli cells of transgenic mice overexpressing the antiapoptotic gene Bcl2 (Yamamoto et al., 2001), mice deficient in inositol polyphosphate 5‐phosphatase (Hellsten et al., 2002), and mice expressing a dominant‐negative form of the WT1 protein in Sertoli cells (M. Rao and M. F. W., unpublished observations). In addition to large Sertoli‐cell vacuoles, all of these mutant mice share other common features, including significant loss of germ cells, visible disruption of the seminiferous tubules, and reduced epididymal sperm counts. This suggests the possibility that these vacuoles develop as a result of the increased Sertoli‐cell phagocytic activity required to engulf the large number of dead germ cells that arise in all of these mutant models. It will be interesting to determine whether these large vacuoles are pathological or simply a normal consequence of massive phagocytosis. In the case of mice deficient in inositol polyphosphate 5‐phosphatase, these vacuoles appear to be ordered structures with well‐defined endosomal‐like membranes and not simply a de facto result of loss of germ‐cell contacts that normally define the cytoplasm of Sertoli cells (Hellsten et al., 2002). Interestingly, the vacuole membranes are coated with actin, contain intact adherens junctions, and stain positive for N‐cadherin and beta‐catenin, suggesting that endosomes may be formed from internalization of germ cell–bound plasma membrane that is engulfed by Sertoli cells (Hellsten et al., 2002). Sox3 binds to the promoter of the gene encoding steroidogenic factor‐1, which may be relevant to its putative function in fetal Sertoli cells (Shen and Ingraham, 2002). However, Sox3’s targets in postnatal and adult Sertoli cells are not known. Once future studies uncover these targets, it will be interesting to know whether Sox3 regulates their transcription by bending DNA, as do other HMG‐box transcription factors. There are more than 20 other known Sox factors, several of which might have a role in spermatogenesis based on their expression patterns. For example, the E group of Sox genes (Sox8, Sox9, and Sox10) are expressed in an overlapping manner in Sertoli cells at diVerent points of postnatal and adult testis development (Schepers et al., 2002). In the adult testis, Sox9 is expressed in a stage‐specific manner (primarily in stage VII) during the seminiferous epithelial cycle (Frojdman et al., 2000). While the functional significance of this postnatal and stage‐specific adult expression is not known, studies in knockout and transgenic mice have shown that Sox9 and Sox8 have essential roles in gonad development during embryogenesis (Bi et al., 1999; Chaboissier et al., 2004; Sock et al., 2001). Both are expressed in Sertoli cells beginning at day 12 and peaking at day 16
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post coitum (p.c.), where they work in concert with GATA factors, WT1, and steroidogenic factor‐1 to activate anti‐Mu¨ llerian hormone expression (Schepers et al., 2003). To determine the postnatal and adult testicular functions of Sox8 and Sox9, it will be necessary to selectively inactivate them in the testis, as targeted disruption of either causes embryonic lethality (Bi et al., 1999; Sock et al., 2001). E. Plzf The zinc‐finger family is the largest class of transcription factors in higher eukaryotes. Mammals are estimated to have at least 700 zinc‐finger genes (Lander et al., 2001). Zinc‐finger proteins have in common a tetrahedral arrangement of four zinc ions that stabilize four amino acids (generally a combination of Cys and His side chains) that form the DNA‐binding domain (Parraga et al., 1988). Several zinc‐finger family members are expressed in the testis (Hirose, 1999; Tohonen et al., 2003), but in most cases the functional relevance of their expression is not known. Here we will focus our attention on the only zinc‐finger transcription factor so far demonstrated to have a clear role in spermatogenesis: promyelocytic leukemia zinc factor (Plzf; also known as Zfp145 [zinc‐finger protein 145]). Plzf was initially characterized as a developmentally regulated embryonic transcription factor that is crucial for limb patterning and skeleton development (Barna et al., 2000). Later it was discovered that Plzf is also expressed in the developing male gonad; it is first expressed on day 17.5 p.c. and is maximally expressed on day 7 p.p. when nondividing gonocytes are undergoing conversion into rapidly dividing spermatogonia (Costoya et al., 2004). Thereafter, Plzf expression is restricted to a subset of spermatogonia in both postnatal and adult testes. In adult testes, Plzf is expressed specifically by spermatogonia that exhibit stem cell‐like properties and express Oct‐4, a transcription factor previously implicated in maintaining stem‐cell populations (Buaas et al., 2004; Costoya et al., 2004). While the restriction of Plzf expression to stem cell‐like spermatogonia is intriguing, the critical test of its functional role is the phenotypic consequences of its loss in vivo. Two studies (Buaas et al., 2004; Costoya et al., 2004) addressed this issue by examining naturally occurring Plzf‐mutant (luxoid) mice and Plzf‐knockout mice generated by targeted deletion, respectively. Both types of Plzf‐mutant mice were found to exhibit a deficit in spermatogonia that progressively worsened as the animals aged. This deficit is not the result of a failure of precursor cells to develop, as the number of gonocytes is normal, but rather is a consequence of loss of self‐renewal, leading to increased spermatogonial apoptosis. Transplantation of spermatogonia from Plzf‐null animals failed to repopulate gonads that had been
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chemically depleted of germ cells, indicating that the defect lies within the germ cells themselves and not the failure of Plzf‐null Sertoli cells to support development (Costoya et al., 2004). As further evidence, Plzf‐deficient mice have no obvious defects in Sertoli cells or seminiferous‐tubule architecture. Microarray analysis of Plzf‐null and control spermatogonia identified several candidate Plzf‐regulated genes. Many of them encode transcription factors, RNA‐binding proteins, enzymes involved in metabolism, and cell‐ cycle control proteins (Costoya et al., 2004). None were likely to be direct targets of Plzf regulation, as none had obvious Plzf‐binding sequences in their promoters. Instead, they are probably indirectly regulated by Plzf via intermediary factors. Interestingly, none of these genes were previously known to be involved in spermatogenesis. The discovery that a subset of them may be regulated by a germ‐cell renewal factor may encourage investigation into this possibility.
F. Dmrt1 Doublesex and Mab‐3‐related transcription factor‐1 (Dmrt1) is a highly conserved vertebrate gene encoding a transcription factor with a DM domain, a cysteine‐rich DNA‐binding motif (Erdman and Burtis, 1993; Zhu et al., 2000). Structurally related genes involved in sexual development (mab‐3, mab‐23, and doublesex) are also found in flies and worms (Raymond et al., 1999). Dmrt1 is first expressed in genital ridge tissues that ultimately give rise to the gonads (De Grandi et al., 2000; Raymond et al., 1999; Smith et al., 1999). In mice, Dmrt1 mRNA appears to be expressed by both Sertoli and germ cells in the primordial gonad (Raymond et al., 1999). Dmrt1 protein levels are upregulated in male germ cells on day 1 p.p., reaching peak levels on day 7 p.p., just before meiosis initiates (Raymond et al., 2000). In adult mouse testes, both premeiotic germ cells (spermatogonia) and Sertoli cells express Dmrt1 protein (Raymond et al., 2000). Phenotypic analysis of Dmrt1‐null mice revealed that Dmrt1 is essential for both postnatal testis diVerentiation and normal Sertoli‐cell proliferation (Raymond et al., 2000). Dmrt1‐null mice have underdeveloped testes, almost no germ cells, and an overabundance of immature Sertoli cells (Raymond et al., 2000). The germ‐cell defect is the result of the failure of gonocytes to migrate to the periphery of the seminiferous epithelium and proliferate, ultimately culminating in their death between days 7 and 10 p.p. Cell type– specific knockouts created using the Cre/loxP system have shown that Dmrt1 is required in a cell‐autonomous manner in both Sertoli and germ cells (D. Zarkower, personal communication). Interestingly, humans possessing only a single copy of the chromosome‐9 region harboring both an ortholog and a paralog of Dmrt1 (DMRT1 and DMRT3, respectively)
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have male reproductive defects similar to those in Dmrt1‐null mice (Veitia et al., 2001). G. CAF1 Chromatin assembly factor‐1 (CAF1) is a highly conserved protein found in mammals, plants, Caenorhabditis elegans, and Saccharomyces cerevisiae that appears to regulate both transcription and mRNA decay (Denis and Chen, 2003). CAF1 was originally genetically defined as a transcription factor that acts together with another factor, CCR4, to regulate the expression of a variety of genes involved in cell‐wall integrity and nonfermentative growth (Draper et al., 1995; Sakai et al., 1992). CAF1 and CCR4 were later shown to be components of a multi‐subunit complex required for the cytoplasmic deadenylation of many mRNA species (Tucker et al., 2001). Because deadenylation is both the initial and the limiting step for the degradation of most yeast mRNAs, CAF1 may be crucial for dictating the overall rate of mRNA decay. CAF1‐deficient male mice were found to be sterile as a result of a testicular defect (Berthet et al., 2004). No other significant abnormalities were observed, which is surprising as CAF1 is ubiquitously expressed in all tissues. The first observable defect during the first wave of spermatogenesis is the accumulation of vacuoles in the cytoplasm of the Sertoli cells at day 19 p.p. By day 22, the number of spermatids is decreased relative to littermate controls. The decrease in spermatids is caused by their degeneration, based on TUNEL and Comet analysis, but whether their death is the result of apoptosis or autophagy remains uncertain. Seminiferous‐tubular morphology worsens with age; the adult testes have disorganized and atrophied tubules, massive Sertoli‐cell vacuolization, and signs of apical germ‐cell sloughing and shedding. In contrast to the homozygous mutant mice, heterozygous CAF1‐deficient males are fertile and give rise to a normal Mendelian ratio of progeny, indicating that haploid CAF1‐deficient spermatids diVerentiate normally and give rise to normal spermatozoa. This implies that CAF1 is not essential for the final (haploid) stages of spermatogenesis and instead suggests that the primary defect lies either in (i) the diploid germ cells or (ii) the supporting Sertoli cells, both of which express CAF1. The former, the Sertoli‐cell vacuolization, could be the indirect result of degenerating spermatids, which could either overwhelm the Sertoli‐cell phagocytic machinery or release protamines, which are known to exert deleterious eVects on epithelial cells (Peterson and Gruenhaupt, 1992). Evidence for the latter case is that the CAF1‐null mice failed to exhibit a stage‐specific arrest of spermatogenesis, a phenotype typically seen for primary germ‐cell defects. Further evidence
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for a primary Sertoli‐cell defect is that the vacuolization of Sertoli cells precedes the first sign of germ‐cell degeneration. A final line of evidence is the finding that Sertoli‐cell toxins cause the same sequence of events that were observed in CAF1‐null mice: vacuolization of the Sertoli‐cell cytoplasm followed by germ‐cell shedding and eventual disappearance of all germ‐cell subsets (Chaboissier et al., 2004). To definitively delineate whether loss of CAF1 causes a primary defect in Sertoli cells or diploid germ cells, it will be necessary to selectively inactivate CAF1 independently in each cell type. CAF1 may be part of a multiprotein complex that regulates reproduction. CAF1 interacts with BTGAPRO proteins in mammalian cells; at least one member of this family, BTG1, is expressed in both Sertoli and germ cells (Raburn et al., 1995). The ortholog of BTGAPRO in C. elegans, FOG‐3, is crucial for C. elegans germ cells to initiate spermatogenesis (Chen et al., 2000). Although limited data exist at present, it is possible that there may be a conserved CAF1/BTG multiprotein complex in both higher and lower eukaryotes that serves to regulate spermatogenesis.
H. Homeobox Genes Homeobox genes encode transcription factors containing a 60‐amino‐acid DNA‐binding motif known as the homeodomain. More than 150 homeobox genes exist in higher eukaryotic genomes; they are grouped in subfamilies based on homeodomain sequence and gene structure. Best known is the highly conserved Hox subfamily, which governs a diverse realm of embryonic developmental processes, including body‐axis formation, organogenesis, and limb development (Weatherbee et al., 1998). The formation of the male urogenital tract is also regulated by some Hox subclass genes (Lindsey and Wilkinson, 1996c; Rao and Wilkinson, 2002). Non‐Hox homeobox subfamily members, including Emx2, Lhx9, Arx, and Pax2, also have a major role in dictating male reproductive tract development (Lindsey and Wilkinson, 1996c; Rao and Wilkinson, 2002). Less is known about the role of homeobox genes in postnatal and adult reproductive events, including spermatogenesis. This is surprising, as over 40 homeobox genes are expressed in postnatal and adult testes (Lindsey and Wilkinson, 1996c; Rao and Wilkinson, 2002). EVorts to clarify the roles of some of these homeobox genes have been clouded by putative functional redundancies and embryonic lethality. Knockout mice for many homeobox genes (mainly Hox genes) do not exhibit any obvious defects in the reproductive tract, including spermatogenesis (Lindsey and Wilkinson, 1996c). It is not known whether this is because of functional redundancy (Hox genes are known to compensate for each other) or because some homeobox genes have little or no role in reproduction (i.e., their expression in the
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reproductive tract is superfluous). In other cases, targeted disruption of homeobox genes, including Oct3, Testis1 (Oct6), HoxA5, and Cart1, causes embryonic lethality, thereby precluding analysis of their role in postnatal and adult events, including spermatogenesis (Lindsey and Wilkinson, 1996c; Rao and Wilkinson, 2002). Although most homeobox gene knockout mice have failed to yield information on the role of homeobox genes in testis function, two homeobox‐null mutant mice have proven useful in this regard. Interestingly, neither disrupts a Hox subclass gene; one (Sperm1) is a member of the POU subclass and the other (Pem/Rhox5) is a member of the newly discovered homeobox subfamily Rhox. As discussed later, targeted disruption of Sperm1 and Pem/Rhox5 specifically impairs the normal function of the testis, not its formation. 1. Sperm1 Sperm1 is expressed exclusively in diVerentiating germ cells in the male reproductive tract, suggesting that it might have a role in spermatogenesis. Consistent with this role, Sperm1‐null male mice are less fertile than littermate controls, based on the average number of progeny generated after cohabitation with normal females for 8 weeks (Pearse et al., 1997). Unfortunately, neither the molecular nor the cellular basis for this subfertile phenotype is known. The testes of these knockout mice develop normally and display normal morphology. Their sperm count and motility are not significantly lower than those of normal mice, and caudal sperm from Sperm1‐null mice are able to eYciently perform in vitro fertilization. Postmeiotic events, including nuclear condensation, acrosome formation, and assembly of the mid‐piece and flagella structures, are all normal in Sperm1‐null animals. Lastly, several germ‐cell diVerentiation markers do not exhibit any significant alterations in Sperm1‐null testes. Clearly, future studies will be required to reveal the underlying mechanism responsible for the subfertility of Sperm1‐null male mice. 2. Pem/Rhox5 Pem/Rhox5 is a homeobox gene originally cloned by subtraction hybridization from a mouse T‐cell lymphoma clone (Lindsey and Wilkinson, 1996c). Consistent with the source of its isolation, Pem/Rhox5 is widely expressed in diverse tumors from many tissues and cell types (Rao et al., 2002a). While aberrantly expressed in tumors, Pem/Rhox5 is normally expressed in a stage‐specific and cell type–specific manner specifically in male and female reproductive tissues (Lindsey and Wilkinson, 1996b; Maiti et al., 1996). An initial analysis of Pem/Rhox5‐null mice detected no obvious
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abnormalities (Pitman et al., 1998), but follow‐up analysis revealed that the mutant males are subfertile (MacLean et al., 2005). Their testes have an increased frequency of apoptotic meiotic spermatocytes across all stages of spermatogenesis, and they have 50% fewer mature spermatids and epididymal spermatozoa than littermate controls. Because Pem/Rhox5 is not expressed by germ cells, but rather is restricted to Sertoli cells in the testis (Lindsey and Wilkinson, 1996b; Pitman et al., 1998; Sutton et al., 1998), Pem/Rhox5 probably regulates the expression of Sertoli‐cell genes that encode secreted or cell‐surface molecules that regulate the survival of germ cells undergoing meiosis. Pem/Rhox5‐null mice also have a depressed frequency of epididymal sperm displaying forward progressive movement. This could be caused by either a Sertoli‐cell defect that does not allow eYcient maturation of motile spermatozoa or a defect in the caput epididymis, the only other site of Pem/ Rhox5 expression in the male reproductive tract besides Sertoli cells (Lindsey and Wilkinson, 1996a). The latter is an attractive possibility, as the caput epididymis is the site where sperm acquire forward‐mobility competence. Gene targets of Pem/Rhox5 have not yet been identified, but microarray analysis has revealed many genes diVerentially expressed as a result of loss of Pem/Rhox5 in the postnatal testis (MacLean and Wilkinson, unpublished). Some of these genes, many of which are involved in cellular metabolism and diVerentiation, are candidates to be directly regulated by Pem/Rhox5. 3. The Rhox Gene Cluster It was discovered that Pem/Rhox5 is not alone on the X chromosome; it is part of a 12 homeobox‐gene cluster selectively expressed in reproductive tissues and placenta (MacLean et al., 2005). These Rhox (reproductive homeobox) genes are located in three subclusters; the genes within each subcluster are expressed in a pattern during testis development corresponding to their position within the subcluster. For instance, the first gene in the subcluster, Rhox1, is the first gene to be expressed during the first wave of spermatogenesis; Rhox2 is expressed second; and so on. The maximal expression of each Rhox gene during testis development also corresponds to its position, such that 50 genes are expressed at higher levels than are 30 genes. This quantitative and temporal colinearity suggests that a global enhancer dictating transcription in a distance‐dependent fashion may exist at one end of each Rhox subcluster. The colinear regulation exhibited by the Rhox gene cluster during postnatal testis development is reminiscent of the colinearity displayed by the well‐characterized Hox gene cluster during embryogenesis (Duboule and Morata, 1994). Most of the Rhox genes are expressed in testicular somatic cells; none are detectably expressed in germ cells postnatally (MacLean et al., 2005). Some
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are expressed when Sertoli cells are proliferating, and others are expressed when Sertoli cells are undergoing the final stages of terminal diVerentiation, suggesting the possibility that they are involved in both Sertoli‐cell mitosis and development. Many aspects of germ‐cell behavior, including their mitosis, meiosis, and diVerentiation, could also potentially be regulated by Rhox genes, as they are expressed in an overlapping manner during all phases of the first wave of spermatogenesis. Targeted disruption or knockdown approaches should reveal the individual and overlapping roles of the Rhox genes in spermatogenesis.
I. Perspective In this section, we discussed transcription factors with well‐defined functions in spermatogenesis. Many other transcription factors will no doubt have important roles in spermatogenesis, but at present there is insuYcient data directly demonstrating this. For example, many transcription factors exhibit developmentally regulated expression during postnatal testis development and/or stage‐specific expression in the adult seminiferous epithelium, implying (but not proving) that they have a role in testis development. Below, we briefly discuss some (but certainly not all) transcription factors that are candidates to having crucial roles in spermatogenesis. One set of transcription factors potentially important for spermatogenesis is the basic helix‐loop‐helix (bHLH) transcription‐factor family. Transfection experiments in cell lines have indicated that the binding element recognized by bHLH factors (called an E‐box) is essential for the expression of several Sertoli‐cell genes, including those encoding AR, FSH receptor (FSHR), transferrin, and steroidogenic factor‐1 (Chaudhary et al., 1997; Kim and Griswold, 2001; Scherrer et al., 2002). Unfortunately, determination of whether bHLH transcription factors have a role in spermatogenesis in vivo has been stymied by the fact that mice lacking these factors die during embryogenesis. Nonetheless, evidence for the importance of bHLH factors comes from studies on the related inhibitor of diVerentiation (Id) proteins, which are naturally occurring inhibitors of bHLH factors. Id proteins lack a functional basic DNA‐binding motif but have an HLH‐interaction domain, allowing them to form transcriptionally inactive heterodimers with bHLH factors (Norton, 2000). An emerging paradigm, based on work in Sertoli cells (Chaudhary et al., 2005) and earlier work in various other cell types (Norton, 2000), is that a delicate balance exists between Id and bHLH factors that ultimately dictates whether a cell undergoes cell division or diVerentiation. All four known Id proteins are expressed in testes. Id1 and Id4 are both expressed in spermatocytes, Id4 is also expressed in type‐A1 spermatogonia, and Id2 and Id3 are expressed in an FSH‐regulated manner
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in Sertoli cells (Chaudhary et al., 2001; Sablitzky et al., 1998; Scobey et al., 2004). This cell type– and stage‐specific expression of the Id inhibitors is consistent with the possibility that they regulate the activities of bHLH transcription factors at various stages of spermatogenesis. Id2 is likely to be involved in spermatogenesis, as Id2‐deficient mice appear to have defects in spermatogenesis (Norton, 2000), but the full details of these defects have not yet emerged. Gene targeting studies have not clarified the in vivo functions of Id1 and Id3 in mouse spermatogenesis, as loss of either factor alone has no obvious eVect on either the testis or male fertility, and loss of both factors causes embryonic lethality (Lyden et al., 1999; Pan et al., 1999; Yan et al., 1997; Yokota et al., 2001). Another family of transcription factors probably involved in spermatogenesis is the zinc finger‐containing GATA factors. Studies examining the eVect of either loss of GATA4 or expression of a dominant‐negative form of GATA4 in vivo have clearly shown that GATA factors have a role in the formation of the male gonad in the embryo (Parker and Schimmer, 2002; Tevosian et al., 2002; Tremblay et al., 2001). One of the gene targets of GATA4 in the embryonic testis is the gene encoding anti‐Mu¨llerian hormone (Tremblay et al., 2001). GATA‐factor expression persists in postnatal and adult testes; at least three of the six family members (GATA1, GATA4, and GATA6) are expressed in Sertoli cells (Kiiveri et al., 2002; Viger et al., 2004). One or more of these GATA factors appear to be essential for normal spermatogenesis, as mice expressing dominant‐negative GATA4 (which inhibits all GATA factors) in Sertoli cells suVer from a late block in spermatogenesis and progressive infertility with advancing age (R. Viger, personal communication). Candidate in vivo targets of GATA factors in postnatal and adult Sertoli cells are promoters of genes shown to be regulated by GATA transcription factors in Sertoli‐cell lines; these include the genes encoding FSHR, inhibin‐, inhibin‐, and Dmrt1 (Feng et al., 2000; Kim and Griswold, 2001; Lei and Heckert, 2004; Rey et al., 2003; Viger et al., 2004). Another candidate target is the Pem/Rhox5 gene. Its proximal promoter contains several consensus GATA sites, some of which have been shown to be essential for maximal expression in transiently transfected Sertoli‐cell lines (A. Bhardwaj and M. F. W., unpublished results; Rao et al., 2003). Other potential targets are genes specifically expressed in stages VII–IX of the seminiferous epithelial cycle, as GATA1 is expressed specifically in these stages (Yomogida et al., 1994). Conditional‐knockout mice lacking GATA1 in Sertoli cells do not suVer from obvious fertility defects (Lindeboom et al., 2003), but this is probably a result of redundancy with other GATA transcription factors, as most GATA factors bind to identical target sequences. The Brd transcription factors are expressed in the adult mouse testis and may therefore also have a role in spermatogenesis. Brdt is expressed
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specifically in testis, Brd2 and Brd3 are expressed selectively but not exclusively in reproductive tissue, and Brd4 is present in most tissues (Shang et al., 2004). Interestingly, the individual members of this family are expressed at diVerent points of male germ‐cell development: Brd4 is expressed in spermatogonia; Brdt is expressed exclusively in mid to late spermatocytes, Brd2 is expressed in spermatogonia and diplotene spermatocytes, and Brd3 is expressed in round spermatids (Shang et al., 2004). The Brd proteins are not true transcription factors, as they do not directly bind DNA; instead they have an evolutionarily conserved bromodomain that binds to acetylated lysine residues found in transcriptionally active chromatin. In the presence of acetylated H3 and H4 histones, Brd2 binds to the cell‐cycle transcription factor E2F and thereby stimulates the promoters of many cell‐cycle regulatory genes in mammary epithelial cells (Crowley et al., 2002; Denis et al., 2000; Dey et al., 2003). Brd4 also appears to be involved in cellular proliferation, based on the phenotypic eVects of a null allele of the Brd4 gene (Houzelstein et al., 2002). By analogy, both Brd2 and Brd4 may also have a role in spermatogonial proliferation in the testis. In addition, Brd factors expressed in spermatocytes (Brdt and Brd2) may respond to or somehow mediate the chromatin modifications that occur during meiosis. While there is, as of yet, no direct evidence for either a meiotic or a proliferative role for Brd proteins in mice, the chromosomal region containing the human BRD2 gene has been linked to azoospermia in humans (Matsuzaka et al., 2002). Another class of proteins that is likely to play a role in spermatogenesis is the cancer/testis antigens. These are encoded by genes enriched on the X chromosome that are highly expressed in malignant tissues but tend to be silent in all normal tissues except the testis (Kouprina et al., 2004). While most work has focused on the use of cancer/testis antigens as immunogenic targets of anticancer vaccines, the fact that they all exhibit expression in distinct cell types within the testis (typically in the germ cells) indicates that they may also have important functions in the normal testis. A direct role in transcriptional regulation has not been elucidated for most cancer/testis antigens, but many of them—including Brdt, SSX, NY‐ESO‐1, and some members of the X‐linked melanoma antigen (MAGE) and SPANX families —are present in the nuclei of spermatogonia and other germ cells (Juretic et al., 2003; Kouprina et al., 2004). The X‐linked Pem/Rhox5 homeobox gene (described earlier in this section) can also be considered a cancer/testis antigen family member, as it is widely expressed in tumors and has shown to be a tumor antigen in mice (Ono et al., 2000; Rao et al., 2002a; Wilkinson et al., 1990). Adjacent to Pem/Rhox5 on the X chromosome are several other homeobox genes expressed in testis, some of which are ectopically expressed in tumor cell lines (MacLean et al., 2005; Wayne et al., 2002). Interestingly, the N‐terminal region of these Pem/Rhox and SPANX
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proteins have significant sequence identity, suggesting they share a common domain (J. A. M. and M. F. W., unpublished observations); this region in Pem/Rhox5 has been shown to regulate the development of early stem cells (Fan et al., 1999). While much progress has been made, the elucidation of the spermatogenic roles of some transcription factors has been hindering by the fact that the ablation of some of them by standard gene targeting (1) fails to yield an observable phenotype; (2) causes embryonic defects that prevent successful mating of adults (such as failure to form reproductive organs or display normal mating behavior); or (3) causes embryonic lethality. To solve the first problem, it will be necessary to generate compound knockout mice, particularly when studying gene families, as their individual members often exhibit functional redundancy. To solve the latter two problems, testis‐ specific knockout and knockdown mice must be generated (e.g., using the Cre/loxP system and dominant‐negative proteins/siRNAs, respectively). For these studies, there is a need to develop a panel of promoters that delivers consistent and high‐level expression to particular cell types in the testis in a developmentally regulated manner. While elucidation of the discrete roles played by individual transcription factors in male gametogenesis is a challenge, it will be even more challenging to determine the relationships between these transcription factors as well as the nature of their upstream regulators and downstream targets. One of the many diYculties will be disentangling the transcriptional control executed by tissue‐specific transcription factors compared with ubiquitous transcription factors. This review has focused on the former, but the latter are also important. For example, the ubiquitously expressed Sp1 and Sp3 transcription factors regulate many genes important for spermatogenesis, including those encoding the Id repressors, homeodomain transcription factors, steroidogenic enzymes, and the Tyro3 receptor tyrosine kinases (which were shown to be required for spermatogenesis [Wong and Lee, 2002]). The transcriptional networks controlling spermatogenesis will no doubt be very complex, and thus their elucidation will require a combination of various approaches, including sophisticated analyses of intracellular regulatory pathways and cell‐to‐cell communication pathways. At present, the field has only begun to scratch the surface.
IV. Testis‐Specific Gene Expression and DNA Methylation An age‐old problem in biology is to understand how genes are expressed in a tissue‐specific manner. One solution to this problem is to hypothesize that it results from the existence of tissue‐specific positive‐acting transcription factors. While nature often employs this strategy, it is not the only means
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by which tissue‐specific expression is achieved. A reciprocal strategy is to selectively repress transcription in inappropriate tissues. In some cases, this may be engendered by transcription factors that function as repressors, but this suVers from the requirement for the repressor factors to be expressed at all times in the nonexpressing tissues to eVect permanent transcriptional repression. A potentially more desirable approach to generate relatively permanent transcriptional repression is to methylate promoters, as this simple modification can potentially shut oV transcription without the requirement for constitutive synthesis of repressors (Jaenisch and Bird, 2003; Siegfried and Cedar, 1997). DNA methylation, which occurs on cytosines in a CpG context in vertebrates, has the added advantage that the transcriptionally repressed state is maintained after cell division by the abundant hemimethylase Dnmt1. This enzyme specifically methylates only those cytosines on newly synthesized DNA that correspond to the cytosines methylated on the parental DNA strand (Jaenisch and Bird, 2003; Siegfried and Cedar, 1997). The notion that DNA methylation might be a mechanism responsible for tissue‐specific expression was originally proposed 30 years ago (Holliday and Pugh, 1975; Riggs, 1975). However, it has received surprisingly little support and thus, has been controversial. One stumbling block with this model is the fact that over half of all vertebrate genes, including many tissue‐ specific genes, are rarely methylated (hypomethylated) in any tissue (Jaenisch and Bird, 2003; Siegfried and Cedar, 1997; Walsh and Bestor, 1999). Because many of these constitutively hypomethylated genes display tissue‐ specific expression, this implies that other mechanisms must be used to direct their tissue‐specific expression. Although the mechanism responsible for their refractoriness to methylation is unknown, this class of genes has a unique distinguishing feature; most contain CpG islands (Ariel et al., 1991; Jaenisch and Bird, 2003; Siegfried and Cedar, 1997). This term may appear to imply an overrepresentation of CpG dinucleotides, but actually these islands have a frequency of CpGs similar to that expected by chance alone. Operationally, a CpG island is defined as a ratio of observed CpG to expected CpG of 0.6 in a stretch of DNA with a GC content of >50% and a length of >200 bp (Gardiner‐Garden and Frommer, 1987). In contrast, the non‐CpG‐island portion of the genome has many fewer CpGs than expected by chance alone. This is probably the result of deamination of methylated cytosines to form thymine (Jaenisch and Bird, 2003; Siegfried and Cedar, 1997). What about the tissue‐specific promoters that contain few or no CpG‐rich islands but still contain suYcient CpGs to be potentially subject to control by DNA methylation? It turns out that while many of these do not display the expected correlation between hypomethylation and transcription, the few that do are commonly testis‐specific promoters. Later, we describe
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individual examples of this class of testis‐specific promoters, all of which share the common characteristic of being hypomethylated and expressed in the testis, and methylated and transcriptionally inactive in other tissues. First, we describe genes expressed in male germ cells, then genes expressed in testicular somatic cells such as Sertoli and Leydig cells (Table II). Both low‐ and high‐density CpG promoters selectively hypomethylated and expressed in testis are also discussed. Finally, we evaluate the available evidence supporting the notion that the testis‐specific expression pattern of these promoters is controlled by DNA methylation.
A. Male Germ Cell–Specific Genes One of the first genes shown to be selective hypomethylated in the testis is phosphoglycerate kinase‐2 (Pgk2). Pgk2 is an autosomal ‘‘retroprocessed’’ gene that is intronless as a result of an ancient retrotransposition event. Pgk2 and the highly related, X‐linked, intron‐containing gene Pgk1 display reciprocal expression patterns. Pgk1 is expressed in all cell types except meiotic and postmeiotic germ cells because of the X‐chromosome inactivation event that occurs at meiosis. In contrast, Pgk2 is a low‐density CpG‐containing gene expressed specifically in meiotic and postmeiotic male germ cells. The stage‐specific and cell type–specific promoter of Pgk2 presumably evolved to selectively provide enzyme activity in the cells that do not express its X‐linked sister Pgk1. In precise correspondence with its expression pattern, Pgk2 is hypomethylated in premeiotic, meiotic, and postmeiotic germ cells and hypermethylated in all other tissues and cell types that have been tested (Ariel et al., 1991). Core promoter sequences have been identified in Pgk2 that direct its demethylation in male germ cells prior to its transcriptional activation in vivo (Zhang et al., 1998). Pdha2 is another intronless, X‐linked autosomal gene selectively expressed and hypomethylated in male germ cells (Iannello et al., 1997). Pdha2 encodes the testis‐specific version of the E1 subunit of pyruvate dehydrogenase. Like Pgk2, Pdha2 has a low‐density CpG‐containing small core promoter that confers testis‐specific expression in transgenic mice (187 bp is suYcient for its activity). Methylation‐dependent silencing of Pdha2 is dictated by a crucial CpG in a CRE within its promoter (Iannello et al., 2000). The gene encoding lactate dehydrogenase C (LDH‐C) is another male germ cell–specific gene driven by a small promoter; a 100‐nt fragment is suYcient for testis‐specific expression in vivo. It contains a CpG‐rich promoter that is hypomethylated in germ cells in the testis and methylated in somatic tissues (Bonny and Goldberg, 1995; Kroft et al., 2001). Curiously, many genes expressed from promoters that are selectively hypomethylated and expressed in male germ cells encode DNA‐binding
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proteins. It is not clear whether this is because it represents a real trend or because it is the result of selection bias, as investigators studying DNA‐ binding transcription factors may be more likely to investigate DNA methylation than those that do not. One example of such a DNA‐binding protein is the ALF transcription factor, which is expressed from a male germ cell– specific, CpG‐rich promoter selectively hypomethylated in testis (Xie et al., 2002). ALF (also called TFIIA) is related to the ubiquitously expressed transcription factor TFIIA. Like many other germ cell–specific genes, ALF has a short promoter (133 nt) that is suYcient to confer testis‐specific expression in vivo (Han et al., 2004). Other germ cell–specific hypomethylated genes encode testis‐specific histones and histone‐like proteins. For instance, the testis‐specific histone TH2B is transcribed from an S‐phase‐expressed promoter selectively hypomethylated and expressed in male germ cells. TH2B is first expressed in spermatogonia (germ cells undergoing mitosis) but is most highly expressed in spermatocytes (germ cells undergoing meiosis) (Choi and Chae, 1991). Another example is H1t, a linker histone involved in chromatin remodeling and formation of higher‐order chromatin structure. The promoter driving the expression of H1t is selectively hypomethylated and expressed in pachytene spermatocytes (Singal et al., 2001). A third example is Tnp1, the gene encoding TP1, one of two transition proteins (the other is TP2) that temporarily replace histones during the chromatin compaction phase in spermatids; they are later replaced by protamines, which are responsible for the final stages of chromatin remodeling that occur to form mature spermatozoa. Like Pgk2, Tnp1 is selectively hypomethylated at its 50 (promoter) end but not its 30 end in germ cells in a developmental pattern that precedes its transcriptional induction during spermatogenesis (Trasler et al., 1990). However, the 50 end of Tnp1 is only partially methylated in somatic tissues, so it is not clear whether this would be suYcient to repress transcription in such tissues. In contrast with the solitary Tnp1 gene, the Tnp2 gene and both protamine‐encoding genes (Prn1 and Prn2) are clustered together at a single chromosomal site where they are heavily methylated even in the testis, the sole site of their expression (Choi et al., 1997). As discussed later, it will be intriguing to know the mechanism by which these three genes avoid methylation‐induced transcriptional repression. Most genes that are selectively hypomethylated and expressed in male germ cells are driven by promoters that lack CpG islands, but there are exceptions. One exception is the Magea1 gene, a member of a large gene family encoding intracellular proteins that are normally expressed specifically in testicular germ cells but aberrantly expressed in tumor cells from many diVerent cell lineages (De Smet et al., 1996, 1999). Because of this unusual expression pattern and the observation that some of the MAGE gene products elicit an immune response, they have been classified as members of a growing class
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of proteins called cancer/testis antigens (see also Section III.I) (Van Der Bruggen et al., 2002). Interestingly, the Mage1a gene is hypomethylated not only in germ cells but also in the tumor cells in which it is expressed. This reduced methylation may represent an extreme example of what often occurs to the genome in tumor cells in general (Feinberg and Tycko, 2004). This hypomethylation may be physiologically relevant, as the resultant abberant expression of MAGE tumor antigens could trigger the immune system and thereby suppress tumor formation. However, this heightened immunosurveillance response is balanced by the fact that tumors sometimes have methylated tumor‐suppressor genes, which instead favors their growth (Feinberg and Tycko, 2004). In contrast to all the male germ cell–specific genes discussed so far, whose promoters are all hypomethylated, the gene Tact1 (also known as Act17b) is hypomethylated in its open reading frame (Hisano et al., 2003). This open reading frame‐specific methylation event occurs in a Cp6 island and is testis‐ specific. It appears to be functionally relevant, as is described later. Like the processed Pgk2 and Pdha2 genes described earlier, Tact1 is an intronless gene that presumably is the product of a primordial retrotransposition event to a new chromosomal site. Because this type of event only transposes cDNA sequences and not classical promoter sequences, it will be interesting to know how these three genes acquired male germ cell–specific promoters. It will also be important to examine whether these three genes acquired the property of tissue‐specific methylation as a product of the transposed cDNA sequences, the environment of the chromosomal site to which they were transposed, or a combination of both.
B. Somatic‐Cell Testis Genes In addition to germ‐cell genes, some somatic‐cell testicular genes are also selectively hypomethylated and expressed in the testis. The best‐studied example is the gene Fshr, which is expressed from a low‐density CpG‐ containing promoter specifically expressed in Sertoli cells in male rodents (Griswold and Kim, 2001). All seven CpG residues in the core Fshr promoter are unmethylated in primary rat Sertoli cells, whereas these residues are methylated in all nontesticular tissues that were tested. Other somatic cell– expressed genes that have been shown to be hypomethylated in testis and hypermethylated in nonexpressing tissues are the male‐determining gene Sry (Nishino et al., 2004), the Sertoli cell–expressed gene encoding clusterin (SGP2) (Rosemblit and Chen, 1994), and the Leydig cell–expressed gene encoding 5‐reductase (Reyes et al., 1997). It has been found that the homeobox gene Pem/Rhox5 is another somatic‐ cell gene that undergoes tissue‐specific methylation. Pem/Rhox5 has two
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promoters: a proximal promoter expressed exclusively in Sertoli and caput epididymal principal cells and a distal promoter expressed in somatic cells in the female reproductive tract (Lindsey and Wilkinson, 1996b; Maiti et al., 1996; Rao et al., 2002b; Sutton et al., 1998). Only 0.3 kb of the proximal promoter’s 50 flanking sequence is required to drive the promoter’s androgen‐dependent expression in Sertoli cells in the testis in vivo (Rao et al., 2003). This 0.3‐kb region is unmethylated in adult testis and methylated in tissues that do not express the proximal promoter, including ovary and liver (S. Shanker, M. Rao, and M. F. W., unpublished observations). A sharp methylation boundary exists at the 50 terminus of this 0.3‐kb region, such that the sequences upstream are methylated in the testis. This methylated region extends beyond the other promoter of Pem/Rhox5—the distal promoter—which is 1 kb upstream of the proximal one. A reciprocal methylation pattern is found in the ovary; the distal promoter (active in the ovary) is unmethylated and the downstream proximal promoter (inactive in the ovary) is methylated. It remains for future research to determine which cis elements are responsible for generating these testis‐ and ovary‐specific methylation boundaries, whether these boundaries represent insulator elements, and whether the diVerential methylation of the Pem/Rhox5 promoters is responsible for regulating the gene’s tissue‐specific expression in vivo. C. Tissue‐Specific Hypomethylation: Cause or Consequence of Transcription? There is often a strong correlation between hypomethylation and transcription in the testis, but this does not necessarily mean that DNA methylation is responsible for dictating the testis‐specific expression of the genes previously described. Instead, the selective DNA hypomethylation in the testis could be merely a consequence or a marker of selective transcriptional activation in the testis. One approach to begin to distinguish between these possibilities is to perform time‐course studies. A detailed time‐course analysis of DNA methylation in the Pgk2 gene found that Pgk2 begins to be demethylated in the core promoter and enhancer on day 18.5 p.c.; the demethylation then spreads 1 kb in either direction from the promoter over the next week (Geyer et al., 2004). Pgk2 transcriptional activation is not apparent until 10–12 days after the initial demethylation event, clearly indicating that DNA methylation of the Pgk2 gene cannot be merely a consequence of its transcription. While this study shows that DNA methylation is not a consequence of transcriptional activation in the case of Pgk2, it remains for future studies to determine whether this is the case for other testis‐specific genes.
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Furthermore, time‐course analysis does not address whether the reverse is true—whether DNA methylation has a causal role in controlling the transcription of testis‐specific genes. Work from nontesticular cell types has provided evidence that DNA methylation inhibits transcription by the following non‐mutually exclusive mechanisms: (1) inhibiting the binding of positive transcription factors to the methylated DNA; (2) allowing the binding of methyl‐CpG‐binding factors that actively repress transcription; and (3) altering the chromatin structure in such a way that it is less supportive of transcription (Siegfried and Cedar, 1997). These three mechanisms may often work together to repress gene transcription. In the case of testis‐specific genes, some evidence exists for the first mechanism, whereas the latter two mechanisms have been barely explored. The following details experiments that have been conducted to examine the role of DNA methylation in testis‐specific gene expression. 1. Transfection experiments to determine whether a promoter construct is expressed in cell lines in which the endogenous gene is transcriptionally silent. High expression from the transfected construct implies that transcription factors are not limiting for transcription and that the endogenous gene is not competent to be transcribed for another reason, such as DNA methylation. This has been shown for some testis‐specific genes, including Magea1 and Pdha2 (De Smet et al., 1996; Iannello et al., 1997). 2. Treatment of nonexpressing cells with the DNA‐methylation inhibitor 5‐aza‐20 –deoxycytidine (5‐AzaC). Induction of the endogenous gene in response to 5‐AzaC suggests that the gene is normally silent because it is methylated (although, as described later, 5‐AzaC can act indirectly). The genes encoding ALF, FSHR, and most cancer/testis antigens (including MAGE‐A1) have been shown to be inducible by 5‐AzaC in cell lines (De Smet et al., 1996, 1999; Griswold and Kim, 2001; Xie et al., 2002). This approach has also been used in vivo: ALF expression in mice is induced in the liver after injection with 5‐AzaC (Han et al., 2004). 3. Transfection of in vitro methylated promoter constructs. If DNA methylation represses transcription from the construct, this provides strong evidence that DNA methylation can repress transcription of the gene. This has been demonstrated for many testis‐specific genes, including those encoding Pdha‐2, ALF, Tact1, MAGE‐A1, H1t, TH2B, LDH‐C, and Sry (Choi and Chae, 1991; De Smet et al., 1999; Hisano et al., 2003; Iannello et al., 1997; Kroft et al., 2001; Singal et al., 2001). In most cases, the transfections have been done in nontesticular cell lines, but for Sry the studies were conducted in primary gonadal cells (Nishino et al., 2004). In the case of Tact1, the transfection studies suggested that its open reading frame, not its 50 ‐flanking (promoter) sequences, is the target of methylation‐mediated transcriptional repression (Hisano et al., 2003).
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4. In vitro methylation of key regulatory elements. If in vitro methylation prevents the binding of a transcription factor crucial for transcription, this provides evidence (but not proof) that DNA methylation represses transcription by preventing crucial transcription factors from binding their target sites. An example of a gene inhibited in this manner is Magea1, which has two Ets family consensus sites critical for transcription. When methylated, the Ets sites no longer bind to nuclear proteins, as judged by electrophoretic mobility shift analysis (De Smet et al., 1996). This type of analysis also showed that methylation of the Fshr promoter reduces the binding of factors to an E‐box site known to be essential for maximal Fshr expression in Sertoli cells (Griswold and Kim, 2001). DNase I footprinting analysis has also been used to study factor binding; this approach showed that methylation of an element essential for maximal Pdha2 transcription (an ATF/CRE site) reduces its ability to bind to testicular nuclear factors (Iannello et al., 1997). The promoters of the genes encoding LDH‐C and TH2B were also shown to have reduced nuclear factor binding when methylated in vitro (Choi and Chae, 1991; Kroft et al., 2001). Reciprocally, in some cases, binding studies have revealed that DNA methylation promotes the binding of nuclear factors to crucial cis elements. This has been shown for both the initiator and E2F elements in the Fshr promoter (Griswold and Kim, 2001). It will be interesting to know whether such proteins are methyl‐CpG‐specific binding proteins that possess repressor activity, such as MeCP2 (Jaenisch and Bird, 2003). 5. Analysis of core promoter sequences necessary for testis‐specific expression and whether they also confer tissue‐specific methylation in vivo. This in vivo experiment is particularly valuable for analysis of germ‐cell genes, as no cell lines that fully recapitulate germ‐cell characteristics have been established. This approach was used to demonstrate that less than 200 nt of the Pgk2 and Pdha2 promoters are necessary to provide testis‐specific hypomethylation and expression in transgenic mice (Iannello et al., 1997; Zhang et al., 1998). D. Perspective Although there are advantages to each of the approaches previously described, they also have some disadvantages. For example, gene activation in response to 5‐AzaC does not distinguish between direct and indirect regulation by DNA methylation; that is, 5‐AzaC could demethylate and induce a positive regulator of a testis‐specific gene rather than inducing the testis‐specific gene itself. The assay for the eVect of methylation on the ability of DNA elements to bind to transcription factors in vitro suVers from the fact that these experiments are typically performed with naked DNA and thus may not reflect the eVect of methylation on DNA in its natural state
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(as part of chromatin). Transiently transfecting methylated substrates into cells provides a more physiological context, but most studies test the transcriptional eVect of methylating the entire plasmid, which does not allow elucidation of which particular elements within a promoter are sensitive to methylation. Transfection experiments are particularly troublesome for analyzing germ cell–expressed genes, as no male germ‐cell lines have been established that fully recapitulate normal germ cells. In addition, although transiently transfected DNA constructs are known to form chromatin, this transient, highly amplified form of DNA is unlikely to form the same type of chromatin as does the DNA of endogenous genes. Hence, there is a need to perform more experiments in stably transfected cell lines, as has been done for Magea1 (De Smet et al., 1996, 1999). Another useful approach is to perform experiments in transgenic mice, as this provides expression data in an in vivo context. However, analysis of transgenic mice containing diVerent promoter constructs is time‐consuming and often generates correlative rather than mechanistic data. What other approaches can be used to address the role of DNA methylation in tissue‐specific expression? One potentially powerful approach is to map testis‐specific demethylation elements and then to ablate them to determine whether this prevents tissue‐specific transcription. This approach will indicate whether a link exists between testis‐specific demethylation and transcription. In addition, by identifying the sites of the specific elements responsible, the approach will also provide a strong foundation for ultimately determining the underlying molecular mechanism, including the nature of the factors that bind to the elements. In the case of genes expressed in somatic cells in the testis, initial analysis can be done in transfected cell lines, but later results would need to be confirmed in transgenic mice. As an ultimate physiological test, ‘‘knock‐in’’ mice could be made in which the demethylation signal is mutated in the endogenous gene. A key aspect of this approach is to perform additional experiments, such as time‐course analysis, to provide further evidence of whether an element mapped by this approach is directly responsible for triggering hypomethylation that subsequently leads to transcriptional activation, or whether the inverse is true. A second approach is to individually mutate the CpGs in crucial regulatory elements to assess which of these CpGs are responsible for repressing transcription in nontesticular tissues. Good candidates would be those CpGs that are conserved in promoter and enhancer elements from diVerent species. This CpG‐mutation approach has been used to identify the specific cytosine residues responsible for repressing transcription of the gene encoding the cytokine interleukin‐2 in noninduced T lymphocytes (Bruniquel and Schwartz, 2003). It has not yet been applied to testis‐specific genes. A third approach is to methylate specific cytosines in promoter constructs in vitro and then transfect them into cell lines to determine whether
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transcription is inhibited. This site‐specific CpG methylation approach has identified specific cytosines that must be methylated to allow inducible expression of the gene encoding brain‐derived neurotrophic factor (Martinowich et al., 2003). Again, this approach has not yet been used to examine the regulation of testis‐specific genes. Once the specific regulatory elements involved in DNA methylation have been identified, it becomes crucial to identify the factors involved in the regulation. The sequence of the elements identified by the analyses just described may be useful toward this goal but will not provide a definitive answer, as some transcriptional elements have the potential to bind a large number of transcription factors. The use of antibodies against specific transcription factors in conjunction with traditional in vitro DNA‐binding assays such as electrophoretic mobility shift analysis will provide an initial answer, but the results should be confirmed in intact cells by chromatin immunoprecipitation analysis. In addition, gain‐of‐function experiments (e.g., transfection of expression plasmids) complemented with loss‐of‐function experiments (e.g., RNA interference and expression of dominant‐negative inhibitors) must be conducted to determine whether the transcription factors implicated by the aforementioned assays actually regulate the transcription and methylation of the testis‐specific gene in question. In cases in which DNA methylation does not inhibit factor binding but instead recruits a new factor, this suggests the involvement of a methyl‐CpG‐ binding protein. The DNA‐binding and chromatin immunoprecipitation assays previously described will determine which particular ones are recruited; currently known examples include MeCP2, MBD1, MBD2, MBD3, MBD4, and KAISO, all of which repress transcription (Jaenisch and Bird, 2003). Gain‐of‐function and loss‐of‐function cotransfection experiments will provide evidence of the particular factors responsible for repressing the transcription of the gene in question. Recapitulation of DNA methylation‐mediated transcriptional repression of testis‐specific gene in vitro provides a potentially powerful approach for ultimately elucidating the underlying molecular mechanism responsible for this regulation in vivo. This may be feasible, as conditions for generating transcriptionally competent chromatin in vitro have been elucidated (Fyodorov and Kadonaga, 2003). Another issue that requires future scrutiny is how some testis‐specific genes are transcribed despite being heavily methylated. As previously described, the Tnp2‐Prn1‐Prn2 gene cluster exhibits this characteristic (Choi et al., 1997). Another example is the human cyclin A gene CCNA1, which is methylated in germ cells in the testis, the site of its preferential expression (Muller‐Tidow et al., 2001). Conversely, CCNA1 is hypomethylated in many tissues that do not express it. This lack of correlation between the methylation and expression patterns of CCNA1 strongly suggests that it is not
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regulated by methylation. Further evidence comes from work on its mouse ortholog, Ccna1, which is exclusively expressed in testis. A 1.3‐kb region of the Ccna1 50 ‐flanking sequence is suYcient to drive its testis‐specific expression (predominantly in pachytene spermatocytes) in transgenic mice containing an enhanced green fluorescent protein reporter gene (Muller et al., 2000). However, the transgene was hypomethylated in many tissues that did not express it, indicating that methylation alone cannot be responsible for preventing the expression of Ccna1 in nontesticular tissues. Further evidence for this is that Ccna1 transcription is not induced by 5‐AzaC in nonexpressing cell lines, whereas it is induced by the histone deacetylase inhibitor trichostatin A (Muller et al., 2000). It is not known how the CCNA1 gene and the Tnp2‐Prn1‐Prn2 gene cluster are transcribed in testis despite their promoters being highly methylated. One possibility is that despite the overall methylation of these genes, CpGs in key regulatory elements in their promoters or enhancers are not methylated. Another possibility is that their transcription depends on transcription factors such as Sp1 whose DNA binding can be impervious to DNA methylation (Zhu et al., 2003). Alternatively, they may have evolved mechanisms to avoid recruiting or activating methyl‐CpG‐binding repressor proteins. Lastly, it is possible that male germ cells are less sensitive to DNA methylation‐mediated repression in general; relevant to this possibility is the finding that the otherwise constitutively expressed methyl‐ CpG‐binding repressor protein MeCP2 is expressed at only trace levels in the testis (Meehan et al., 1992). How commonly is DNA methylation used to dictate tissue‐specific expression? Has this mode of gene regulation been co‐opted almost exclusively by some germ cell–specific genes? Or is it a mechanism also used by some somatic cell–specific genes? It certainly makes sense that this mechanism would be used by germ cell–expressed genes, as such genes would be hypomethylated in germ cells and thus would avoid losing CpGs in their promoters as a result of deamination over evolutionary time (as only germ‐cell genes are passed on to progeny) (Jaenisch and Bird, 2003; Siegfried and Cedar, 1997). Thus, by being impervious to this CpG‐loss mechanism, germ cell–specific genes have the unique advantage of being able to maintain their ability to be regulated by DNA methylation. In contrast, a somatic cell– specific gene methylated in all nonexpressing cell types would, by definition, have a promoter highly methylated in germ cells. This would lead to a potentially unstable situation in which its promoter would have a tendency to progressively lose cytosines over future generations. Thus, for a somatic cell–specific gene to remain regulated by DNA methylation, it must somehow prevent (or at least reduce) this loss of CpGs. We predict that one category of such somatic cell–specific genes is one in which DNA methylation control was so quickly established over evolutionary time that few
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CpGs were lost by deamination. Once established, it is hypothesized that the crucial CpGs required for this tissue‐specific expression would have been maintained by strong selection pressure. A second category of somatic cell– specific genes that would escape loss of CpGs crucial for methylation regulation is ones that are hypomethylated not only in a particular somatic‐cell type but also in germ cells. By being hypomethylated in germ cells, these genes will avoid losing CpGs over evolutionary time. A potential example of this type of gene is the Pem/Rhox5 homeobox gene, which is specifically expressed in Sertoli cells in the testis but is also hypomethylated in testicular germ cells (Lindsey and Wilkinson, 1996b; S. Shanker and M. F. W., unpublished observations). In conclusion, many testis‐specific genes exhibit a striking association between hypomethylation and transcription, making them potentially useful for understanding how tissue‐specific gene expression is achieved. A causal link likely exists between these two phenomena, but it remains to be definitively determined whether demethylation allows transcription or whether activation of transcription elicits demethylation. Growing evidence supports the former scenario for some testis‐specific genes, including time‐course, in vitro methylation, and in vivo demethylation studies. Further analysis by these approaches, along with gene‐ and element‐targeting approaches in mice, should provide a complete picture of whether DNA methylation regulates the transcription of testis‐specific genes and how this is achieved at the molecular level.
Acknowledgments We thank many people for providing experimental results prior to publication and giving us input on writing this review, including Bob Braun, Jaideep Chaudhary, Charles De Smet, Erv Goldberg, Larry Jameson, John McCarrey, Carsten Mu¨ ller, Roy Parker, Philippa Saunders, Michael Skinner, Jacquetta Trasler, Robert Viger, William Walker, Debra Wolgemuth, and David Zarkower. We also thank Pierrette Lo for assistance in preparation of this manuscript. This work was supported by National Institutes of Health grants HD045595, HD042714, and CA16672.
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Modeling Age‐Related Diseases in Drosophila: Can this Fly? Kinga Michno,*,{ Diana van de Hoef,*,{ Hong Wu,* and Gabrielle L. Boulianne*,{ *Program in Developmental Biology, The Hospital for Sick Children, Toronto, Ontario Canada M5G 1X8 { Department of Molecular and Medical Genetics, University of Toronto, Toronto, Ontario, Canada M5G 1X8
I. II. III. IV. V. VI. VII. VIII.
The Challenge of Studying Age‐Related Diseases Why Study Human Neurodegenerative Diseases in Drosophila? Huntington’s Disease and Polyglutamine Disorders Parkinson’s Disease Alzheimer’s Disease Amyotrophic Lateral Sclerosis What Can We Do with a Fly Model of Neurodegenerative Disease? The Next Frontier Acknowledgments References
Human neurodegenerative diseases are characterized by progressive neuronal cell loss often resulting in memory and cognitive decline, motor dysfunction, and ultimately premature death. Despite the prevalence of these diseases, there are no eVective cures. Insight into many of these syndromes has come from the identification of single gene mutations that are associated with inherited forms of the disease. This has led to the development of animal models in which the pathogenesis caused by these genes can be rigorously examined. Due to their short life span and powerful genetic potential, several attempts have been made to model neurodegenerative diseases in the fruit fly Drosophila melanogaster. This review will describe how these models were generated and how faithfully they recapitulate human disease. In addition, how fly models can be used to identify genetic modifiers of known disease genes and what these have revealed about the biochemical pathways underlying disease pathogenesis is discussed. Finally, the review will describe how fly models can be used to identify new therapeutic targets and test the eVectiveness of new drugs. ß 2005, Elsevier Inc.
Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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I. The Challenge of Studying Age‐Related Diseases During the past few decades, advances in modern medicine have led to dramatic improvements in human health and, as a result, a commensurate increase in human life expectancy. An unexpected consequence of this however, has been an increase in the number of age‐related diseases and in particular, neurodegenerative diseases such as Alzheimer’s disease. In fact, studies estimate that 4.5 million Americans are currently aZicted with Alzheimer’s disease and that this number will exceed 11 million by 2050 (Hebert et al., 2003). Similar increases in the number of individuals aZicted with other neurodegenerative diseases have also been predicted. Despite their prevalence and the huge emotional and financial burdens imposed by these diseases, no eVective therapies exist. This therapeutic void in part reflects an incomplete understanding of the biochemical pathogenesis of these diseases. One approach that has been successful in revealing the mechanisms underlying many neurodegenerative diseases has been to identify genes that are mutated in familial forms of the disease. However, mutations in these genes often account for less than 5% of all disease cases, while the remaining 95% are sporadic and late onset. The low frequency of familial forms and the late onset of sporadic forms suggests that neurodegenerative diseases are complex, and that other genetic and environmental factors must play additional roles as causative or risk factors in these various disorders. In addition, the mechanisms by which mutations in these genes lead to pathogenesis remain unclear. A powerful method that can be used to understand the basic mechanisms underlying neurodegenerative diseases is to generate animal models based on manipulating the expression of single genes that are causative in disease. This approach has been facilitated by the fact that many neurodegenerative diseases are inherited as autosomal dominant traits, and therefore, expression of the mutant gene in a model organism might be expected to recapitulate the disease. In fact, vertebrates, and in particular mice, have provided many useful models of human neurodegenerative diseases. However, genetic analysis in mice is still slow as compared to simpler organisms like the nematode Caenorhabditis elegans or the fruit fly Drosophila melanogaster. But is it possible to recapitulate a complex human disorder in such a simple model system?
II. Why Study Human Neurodegenerative Diseases in Drosophila? There are numerous features of Drosophila that make it particularly amenable for studying human neurodegenerative diseases. First, Drosophila is a genetically tractable organism with a generation time of approximately 10
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days and a short life span of 60–80 days. This makes it an attractive model to study aging and age‐related diseases. Drosophila also has a relatively small genome (1=20 of human) contained on four chromosomes that have been completely sequenced. Importantly, comparative genome analysis reveals that approximately 75% of human disease genes have a Drosophila ortholog (Fortini et al., 2000; Reiter et al., 2001). Many methods are also available to manipulate genes in Drosophila using either loss‐of‐function mutations or RNA‐interference techniques to study the normal function of a gene or by generating transgenics to examine the eVect of misexpressing wild‐type or mutant genes. Importantly, these ectopic expression studies are greatly enhanced by the availability of tissue‐specific promoters. Genes that are thought to be vital to the survival of the organism can also be studied using the GAL4/UAS system (Brand and Dormand, 1995; Brand and Perrimon, 1993). Briefly, the GAL4 system consists of two independent components. The first component consists of GAL4‐enhancer trap lines that express the yeast transcriptional activator, GAL4, in a tissue‐ or cell‐specific manner. The second component consists of transgenic lines in which a transgene is expressed under the transcriptional control of the GAL4 target sequence, the upstream activating sequence (UAS), which is inactive in flies lacking GAL4. The progeny of a cross between these two lines, which now contain both the UAS transgene and the GAL4, then express the transgene in a specific pattern. The advantages of using this system are: (1) it allows us to target the expression of transgenes to a variety of specific cell types at any developmental stage; and (2) lethal eVects of expressing the transgenes are circumvented. This method has been particularly useful to study the eVect of expressing genes implicated in neurodegenerative diseases given that expression of these genes might be expected to cause progressive neuronal cell loss and eventually premature organismal death. The anatomy and development of Drosophila has also been extensively characterized and many reagents are available to determine the eVect of mutations on specific cell/tissue types. Drosophila also has a relatively complex nervous system and many markers are available to identify neuronal cells at various developmental time points and to monitor neuronal cell death. Methods to measure neuronal function and in particular synaptic transmission are also available. At present, the most widely used preparation is the Drosophila neuromuscular junction (NMJ), which has been particularly useful for revealing morphological and functional defects associated with mutations or misexpression of genes that aVect synaptic morphology and neurotransmission. Similar methods are also rapidly being developed to measure synaptic transmission at central synapses. In addition, methods to measure intracellular calcium dynamics within this preparation have also been developed (Macleod et al., 2002). Together these tools make it possible to determine whether the expression of mutant genes implicated in
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neurodegenerative diseases give rise to early defects in synaptic transmission in addition to progressive neuronal cell loss. Drosophila also exhibits complex behaviors, which can be monitored relatively easily as can the eVect of specific genes on learning and memory. Taken together, these features make it possible to examine the progressive nature of neurodegenerative diseases from potential early eVects on synaptic function, leading to memory and cognitive decline, and ultimately to neuronal cell death. Perhaps one of the major strengths of a Drosophila model is the ability to perform large‐scale genetic screens to identify modifiers of known disease‐ causing genes. These modifiers not only provide insight into the biological functions of disease‐causing genes during normal development but also provide clues as to how they contribute to disease pathogenesis. More importantly, genetic modifiers are themselves excellent candidates for additional risk or causative factors in disease and can provide novel targets for therapeutic intervention. Finally, Drosophila models of human disease may also be useful to screen for novel therapeutic drugs. Currently, screening for drugs and therapeutics takes place sequentially with the first steps carried out in cell‐free or cultured cell models. Although these models may reflect some aspect of the disease, they do not recapitulate what occurs in humans and are of limited use. To further validate the potential therapeutic value of these compounds, they are then evaluated in animal models where the in vivo eYcacy of the trial compounds can be determined. The majority of compounds are often abandoned at this stage because they fail to ameliorate the disease process in vivo or exhibit harmful side eVects. Altogether, the process is slow and expensive and contributes to the time and overall cost of developing new drugs. This could proceed more rapidly and be more cost eVective if drugs could be screened using invertebrate models of human disease. Although there are few examples of drug screening using Drosophila as a model, previous studies have shown that the peptidomimetric FTI‐254, which inhibits farnesyl protein transferase, can suppress dominant phenotypes generated in the Drosophila eye as a result of expression of a mutant Ras oncogene. These early studies suggested that Drosophila could be used to screen for drugs that inhibit Ras‐dependent cell transformation and therefore might function as anticancer agents in humans (KauVmann et al., 1995). Similarly, fly models of epilepsy have been used to test the eYcacy of known antiepileptic drugs. These studies have shown that the antiepileptic drugs gabapentin, phenytoin, valproate, and potassium bromide were all able to suppress seizure behavior in flies (Kuebler and Tanouye, 2002; Reynolds et al., 2003; Tan et al., 2004). These studies suggest that Drosophila models of disease may provide powerful methods to test the eVectiveness of known compounds and also screen for new therapeutic drugs.
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Taken together, there are many reasons why invertebrate models such as Drosophila oVer unique opportunities to study the biological mechanisms underlying human neurodegenerative diseases. Of course, this depends entirely on the ability to generate a model in flies that faithfully reproduces the human condition. Several attempts have been made to generate invertebrate models of several human neurodegenerative diseases (Table I). Some of these have met with promising results while others have failed to recapitulate essential features of the disease.
III. Huntington’s Disease and Polyglutamine Disorders Huntington’s disease (HD) is a dominant, late‐onset neurodegenerative disease caused by expansion of a CAG triplet repeat within the Huntingtin (Htt) protein (Group, 1993). Htt is a large 350‐kDa protein that is ubiquitously expressed (Li et al., 1993; Strong et al., 1993) and normally carries a repeat of between 8 and 25 glutamines in the N‐terminus of the protein. Individuals that have 35 or fewer copies of the CAG repeat are asymptomatic, whereas individuals carrying greater than 40 repeats develop disease (Bates et al., 2002; Gusella and MacDonald, 1995) and those with greater than 65 repeats develop a juvenile form of disease (Penney et al., 1997). In addition to HD, several other neurodegenerative diseases have been shown to be caused by expanded CAG repeats that encode expanded polyglutamine repeats that are collectively referred to as polyQ diseases (Gusella and MacDonald, 2000; Paulson, 1999; Perutz, 1999; Price et al., 1998a). The key features of HD and polyQ diseases are that they are late onset and display progressive neurodegeneration that is associated with memory loss, cognitive defects, and, often, movement disorders. As in HD, the age of onset for polyQ diseases correlates with the length of the CAG repeat with a sharp threshold separating disease from nondisease states. A major pathological hallmark of HD and polyQ diseases is the presence of visible protein aggregates or inclusions (Davies et al., 1997; DiFiglia et al., 1997). In addition to aggregated mutant polyQ protein, these inclusions often contain many other cellular proteins, including chaperones, transcriptional coactivators, proteasome subunits, ubiquitin, and in particular CBP, an acetyl transferase (Kazantsev et al., 1999; Nucifora et al., 2001; SteVan et al., 2000). During the past several years, models of HD and polyQ diseases have been generated in Drosophila (Marsh et al., 2003). The most common approach has been to engineer flies that express either a wild‐type gene or a gene that encodes various numbers of CAG repeats under the transcriptional control of the GAL4/UAS system. Typically, two diVerent GAL4 drivers have been utilized to drive expression of a wild‐type or mutant transgene. Elav‐GAL4
Table I
Invertebrate Models of Human Neurodegenerative Diseases
Human Pathology PolyQ disorders Disease gene homologue Movement deficits Neuronal degeneration Protein aggregation Premature death Parkinson’s disease Disease gene homologue Movement deficits Neuronal degeneration Protein aggregation Premature death Alzheimer’s disease Disease gene homologue Learning deficits
Drosophila Models
C. elegans Models
HD, SCAs (Rubin, 2000)
MJD1, SCAs (Rubin, 2000)
Larval (Lee, 2004) adult (Marsh, 2003) locomotory deficits Eye (Marsh, 2000; Warrick, 1998) sensory bristles (Marsh, 2000) Nuclear (Warrick, 1998) cytoplasmic (Lee, 2004) Yes (Warrick, 1998; Marsh, 2000; Lee, 2004)
Progressive loss of motility (Morley, 2002)
PARK2, UCHL1 (Rubin, 2000)
PARK2, UCHL1 (Rubin, 2000)
Locomotor deficits (Feany, 2000; Greene, 2003) Dopaminergic cell loss (Feany, 2000; Yang, 2003) Cytosolic inclusions (Feany, 2000)
Motor deficits (Lakso, 2003)
Sensory neurons (Faber, 1999) Cytoplasmic (Satyal, 2000) perinuclear (Parker, 2001) ND
Dopaminergic cell loss (Lakso, 2003) Intracellular inclusions (Lakso, 2003)
Yes in parkin null mutants (Greene, 2003)
ND
PS, APP, TAU (Rubin, 2000)
PS, APP, TAU (Rubin, 2000)
Induced by A misexpression (Greeve, 2004; Iijima, 2004)
ND
Neuronal degeneration Protein aggregation Premature death Amyotrophic lateral sclerosis Disease gene homologue Movement deficits Neuronal degeneration Oxidative stress Protein aggregation Premature death
Yes, age‐dependent (Wittmann, 2001; Greeve, 2004; Iijima, 2004) A plaques (Greeve, 2004; Iijima, 2004) & NFTs (Jackson, 2002) Yes (Wittmann, 2001; Greeve, 2004; Iijima, 2004)
Yes, induced by tau misexpression (Kraemer, 2003) A, fibrillar deposits in muscle (Link, 2001) Yes with tau misexpression (Kraemer, 2003)
SOD1 (Rubin, 2000)
SOD1 (Rubin, 2000)
Yes for SOD1 null (Mockett, 2003) & dEAAT LOF (Rival, 2004) Yes but not motor neurons (Phillips, 1995; Rival, 2004) Sensitivity in SOD1 null (Phillips, 1989) dEAAT LOF (Rival, 2004) ND
ND
Yes for SOD1 null (Mockett,2003) & dEAAT LOF (Rival, 2004)
ND Vulnerability in fALS‐SOD1 transgenic (Oeda, 2001) Concomitant with oxidative stress (Oeda, 2001) ND
The purpose of this table is to provide a brief comparison between two invertebrate models and is not comprehensive. For an extensive outline of human disease genes suited to analysis in Drosophila, see Bier, 2005. For full reference information, see corresponding bibliography. A, beta‐amyloid; APP, amyloid precursor protein; dEAAT1, Drosophila excitatory amino acid transporter; FALS, familial amyotrophic lateral sclerosis; HD, Huntington’s disease; LOF, loss of function; MJD1, Machado‐Joseph disease; ND, not determined; NTF, neurofibrillary tangles; PARK, Parkinson; PolyQ, polyglutamine; PS, presenilin; SCA, spinocerebellar ataxia; SOD1, superoxide dismutase 1; UCHL1, ubiquitin‐C‐terminal hydrolase.
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drives expression of the transgene in every cell of the nervous system from embryogenesis onwards (Robinow and White, 1988) and is therefore widely used to examine the eVect of neuronal expression of any given transgene. Gmr‐GAL4 is expressed in all cells of the eye including both neurons and support cells (Ellis et al., 1993). The major advantage of this driver is that the eye is not required for viability and any toxic eVect of the transgene can be easily observed in the adult as a rough eye phenotype. Importantly, flies that express mutant polyQ proteins display a neurodegenerative phenotype that, like humans, depends on the length of the polyQ repeat and shows similar thresholds of repeat lengths to those that cause human disease (Jackson et al., 1998; Marsh et al., 2000; Takeyama et al., 2002; Warrick et al., 1998). As in humans, neurodegeneration in the fly models is late onset, progressive, and ultimately results in premature death (Jackson et al., 1998; Marsh et al., 2000; Warrick et al., 1998). In addition, transgenic flies expressing mutant polyQ proteins also accumulate protein aggregates. Thus, it seems possible to recapitulate the key features of HD and other polyQ diseases in flies. Similar neurodegenerative phenotypes have also been shown in C. elegans expressing mutant polyQ proteins (Morley et al., 2002), suggesting that the mechanisms leading to neurodegeneration are conserved between species. The ability to generate a fly model of HD has also made it possible to investigate the mechanisms by which mutant proteins lead to pathogenesis. In particular, it has been possible to test the hypothesis that the sequestration of critical proteins within nuclear inclusions in HD can contribute to the pathogenesis of the disease. As mentioned earlier, a prominent component of nuclear inclusions are acetylases such as CBP. If sequestration of acetylases contributes to HD pathology, then inhibiting deacetylases using either genetic or pharmacological methods might be expected to compensate and suppress the HD phenotype in Drosophila. In fact, not only did inhibition of deacetylases suppress the Drosophila HD phenotype (Hughes, 2002; SteVan et al., 2001), subsequent studies have shown that it is also eVective in mammals (Hockly et al., 2003). Other major components of inclusions are molecular chaperones and various components of the ubiquitination machinery, suggesting that protein processing and degradation may also be compromised in the disease state. If so, then overexpression of chaperones might be able to suppress the neurodegeneration induced by polyQ mutant proteins in flies. In fact, chaperones have been identified in several genetic screens as suppressors of many polyQ disease phenotypes in flies (Fernandez‐Funez et al., 2000; Kazemi‐Esfarjani and Benzer, 2000; Warrick et al., 1999). Similarly, modulation of the ubiquitin or SUMO pathways also modifies disease pathology (SteVan et al., 2004). Finally, these fly models of HD and polyQ diseases have made it possible to test the potential eVectiveness of specific drugs. In particular, the
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eVectiveness of compounds that might block or disrupt aggregate formation have been evaluated in cell‐free systems, cell‐culture models, and in vivo in a fly model of polyQ disease (Apostol et al., 2003). In all cases, two compounds, cystamine and Congo red, suppressed the pathology associated with expression of the mutant protein. These studies demonstrate the potential that fly models have to reveal the mechanisms underlying disease and to identify and evaluate new therapeutic interventions.
IV. Parkinson’s Disease Parkinson’s disease (PD) is the second most common human neurodegenerative disorder. Like most other neurodegenerative disorders, PD is mostly sporadic and typically aVects individuals between 50 and 60 years of age. The disease is associated with selective death of dopaminergic neurons resulting in a movement disorder characterized by muscle rigidity and resting tremor. In addition to the observed degeneration of specific brain regions, patients also display two neuropathological hallmarks called Lewy bodies and Lewy neurites, which are ubiquitin‐containing protein inclusions adjacent to the nucleus and in neuronal processes. As with many other neurodegenerative diseases, the etiology is unclear. The best characterized form of familial Parkinson’s disease has been linked to mutations in ‐synuclein (Kruger et al., 1998; Polymeropoulos et al., 1997), a small, phosphorylated protein that localizes to presynaptic nerve terminals. At present the precise mechanism by which ‐synuclein leads to PD is unclear; however, ‐synuclein has been found to form insoluble filaments under defined conditions. Moreover, mutations in ‐synuclein that are associated with PD display a higher rate of insoluble filament assembly compared to wild‐type protein, suggesting that protein aggregates composed of these filaments contribute to disease pathology. Importantly, ‐synuclein has been shown to be a central component of Lewy bodies in patients with both familial and sporadic PD (Spillantini et al., 1997). Therefore, understanding the mechanism by which ‐synuclein leads to the degeneration of dopaminergic neurons may shed light on all forms of PD. Not surprisingly, the first attempts to generate a Drosophila model of PD were based on generating transgenic flies that expressed either wild‐ type or mutant forms of ‐synuclein in neurons using the GAL4/UAS system (Feany and Bender, 2000). Because there is no obvious ortholog of ‐synuclein in Drosophila, these studies focused on expressing the human gene in flies. Interestingly, expression of ‐synuclein within all neurons did not aVect overall brain morphology or lead to overall degeneration. Rather a specific and progressive loss of dopaminergic neurons was observed regardless of whether flies were expressing wild‐type or mutant protein,
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suggesting that high levels of ‐synuclein are responsible for the disease phenotype. Other aspects of the disease, however, were only observed in flies expressing mutant protein including cytosolic inclusions reminiscent of Lewy bodies and locomotory defects (Feany and Bender, 2000). Similar eVects of expressing mutant ‐synuclein were also observed in C. elegans. The eVects of expressing wild‐type and mutant ‐synuclein in the Drosophila eye were also examined. As was observed in the adult brain, expression of mutant ‐synuclein in the eye gave rise to progressive degeneration of photoreceptor neurons. Several other genes have also been identified that are causal of parkinsonism. Many of these encode proteins involved in ubiquitin‐ and proteasome‐ mediated protein degradation implicating the ubiquitin machinery in the etiology of PD. Mutations in parkin, which encodes an E3‐ubiquitin ligase (Shimura et al., 2000), account for most forms of juvenile and early onset PD. In addition, mutations have also been found in ubiquitin C‐terminal hydrolase L1 (UCHL1) (Wintermeyer et al., 2000), which encodes a thiol protease that hydrolyzes C‐terminal bonds between ubiquitin and small adducts. At present, the mechanism by which mutations in these genes lead to selective loss of dopaminergic neurons is still unknown. In an attempt to generate a new model of PD in Drosophila, loss‐of‐ function mutations in Drosophila parkin were generated. Mutant flies had reduced longevity and exhibited defects in flight and climbing activity. The defects in motor function appear to be due to an apoptotic muscle degeneration that is associated with prominent mitochondrial swelling and disruption of the inner membrane (Greene et al., 2003). Although this does not resemble PD, mitochondrial dysfunction has been implicated in the progression of the disease. More recently, studies have shown that pan‐neuronal expression of the Parkin substrate Pael‐R causes age‐dependent selective degeneration of Drosophila dopaminergic (DA) neurons (Yang et al., 2003). Moreover, coexpression of Parkin and Pael‐R causes degradation of Pael‐R and suppresses its toxicity, whereas loss‐of‐function mutations in Parkin promote Pael‐R accumulation and augment its toxicity. Finally, overexpression of Parkin can mitigate ‐synuclein‐induced neuritic pathology and suppress its toxicity. Taken together, these studies suggest that Parkin may be a central player in PD and that the mechanisms by which mutations in Parkin lead to pathology can be studied in a Drosophila model.
V. Alzheimer’s Disease Alzheimer’s disease (AD) is the most common cause of dementia in the aging population. The disease is characterized by two neuropathological hallmarks: neurofibrillary tangles (NFTs) and extracellular amyloid plaques
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(Probst et al., 1991; Selkoe, 1994; Yankner, 1996). NFTs consist of paired helical filaments composed primarily of the abnormally phosphorylated, microtubule‐associated tau protein (Goedert et al., 1988, 1992; Greenberg and Davies, 1990; Kosik et al., 1988; Lee et al., 1991). Amyloid plaques (also called senile plaques) can be divided into two categories, ‘‘diVuse’’ and ‘‘compact,’’ and these can be distinguished both morphologically and with the use of histochemical stains. Both types of plaques are composed primarily of ‐amyloid (A) peptide deposits that are produced as a result of the processing of a larger precursor called the amyloid beta protein precursor (APP) (Kang et al., 1987). DiVuse plaques tend to be amorphous and are not associated with significant cell loss, whereas compact or dense plaques can be associated with neuronal cell loss. Although much eVort has gone into understanding the composition of plaques and tangles, their role in the etiology of the disease remains controversial. An alternative approach to identify causative factors in AD has been to search for cases that are inherited as an autosomal dominant trait. This search has led to the identification of three human genes, APP, presenilin 1, and presenilin 2 (PS1 and PS2), for which in‐frame, missense mutations have been associated with early onset Familial AD (FAD). Together mutations in APP and PSs account for greater than 50% of FAD, with PS1 mutations being the most common. APP encodes a type I integral membrane protein that is initially cleaved by proteases known as ‐ and ‐secretase to give rise to an N‐terminal extracellular fragment and a membrane tethered C‐terminal fragment. PSs encode multipass transmembrane proteins that function as part of a multimolecular complex called ‐secretase (reviewed in Sisodia and St. George‐Hyslop, 2002), which cleaves single‐pass transmembrane proteins including Notch and APP. In the case of APP, ‐secretase cleaves the membrane‐bound C‐terminal fragment resulting in the release of secreted A peptides and the APP intracellular domain referred to as the AICD. FAD mutations in PSs appear to cause elevated levels of total ‐amyloid peptide (A 1–40 and A1–42) or specifically elevate the levels of the longer form of the peptide (A 1–42), which is thought to form amyloid plaques in the brains of patients with AD and has been widely suggested as a primary cause for AD‐related neurodegeneration (Price et al., 1998b). Although less is known about the function of the AICD, studies have shown that it can translocate to the nucleus and activate transcription of downstream targets including tetraspanin KAI1/CD82 that is known to modulate cytoskeletal function. Not surprisingly, attempts to create a Drosophila model of AD have focused on generating transgenic flies that express various forms of PS or APP with the goal of observing age‐dependent accumulation of amyloid plaques and, as a result, neuronal degeneration. Unlike vertebrates or C. elegans, there is a single PS homologue in Drosophila (Boulianne et al.,
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1997). Null mutations in Drosophila psn give rise to a Notch‐like phenotype as was observed in C. elegans and mice, indicating that in addition to a high degree of sequence homology, the function of PS is also conserved between species (Guo et al., 1999; Struhl and Greenwald, 1999; Ye et al., 1999). Despite the similarities, however, misexpression of either wild‐type or mutant PS does not give rise to AD‐like pathology. Rather, the phenotypes observed resemble those of Notch loss‐of‐function mutants, suggesting that the phenotype is due to dominant negative eVects of psn (Ye and Fortini, 1999). Interestingly, while misexpression of FAD mutations in psn also gave rise to Notch‐like phenotypes, they were less severe than those observed using a wild‐type psn transgene, suggesting that the FAD mutations in PS are loss‐of‐function. These flies may therefore provide a useful model to determine how loss‐of‐psn function may contribute to AD pathology. In addition to models based on PS expression, several attempts have been made to generate AD models based on manipulating APP in flies. Importantly, while APP homologs are found in Drosophila and C. elegans, they do not contain sequences homologous to human A peptide. Therefore, models based on expression of invertebrate forms of APP are unlikely to recapitulate the essential pathological features of AD. However, they can provide valuable insight into the normal function of APP, which in vertebrates has proven elusive. These studies revealed that null mutations in Drosophila appl are viable but give rise to defects in phototaxis that can be rescued by Drosophila or human APP (Luo et al., 1992). More recently, further analysis of appl mutants has suggested that APPL may also function as a vesicular kinesin 1 receptor and play a role in axonal transport (Gunawardena and Goldstein, 2001; Torroja et al., 1999). To generate a Drosophila model of AD that will provide insight into the mechanisms by which A might lead to toxicity in AD, transgenic flies have been generated that misexpress either a wild‐type human APP transgene or one containing mutations associated with FAD. These studies have demonstrated that overexpression of a wild‐type human APP transgene that included the A domain resulted in the production of A40, suggesting that Drosophila have some aspects of the APP‐processing machinery (Fossgreen et al., 1998). In addition, increases in cell death were also observed within the larval brain that depended on the presence of the A region and the carboxy‐ terminal domain of APP, which is thought to bind to kinesin 1. However, amyloid plaques were not observed. In contrast, misexpression of the longer, more toxic A42 peptide in the Drosophila brain resulted in diVuse plaques, age‐dependent learning defects, and extensive neurodegeneration that ultimately resulted in reduced life span (Finelli et al., 2004; Greeve et al., 2004; Iijima et al., 2004). While similar age‐dependent learning defects were also
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observed in transgenic flies expressing the less toxic A40 peptide, this was not associated with either amyloid plaques or neurodegeneration. Of note, the amyloid plaques observed in flies expressing A42 were diVuse in contrast to the dense core plaques regularly observed in patients with AD. At present, it is unclear whether this reflects the fact that flies do not live long enough to develop dense cored plaques or that they are lacking essential cofactors. Nonetheless, flies do exhibit both learning defects and neurodegeneration in the absence of dense cored plaques, suggesting that, although these may correlate with disease pathology, they may not be essential for all of the associated pathologies. Interestingly, the phenotype of flies expressing A peptides could be ameliorated by secretase inhibitors or by mutations in neprilysin, a metalloprotease that is thought to play a role in the catabolism of A peptides in vivo. This observation suggests that it may be possible to use these fly models to screen for drugs that are of therapeutic value in AD. Finally, several attempts have also been made to generate a fly model of AD based on misexpression of tau, the primary component of NFTs, which are a hallmark of AD and are prominent in many other neurodegenerative diseases. For example, mutations in tau have been found in families with inherited frontotemporal dementia with parkinsonism linked to chromosome 17 (FTDP‐17), suggesting that tau dysfunction contributes to neurodegeneration (Hutton et al., 1998). Encouragingly, when either wild‐type or mutant tau was expressed in transgenic flies, several key features of human neurodegenerative disease were observed including neuronal accumulation of phosphorylated tau, age‐dependent neuronal degeneration, and premature death (Wittmann et al., 2001). However, NFTs did not develop. In contrast, if tau is overexpressed along with GSK‐3, a protein kinase known to phosphorylate tau, NFTs were observed, suggesting that the limiting factor when generating NFTs are the levels of tau phosphorylation (Jackson et al., 2002). Consistent with this model, protein kinases were the most frequent class of molecules identified as enhancers in a modifier screen of a Drosophila model of tauopathy (Shulman and Feany, 2003). In addition, studies have shown that the kinase PAR‐1 can enhance tau toxicity by triggering a phosphorylation process that involves phosphorylation of tau at S262 and S356 (Nishimura et al., 2004). Furthermore, phosphorylation by PAR‐1 is a prerequisite for subsequent phosphorylation of tau by GSK‐3 and Cdk‐5. Together, the action of these kinases may lead to the high levels of tau phosphorylation necessary to form NFTs. Interestingly, PSs containing FAD mutations bind avidly to GSK‐3 and this may enhance the ability of the kinase to phosphorylate tau. Therefore, in AD, PS mutations may contribute to the formation of NFTs by accelerating the ability of protein kinases to phosphorylate tau.
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VI. Amyotrophic Lateral Sclerosis Not all attempts to generate a fly model of a neurodegenerative disease have succeeded. Cases in point are studies aimed to develop a Drosophila model of amyotrophic lateral sclerosis (ALS). ALS is a progressive and ultimately fatal neurodegenerative disease that selectively targets motor neurons. Most ALS cases occur sporadically (sALS), but 5–10% are inherited in an autosomal dominant fashion referred to as familial ALS (fALS), which is indistinguishable from sALS from a neuropathological perspective. All ALS patients succumb to progressive paralysis but are spared from any apparent cognitive decline. Because there is currently no cure for ALS, identifying the unique molecular events that result in selective motor neuron death has been the subject of intense research. Several gene mutations have been identified for fALS cases, leading to the implication of several toxic cellular processes in promoting motor neuron degeneration, including protein aggregation (Johnston et al., 2000; Watanabe et al., 2001), oxidative stress (Yim et al., 1996), and glutamate excitoxicity (Rothstein et al., 1992). The most intensely studied fALS‐associated gene is superoxide dismutase 1 (SOD1). Mutations in SOD1 account for approximately 20% of fALS cases and over 100 diVerent fALS‐associated mutations have been identified (Rosen et al., 1993). SOD1 is an antioxidant enzyme that catalyzes the dismutation of superoxide free radicals. Since increased oxidative damage has been observed in ALS patients, this suggested that the mechanism by which fALS associated mutations in SOD1 gives rise to motor neuron dysfunction is due to increased oxidative damage. However, most fALS mutations in SOD1 do not aVect the dismutase activity of the enzyme but appear to confer a toxic dominant gain‐of‐function eVect. Consistent with this model, mice harboring loss‐of‐function mutations in SOD1 are viable and do not display any motor neuron disease (Reaume et al., 1996). In contrast, transgenic mice that express SOD1 containing fALS‐associated mutations develop a motor neuron syndrome that bears many similarities with human ALS (Bruijn et al., 1998; Gurney et al., 1994; Wong et al., 1995). Although the nature by which SOD1 mutations give rise to toxic eVects is still unclear, mutant SOD1 enzyme has been shown to exhibit increased peroxidase activity (Wiedau‐ Pazos et al., 1996). In addition, mutant SOD1 also appears to form insoluble aggregates that have also been observed within inclusion bodies of fALS patients. Surprisingly, similar attempts to recapitulate motor neuron disease by expressing fALS‐associated mutations in transgenic flies have failed despite the fact that Drosophila contains a SOD1 gene that shows a high degree of sequence homology with its vertebrate counterparts. As was observed in mice, loss‐of‐function mutations in Drosophila SOD1 do not give rise to
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motor neuron disease. However, unlike mutant mice, which are viable, Drosophila SOD1 mutants exhibit a severely reduced life span (7–8 days) as compared to wild‐type and display enhanced sensitivity to oxidative stress (Phillips et al., 1989). The short life span can be rescued by introducing a wild‐type Drosophila, bovine (Reveillaud et al., 1994), or human SOD1 gene (Parkes et al., 1998), demonstrating that, in addition to sequence homology, Drosophila and vertebrate SOD1 genes are also functionally homologous. Interestingly, expression of human SOD1 exclusively within motor neurons can rescue the short life span of SOD1‐null flies and increase the life span of wild‐type flies by more than 40% (Parkes et al., 1998). In both cases, the increase in life span is associated with increased resistance to oxidative stress. These studies suggest that although SOD1 is a ubiquitously expressed enzyme, its function is particularly critical in motor neurons. This is interesting because these cells are the precise targets of ALS. However, neither motor neuron‐specific, nor ubiquitous expression of human fALS‐SOD1 alleles gives rise to dominant gain‐of‐function fALS‐like phenotypes in flies (Elia et al., 1999; Mockett et al., 2003). In fact, both ubiquitous and motor neuron expression of fALS alleles of SOD1 partially rescues the life span of SOD1‐ null flies. While the precise mechanism is unknown, the ability of the mutant SOD1 to rescue life span is likely due to the positive eVects of reintroducing dismutase activity. Hence, while both Drosophila SOD1 mutants and human fALS‐SOD1 mutants give rise to recessive phenotypes in flies, inducing dominant gain‐of‐function eVects has not been successful. Interestingly, similar attempts to establish a model of ALS in the nematode C. elegans have been far more successful. Specifically, ubiquitous expression of human fALS‐SOD1 alleles in C. elegans using a heat shock promoter gave rise to worms with elevated sensitivity to oxidative stress (Oeda et al., 2001). Following muscle‐specific fALS‐SOD1 expression, oxidative stress promoted aggregate formation concomitant with impairments in the degradation of fALS‐SOD1 protein. Notably, in the absence of oxidative stress, no phenotypes were detected, which the authors attribute to the short life span of the organism. In future studies it would be interesting to see whether other ubiquitous drivers produce similar ALS‐like phenotypes in worms given that heat shock itself is known to have negative eVects on longevity. The obvious question is why does expression of fALS‐SOD1 give rise to dominant ALS‐like phenotypes in mice and worms but not in flies? One possibility is that only a subset of fALS mutations in SOD1 can confer motor neuron dysfunction in invertebrates. However, similar SOD1 mutations were used in both the fly and worm studies previously described. In addition, studies have expressed a variety of fALS alleles in transgenic flies and none have given rise to any notable pathology. Another likely possibility is that motor neuron dysfunction in model organisms only occurs when
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mutant SOD1 is expressed at very high levels. Consistent with this hypothesis, studies in transgenic mice have shown that the level of expression of the fALS‐SOD1 transgene is critical with respect to onset of symptoms, such that founder mice with low levels of transgene expression are reported to be asymptomatic. This point is particularly important when taking into consideration the fact that ubiquitous expression of fALS‐SOD1 in flies was most often accomplished using the endogenous SOD1 promoter to drive expression of a single copy transgene in the context of the GAL4‐UAS system and this may not result in suYciently high levels of SOD1 expression. In the worm model, an inducible heat shock promoter was used to induce transgene expression from multicopy extrachromosomal arrays. It is important to bear in mind that in humans, a single mutant SOD1 allele is suYcient to generate motor neuron degeneration. The failure to do so in animal models may be due to the fact that these organisms do not live long enough to display symptoms unless the transgenes are expressed at high levels to compensate. Hence, future attempts to generate a Drosophila model of fALS may require the use of stronger promoters or the presence of multiple copies of the transgene. Alternatively, it may be that ubiquitous or even motor neuron expression of fALS‐SOD1 in flies is not suYcient to cause motor neuron degeneration. In fact, motor neuron expression of fALS‐SOD in transgenic mice also failed to induce ALS‐like phenotypes (Lino et al., 2002), suggesting that SOD1 expression in other cell types plays a role in motor neuron dysfunction. Glial cells are excellent candidates as they have also been implicated in ALS pathology. Specifically, deficits in glutamate uptake by glial glutamate transporters can lead to glutamate excitotoxicity, which is believed to promote motor neuron cell death (Rothstein et al., 1995). While the role of glial cells in motor neuron disease has not yet been investigated in flies, inactivating the function of the Drosophila glutamate transporter, dEAAT1, by RNA interference gives rise to characteristic ALS pathology including decreased longevity, increased sensitivity to oxidative stress, locomotory defects, mitochondrial damage, as well as neuronal cell death (Rival et al., 2004). Although progressive motor neuron degeneration is not observed, this study has produced a model that, in future studies, could significantly further our understanding of the role of glutamate excitoxicity in promoting neuronal degeneration. Finally, it remains possible that the discrepancy between flies and worms reflects an inherent diVerence between the sensitivity of these two organisms to specific forms of stress. Alternatively, flies may be more resistant to SOD1 aggregate formation. Abnormal protein folding and degradation has previously been implicated in ALS pathology and inclusion bodies form in both fALS and sALS patients as well as in mouse ALS models. Some of these inclusions stain positively for ubiquitin and molecular chaperones. Adult
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flies are known to express high levels of various chaperones including small heat shock proteins. As such, it would be interesting to determine whether expression of fALS‐SOD1 in a background in which the levels of specific chaperones are reduced is suYcient to induce motor neuron disease. These studies would provide further insight into the mechanisms by which mutant SOD1 can lead to motor neuron dysfunction and provide an additional model for studying this disease.
VII. What Can We Do with a Fly Model of Neurodegenerative Disease? There are numerous advantages to having a genetically tractable model of a human neurodegenerative disease. First and foremost is the ability to perform large‐scale genetic screens to identify genes that, when mutated, can enhance or suppress the disease phenotype in the fly. Although it is possible to identify modifier loci in mice and humans using a candidate gene approach, it is currently impossible to carry out systematic, large‐scale genetic screens to identify new modifier loci in vertebrates. In contrast, such modifier screens are common in invertebrates such as Drosophila. In the past, several such screens have been performed in Drosophila and C. elegans to identify modifiers of Huntington’s, polyQ, and PD disease genes (Driscoll and Gerstbrein, 2003; Hughes, 2002). Because aggregate formation is a common feature of most human neurodegenerative diseases, it is perhaps not surprising that chaperones were often identified in these screens. However, modifiers have also been identified that reveal mechanisms specific to each disease including histone deacetylases (polyQ diseases) and various kinases (PD). Because Drosophila models of AD have only recently been developed, less is known about the nature of any A modifiers. However, a neprilysin homolog was identified in a genetic screen for modifiers of A‐induced toxicity in Drosophila (Finelli et al., 2004). Neprilysin is the prototypical member of the M13 family of zinc‐dependent metalloproteases. In vertebrates, Neprilysin is expressed in the brain and can hydrolyze neuropeptides at synapses (Turner et al., 2001). More recently, Neprilysin has been shown to degrade both intracellular and extracellular A peptides in vivo when Neprilysin transgenic mice were crossed to APP transgenic mice, and the progeny exhibited a marked reduction in amyloid plaques (Leissring et al., 2003). Taken together, these studies suggest that proteases such as Neprilysin may be of therapeutic value in AD. Moreover, the fact that Neprilysin was identified in a genetic screen in Drosophila suggests that this model may be useful in identifying additional factors involved in A toxicity and AD.
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Drosophila models may also be useful to identify and evaluate potential therapeutic drugs. While such studies are commonly performed using mouse models, they are time‐consuming, labor intensive, and costly. In contrast, it is easy and inexpensive to maintain large populations of flies. Therapeutic drug treatment of flies with neurological disorders such as epilepsy has also been extensively described. More recently, studies have shown that pharmacological reduction of HDAC can reduce pathology in flies as well as in HD and related diseases. As further models of disease are developed, it should be possible to screen for drugs that not only are eVective in one disease, but may also be of value in other neurodegenerative disorders.
VIII. The Next Frontier Despite the relative success in developing fly models of neurodegenerative diseases, several important challenges remain. For example, in many instances, neurodegeneration in fly models occurs regardless of whether a wild‐type or mutant protein is expressed. While the severity or progression of the phenotype increases when the mutant rather than the wild‐type protein is expressed, it remains possible that the toxicity observed in the fly is simply due to overexpressing a given protein. This is a particular concern when the mutant protein expressed is not normally found in flies, as is the case for A. Even in the case in which a fly model has been established based on expression of a highly conserved gene, it is not always possible to fully recapitulate all of the features of the disease (Table I). For example, it has been diYcult to generate a fly that exhibits NFTs. However, similar diYculties have been encountered when generating mouse models. Whether this is due to the short life span of these organisms as compared to humans remains unclear. Alternatively, some of the biochemical pathways required to initiate disease pathology may not be conserved. Another major challenge will be to demonstrate neuronal specificity in any fly model of neurodegenerative disease. To date, many of the fly models were developed by expressing mutant proteins in the fly eye and examining the eVects on photoreceptor degeneration. However, in humans, often only selective populations of neurons are aVected in any given disease. Whether this level of specificity can be achieved in a fly model remains unclear but would be useful in understanding why some neuronal populations are more susceptible to disease, whereas others are relatively unaVected. Finally, although drug discovery and testing may be a distinct possibility in fly models, successful use of a drug in flies may not be a predictor of eYcacy in human clinical trials. While there are many advantages to fly models, there are also important diVerences in their metabolism, complexity of their nervous systems, and life span that might preclude the observation of potentially noxious eVects.
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Nevertheless, Drosophila models could provide the basis for the development of high‐throughput screens of drugs and chemical libraries that could then be further characterized and validated in vertebrate models and finally, by clinical trials. Clearly, Drosophila models of neurodegenerative diseases are rapidly making significant contributions to understanding the biochemical mechanisms underlying pathogenesis. Over time it will be interesting to see whether they also provide venues for novel therapeutic interventions.
Acknowledgments We thank Dr. William Trimble for critically reading the manuscript and various members of the Boulianne lab for helpful discussions. K.M. is the recipient of a Canada Research Scholarship. D.vdH was partially supported by a University of Toronto Open Scholarship 2001 and the OSOTF Scace graduate fellowship in Alzheimer’s disease. H.W. was supported by a CIHR fellowship award and received a Postdoctoral Fellowship, in part, through the Hospital for Sick Children Research Training Centre. G.L.B. is a CIHR Investigator.
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Warrick, J. M., Chan, H. Y. E., Gray‐Board, G. L., Chai, Y., Paulson, H. L., and Bonini, N. M. (1999). Suppression of polyglutamine‐mediated neurodegeneration in Drosophila by the molecular chaperone HSP70. Nat. Genet. 23, 425–428. Warrick, J. M., Paulson, H. L., Gray‐Board, G. L., Bui, Q. T., Fischbach, K.‐F., Pittman, R. N., and Bonini, N. M.wat (1998). Expanded polyglutamine protein forms nuclear inclusions and causes neural degeneration in Drosophila. Cell 93, 939–949. Watanabe, M., Dykes‐Hoberg, M., Culotta, V. C., Price, D. L., Wong, P. C., and Rothstein, J. D. (2001). Histological evidence of protein aggregation in mutant SOD1 transgenic mice and in amyotrophic lateral sclerosis neural tissues. Neurobiol. Dis. 8, 933–941. Wiedau‐Pazos, M., Goto, J. J., Rabizadeh, S., Gralla, E. B., Roe, J. A., Lee, M. K., Valentine, J. S., and Bredesen, D. E. (1996). Altered reactivity of superoxide dismutase in Familial Amyotrophic Lateral Sclerosis. Science 271, 515–518. Wintermeyer, P., Kruger, R., Kuhn, W., Muller, T., Woitalla, D., Berg, D., Becker, G., Leroy, E., Polymeropoulos, M. H., Berger, K., Przuntek, H., Schols, L., Epplen, J. T., and Riess, O. (2000). Mutation analysis and association studies of the UCHL1 gene in German Parkinson’s disease patients. Neuro.Rep. 11, 2079–2082. Wittmann, C. W., Wszolek, M. F., Shulman, J. M., Salvaterra, P. M., Lewis, J., Hutton, M., and Feany, M. B. (2001). Tauopathy in Drosophila: Neurodegeneration without neurofibrillary tangles. Science 293, 711–714. Wong, P. C., Pardo, C. A., Borchelt, D. R., Lee, M. K., Copeland, N. G., Jenkins, N. A., Sisodia, S. S., Cleveland, D. W., and Price, D. L. (1995). An adverse property of a Familial ALS‐linked SOD1 mutation causes motor neuron disease characterized by vacuolar degeneration of mitochondria. Neuron 14, 1105–1116. Yang, Y., Nishimura, I., Imai, Y., Takahashi, R., and Lu, B. (2003). Parkin suppresses dopaminergic neuron‐selective neurotoxicity induced by Pael‐R in Drosophila. Neuron 37, 911–924. Yankner, B. A. (1996). Mechanisms of neuronal degeneration in Alzheimer’s disease. Neuron 16, 921–932. Ye, Y., and Fortini, M. E. (1999). Apoptotic activities of wild‐type and Alzheimer’s disease‐ related mutant presenilins in Drosophila melanogaster. J. Cell Biol. 146, 1351–1364. Ye, Y., Lukinova, N., and Fortini, M. E. (1999). Neurogenic phenotypes and altered Notch processing in Drosophila presenilin mutants. Nature 398, 525–529. Yim, M. B., Kang, J.‐H., Yim, H.‐S., Kwak, H.‐S., Chock, P. B., and Stadtman, E. R. (1996). A gain‐of‐function of an amyotrophic lateral sclerosis‐associated Cu,Zn‐superoxide dismutase mutant: An enhancement of free radical formation due to a decrease in Km for hydrogen peroxide. Proc. Natl. Acad. Sci. USA 93, 5709–5714.
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Cell Death and Organ Development in Plants Hilary J. Rogers School of Biosciences, CardiV University, CardiV United Kingdom CF10 3TL
I. Introduction II. Seed and Embryo Development A. Cell Death in the Endosperm B. Cell Death in the Embryo and Suspensor III. Leaf, Stem, and Root Development A. Leaf Sculpting B. Leaf Senescence C. Tracheary Element DiVerentiation D. PCD in Aerenchyma Formation E. Cell Death in the Root Cap IV. Flowering and Reproduction A. Sexual Organ Abortion B. PCD in the Male Sexual Organs C. PCD in the Female Gametophyte D. PCD During Self‐Incompatibility Interactions E. Petal Senescence and Cell Death V. Conclusions and Future Directions Acknowledgments References
Programmed cell death (PCD) is an important feature of plant development; however, the mechanisms responsible for its regulation in plants are far less well understood than those operating in animals. In this review data from a wide variety of plant PCD systems is analyzed to compare what is known about the underlying mechanisms. Although senescence is clearly an important part of plant development, only what is known about PCD during senescence is dealt with here. In each PCD system the extracellular and intracellular signals triggering PCD are considered and both cytological and molecular data are discussed to determine whether a unique model for plant PCD can be derived. In the majority of cases reviewed, PCD is accompanied by the formation of a large vacuole, which ruptures to release hydrolytic enzymes that degrade the cell contents, although this model is clearly not universal. DNA degradation and the activation of proteases is also common to most plant PCD systems, where Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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they have been studied; however, breakdown of DNA into nucleosomal units (DNA laddering) is not observed in all systems. Caspase‐like activity has also been reported in several systems, but the extent to which it is a necessary feature of all plant PCD has not yet been established. The trigger for tonoplast rupture is not fully understood, although active oxygen species (AOS) have been implicated in several systems. In two systems, self incompatibility and tapetal breakdown as a result of cytoplasmic male sterility, there is convincing evidence for the involvement of mitochondria including release of cytochrome c. However, in other systems, the role of the mitochondrion is not clear‐cut. How cells surrounding the cell undergoing PCD protect themselves against death is also discussed as well as whether there is a link between the eventual fate of the cell corpse and the mechanism of its death. ß 2005, Elsevier Inc.
I. Introduction Programmed cell death (PCD) is now recognized as an important event in shaping plant organs (Jones and Dangl, 1996; Pennell and Lamb, 1997). In animal cells PCD has been defined as cell death that is part of the normal life cycle of the organism, is triggered by specific physiological signals, and involves de novo gene transcription (Ellis et al., 1991). This distinction is important in diVerentiating developmental PCD from necrotic cell death that may occur as a result of injury or exposure to toxic substances. In plants an important response to pathogenic attack is the hypersensitive response (HR) in which selected cells around the site of infection undergo PCD to prevent spread of the pathogen. Although the cellular features of HR‐PCD show many similarities to developmental PCD, this area has been reviewed (Greenberg and Yao, 2004) and will not be dealt with in detail here. In plants it is also important to distinguish between PCD and senescence, terms that have led to much controversy in this field and have been the focus of review (van Doorn and Woltering, 2004). In this review, PCD will be used to indicate cellular death as opposed to the death of whole organs or individuals. In some plant organs the eVect of PCD is particularly evident. Thus, although in animals, sculpting of the fingers is often quoted as the classic example of the action of PCD in organ development, in plants an obvious example is the prominent holes in the leaves of the house plant Monstera (Melville and Wrigley, 1969). However, the phenomenon is much more widespread, and examples are found at all stages of plant development in which the targeted removal of specific cells or cell types is a requirement for normal development.
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In animal cells at least two forms of PCD have been described: apoptosis and autophagy. A number of cytological features describe apoptosis. These include nuclear condensation and marginalization, chromatin condensation, followed by fragmentation of DNA into nucleosomal units known as DNA laddering, and the formation of membrane inclusions known as apoptotic bodies. In animals the apoptosed cell remains are finally engulfed by neighboring cells through phagocytosis and the cell corpse disappears (Cohen, 1993). In a few cases PCD may be defined as truly autonomous as, for example, in the very tightly regulated cell death in Caenorhabditis elegans (Yuan and Horvitz, 1990). However, in many cases, signals external to the cell, such as changes in hormone levels, trigger PCD (Cohen, 1993). Once PCD has been triggered (for an overview of animal PCD, see Krishnamurthy et al., 2000), a complex network of regulators is switched on involving increases in cytosolic calcium concentrations, an oxidative burst, and release of pro‐PCD factors such as cytochrome c from the mitochondrion. Release of cytochrome c is regulated by a growing family of Bcl‐2 proteins that interact with the mitochondrial membrane, facilitating or inhibiting its release. These intracellular events activate a family of cysteine aspartate specific proteases known as caspases, which are both regulators and the eVectors of cell death. Caspases act on a plethora of targets initiating cell condensation, nuclear fragmentation, and DNA breakdown. The characteristic DNA laddering occurs as a result of cleavage at the nucleosome‐linker sites, by DNAases, which are activated both by the caspases and more directly by increases in cytosolic calcium levels (Peitsch et al., 1993). Autophagy in mammalian cells (reviewed by Stromhaug and Klionsky, 2001) was originally associated with a response to starvation. It is characterized by the formation of vesicles containing proteins and organelles, which are transported to the lysosome. Known as autophagosomes, these vesicles have a short half‐life (in the range of a few minutes) and their contents are then digested by the hydrolase‐packed lysosomes to generate monomeric building blocks. The signaling pathways leading to autophagy are much less well‐defined than those involved in apoptosis; however, heterotrimeric G proteins (Ogier‐Denis et al., 2000) and type III phosphoinositide 3‐phosphates (Petiot et al., 2000) have been implicated. Interestingly, autophagy and apoptosis may not be completely independent mechanisms. Beclin 1, which interacts with Bcl‐2 family proteins, is associated with type III phosphoinositide 3‐phosphates and may have a role in sorting proteins destined for the autophagosome (Kihara et al., 2001). Also, sphingosine, known to activate apoptosis, increases the activity of lysosomal proteases such as cathepsin B, which in turn are caspase activators (Ferri and Kroemer, 2001). Notably the lysosomes appear to act upstream of the mitochondria, suggesting a regulatory role in apoptosis (Yuan et al., 2002).
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Few genes with sequence homology to those involved in animal PCD have been identified in plants despite the availability of genome sequences from representatives of the major taxonomic divisions of flowering plants, namely Arabidopsis representing dicotyledonous plants (The Arabidopsis Genome Initiative, 2000) and rice representing the monocotyledons (GoV et al., 2002). Putative homologues include those for Bax‐Inhibitor‐1, an inhibitor of the proapoptotic Bcl‐2 family member Bax (Kawai et al., 1999) involved in apoptosis, Beclin (Laporte et al., 2004; C. WagstaV, unpublished results) involved in autophagy, and dad‐1 (Orza´ ez and Granell, 1997), although doubt has been cast on the regulatory role of dad‐1 (Kelleher and Gilmore, 1997). Although homologues of caspases have not been identified from the genome sequences (Lam and Del Pozo, 2000), plant metacaspases members of a related superfamily are found in plant genomes (Lam, 2004) and can trigger apoptosis‐like cell death in yeast (Watanabe and Lam, 2005). In addition, caspase inhibitors inhibit plant PCD in several systems (see following). Thus, whether homologues to the other components of the apoptotic or autophagic machinery in animal cells are present in plants but with insuYcient homology to be identifiable through database searching, or whether the whole mechanism of cell death is diVerent in plants remains as yet unresolved. A number of features diVerentiate plant and animal cells and these may account for the divergence in cell death mechanisms, one major diVerence being the cell wall. However, parallels to apoptosis are evident at a cytological level in some types of plant PCD and proteases are prominently upregulated in both plants and animals. Hence, a mechanism may emerge for plant PCD that is divergent from that in animals but shares some common features. As examples of PCD are examined from diverse developmental processes questions about the input signals triggering PCD, the process of cell death, and the consequences of the death of the cell can be asked.
II. Seed and Embryo Development Several seed tissues undergo PCD as part of their normal development, and best studied of these is the endosperm. The endosperm is a transient nutritive triploid tissue formed from the secondary fertilization event characteristic of higher plants. It is essential for embryo development, but endosperm life span is limited and cell death progresses until in some species all the cells in the tissue are dead as the seed reaches maturity. The life span of the endosperm is species specific; in dicotyledonous seeds such as beans, it is barely detectable and the nutrients are transferred to the embryo before the seed reaches maturity. In cereal seeds, however, the endosperm is a prominent component and a layer of the endosperm, the aleurone layer, is still
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viable at seed maturity. Aleurone cells, however, are nondividing, and the whole tissue disappears following seed germination. The aleurone has a secretory role, producing the hydrolytic enzymes required to break down the reserves stored in the endosperm, and is tightly regulated through hormonal control (Huttly and Phillips, 1995). Both have been intensely studied, due to their importance for seed development and also as models for plant PCD (Young and Gallie, 2000a). Another well‐studied seed tissue with limited life span is the suspensor. This is composed of a string of cells connecting the embryo to the rest of the seed and acts as a conduit for the nutrients mobilized from the endosperm to reach the developing embryo (Yeung and Meinke, 1993). Once this function has been performed, its cells undergo PCD. Suspensor cell death has proven to be a particularly fruitful tissue for PCD studies in that it is possible to induce embryo formation from isolated cells in culture and to manipulate the formation and degeneration of the suspensor cells.
A. Cell Death in the Endosperm In seeds with little or no endosperm, the lack of this tissue is due either to arrested development or to its early degeneration (Young and Gallie, 2000a). Thus, although the endosperm cells undergo cell death in all species, the timing of this event and the size of the tissue vary. Although the fate of the endosperm in noncereals is interesting, most studies have focused on cereal endosperm, especially in wheat and maize. Development and cell death in both species is not uniform, and the pattern of cellular change may be related to species‐specific signaling events (Young and Gallie, 2000b). 1. The Aleurone PCD in the aleurone is stimulated by Gibberellic acids (GAs), a class of plant growth regulators (Bethke et al., 1999; Kuo et al., 1996; Wang et al., 1996), and the process can be simulated by the incubation of isolated aleurone layers, or protoplasts with GA (Fath et al., 2000). In vivo GAs are produced by the embryo and stimulate the aleurone layer to release hydrolytic enzymes that make available nutrients stored in the starchy endosperm for embryo growth (Fincher, 1989). The hydrolases are synthesized de novo from the breakdown of storage proteins found in protein storage vacuoles (PSVs) within the aleurone cells. During GA stimulation, the PSVs coalesce to form a single large vacuole and change from storage to lytic organelles. This is accompanied by acidification of the PSVs (Swanson and Jones, 1996) and accumulation of hydrolytic enzymes. Although at first these are required for
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storage protein mobilization, later they are implicated in death of the aleurone cell through autolysis and removal of the aleurone cell corpse. One possible mechanism for this autolysis is that the acidification activates specific proteases, which initiate a protease cascade within the PSVs (Fath et al., 2000). The protease phytepsin is upregulated following GA treatment in aleurone cells (Bethke et al., 1996) and was shown to process probarley lectin in roots (Sarkkinen et al., 1992). Thus, protease activation by specific proteases may be an important mechanism in aleurone PCD. However, although protease cascades are reminiscent of the well‐characterized caspase cascades in animal PCD (Salvesen and Dixit, 1999), caspases are unlikely to play an important role here. Caspases, which accumulate at the right time to be involved in PCD, have been identified in aleurone cells, but they are not GA‐induced (Fath et al., 2000). Furthermore, apoptotic bodies as seen in apoptotic animal cells have not been observed (Fath et al., 2000), so GA‐induced PCD in aleurone cells diverges significantly from animal cell apoptosis. The final step of aleurone cell PCD is loss of integrity of the plasma membrane (Fath et al., 2000, 2002) leading to loss of turgor and shrinkage of the cell corpse. AOS may play the final role in the execution of aleurone PCD by causing membrane lipid peroxidation resulting in loss of membrane integrity (Fath et al., 2001). There is indeed a body of evidence implicating AOS in aleurone PCD. GA‐treated protoplasts become more sensitive to exogenous hydrogen peroxide, and incubation of GA treated aleurone protoplasts with antioxidants such as butylated hydroxy toluene (BHT) reduced mortality (Fath et al., 2002). The same study also revealed that both transcripts and activities of enzymes involved in AOS metabolism decrease on treatment of aleurone protoplasts with GA. The source of AOS was also investigated, and both an increase in glyoxylate activity and a change in the electron transport chain away from the alternative oxidase pathway were associated with GA treatment. Both of these are associated with increased AOS production. Thus, both an increase in AOS production and a reduction in AOS metabolism are associated with PCD following GA treatment, and membrane damage leading to PCD that may occur when AOS production exceeds removal of AOS by metabolic routes. Nitric oxide (NO) may also be involved as a signaling molecule and/or with an antioxidant role. NO can act as an antioxidant reducing membrane damage by interacting with lipid peroxyl radicals, or it can promote cell death (Beligni et al., 2002). Incubation of barley aleurone cells with a NO scavenger, 2‐(4‐carboxyphenyl)‐4,4,5,5‐tetramethylimidazoline‐1‐oxyl‐3‐oxide (cPTIO), accelerated GA‐induced PCD, suggesting that endogenous NO was retarding PCD. This was supported by the detection of endogenous NO production using NO‐sensitive fluorescent dyes (Beligni et al., 2002), although there was no apparent diVerence in NO production between GA or cells treated with another plant growth regulator abscisic acid (ABA), which protects
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endosperm cells from PCD (see later). Exogenous application of NO also retarded PCD in GA‐treated cells in a similar way to BHT, suggesting an antioxidant role, and NO also delayed the downregulation of AOS metabolism gene expression. While aleurone cells become highly vacuolated, degradation of their nuclear DNA is detectable. It proceeds rapidly and they lose over 50% of their nuclear DNA within 5 days after GA treatment. However, although TUNEL staining is positive indicating the presence of free 30 OH ends, controversy exists over whether DNA laddering takes place. One study (Fath et al., 1999) did not find DNA laddering in barley aleurone protoplasts, whereas more recently DNA laddering was detected in preparations from both wheat and maize intact aleurone cells (Domı´nguez et al., 2004). Whether there is interspecific divergence in the mechanism for nuclear DNA degradation in aleurone cells, or whether the contrasting results are due to diVering methodologies remains unclear. Where it has been detected, the timing of DNA laddering is in agreement with the TUNEL data, coinciding with extensive vacuolation. Candidate nucleases have been identified; at least three nucleases are induced by GA treatment, and their induction is delayed by ABA treatment. The timing of their activity fits in with the data on DNA degradation. Thus, strong upregulation of their activity occurs after hydrolase release is essentially complete (Fath et al., 2000). It would seem, moreover, that the nuclease activity and consequent DNA degradation are a requirement for PCD in these cells. Hydrolase release, nuclease activity, and DNA degradation were all inhibited by the guanylyl‐cyclase inhibitor LY83583 (Bethke et al., 1999; Fath et al., 1999), indicating that cGMP may also play a regulatory role in aleurone cell PCD. The link between hydrolase release and cell death is not, however, simple because at least in the earlier stages of hydrolase release, it can be uncoupled from PCD. When aleurone cells are treated with ABA, cell death is delayed even if the cells have previously been treated with GA and induction of hydrolase release has begun (Fath et al., 2000). Furthermore, in aleurone protoplasts, accumulation of proteases in the medium does not result in PCD. Thus, freshly prepared protoplasts were incubated in medium that had been recovered from GA‐treated cells and that was rich in hydrolases; however, this did not accelerate PCD in the protoplasts (Fath et al., 2000). Ca2þ and protein phosphorylation have been implicated in the signal transduction mechanism that leads to GA‐induced PCD. Okadaic acid, a protein phosphatase inhibitor, blocked GA‐induced cytosolic Ca2þ increase (Kuo et al., 1996) and microinjection of a syntide‐2, synthetic substrate for calmodulin and Ca2þ‐ dependent protein kinases, inhibited the development of the large vacuole, an essential precursor of PCD in the aleurone cells (Ritchie and Gilroy, 1998).
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2. The Starchy Endosperm Unlike the aleurone, the starchy endosperm cells are dead by the time the seed reaches maturity. Researchers (Gallie and Young, 2004) suggested that the death of these cells may be necessary to allow access for the hydrolases, which break down the starch reserves, making the nutrients available to the embryo. Cell death of the starchy endosperm is thus unusual in that the corpse of the dead cell is not eliminated and persists in the developed seed. Cell death in this tissue needs to be correctly timed such that starch filling is complete before PCD is initiated. Moreover, the starch filling is not synchronous across the whole tissue; it occurs in a wave starting in the central/ proximal region of the seed and moving toward the pedicel end, and PCD follows in the same pattern (Young and Gallie, 2000a). Thus, in close proximity the starchy endosperm cells are dying while the aleurone cells and the embryo remain living, suggesting a programmed coordination of signals. The plant‐growth regulator ethylene plays an important role in regulating PCD in the starchy endosperm both spatially and temporally. Evidence comes from the use of inhibitors of ethylene synthesis such as AVG ([2‐aminoethoxyvinyl]glycine) and of ethylene action such as 1‐MCP (1‐methylcyclopropane), both of which delay PCD, whereas applications of exogenous ethylene accelerate PCD in this tissue (Young et al., 1997). Two peaks of ethylene production are detected in the endosperm coinciding with waves of PCD. The first is in the central/proximal region of the seed, while the second late peak occurs at the pedicel end of the seed (Young et al., 1997). Evidence for involvement of ethylene in endosperm PCD also comes from starch‐deficient mutants. In the maize shrunken 2 mutant endosperm, PCD occurs prematurely (Young et al., 1997) and ethylene levels are elevated coinciding with the wave of PCD. However, to achieve correct timing of PCD in diVerent cells, and furthermore to protect the aleurone and the embryo from the ethylene signal, the cells must either be exposed diVerentially or sense the same ethylene levels diVerentially. Researchers (Gallie and Young, 2004) have isolated cDNAs both for ethylene biosynthetic genes and ethylene receptor genes. They have shown that the expression of ethylene receptors is much lower in the endosperm compared to the embryo, suggesting that the endosperm tissue is responsive to lower levels of the plant growth regulator compared to the embryo. This fits in with other experiments showing that if ethylene levels are suYciently high, then the embryo cells will also enter PCD (Young et al., 1997). As well as ethylene, ABA may also be important in the regulation of PCD in the starchy endosperm. ABA is important during the desiccation of the seed as it reaches maturity. Evidence comes from two mutants in which the endosperm PCD occurs prematurely (Young and Gallie, 2000b). One of these mutants (vp9) is defective in ABA production, whereas the other
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(vp1) is defective in ABA perception. Thus, it would seem that in the wild‐ type ABA signaling is protecting the endosperm cells from PCD. Ethylene production increases concomitantly with premature PCD in these mutants, suggesting a link between the two plant‐growth regulators, although the exact nature of the association is as yet unclear. Two other types of signals have also been studied in relation to endosperm PCD. AOS may play a role, but again the evidence is indirect: production of an enzyme involved in AOS removal in barley, peroredoxin (Per1), is only expressed in the embryo and the aleurone, but not in the endosperm (Stacy et al., 1996). Also SOD production is reduced in the viviparous mutants previously described, suggesting that it may have a protective role against premature PCD (Guan and Scandalios, 1998). However, further work is required to obtain firmer evidence for a role of AOS in PCD of this tissue. The other possible trigger for PCD is endoreduplication. During endosperm development there is a wave of endoreduplication starting at the central region, and followed by the wave of PCD. In support of a role for endoreduplication in the regulation of PCD is the observation that in lily endosperm there is no endoreduplication and the endosperm cells remain alive at maturity (Young and Gallie, 2000a). However, again the evidence is indirect and would require further data to support a causative link. The mechanism of PCD in the starchy endosperm shares features with other types of PCD. DNA laddering is detected once the wave of cell death begins, and fragmentation of the DNA into larger fragments of 50–300 kb is detectable much earlier before cell death becomes evident (Young and Gallie, 2000a). There is also a clear link between ethylene and DNA laddering: exposure to ethylene accelerated DNA fragmentation in wild‐type maize. Furthermore, in maize starch‐deficient mutant shrunken2 kernels where PCD and DNA laddering progress more rapidly than wild‐type, treatment with AVG reduced the DNA breakdown. Nuclease activity is also detected in the endosperm at levels 10‐fold higher than in the embryo (Young et al., 1997). 3. The Endosperm of Ricinus communis Although the machinery associated with protein breakdown has not been a major focus in work on cereal endosperm PCD, some very interesting results have emerged from the study of endosperm PCD in the castor bean (Ricinus communis). In this species the endosperm persists into the mature seed and then dies following germination as the protein and lipid reserves it contains are depleted. The lipid reserves are mobilized via the formation of glyoxysomes, but additionally the endosperm cells also contain another organelle that develops from the ER and has been called the ricinosome (Schmid et al., 1999, 2001). Endosperm degeneration is accompanied by
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DNA fragmentation, and the number of ricinosomes dramatically increases as DNA fragmentation becomes evident (Schmid et al., 1999). A papain‐like endopeptidase (Cys‐EP), which shows homology to cysteine proteases and is associated to other plant PCD events, is also found in ricinosomes (Gietl et al., 1997; Than et al., 2004). The Cys‐EP is formed as a 45-kDa proenzyme, which is later cleaved to a 35 kDa mature form. Antibodies to Cys‐EP can be used to label the ricinosomes and follow the fate of the Cys‐EP as PCD progresses (Schmid et al., 1998). Labeling shows that the proenzyme accumulates in the intact ricinosomes in parallel with the increase in DNA fragmentation. However in the final stages of PCD, the ricinosomes rupture releasing the mature 35-kDal form. Thus, the ricinosomes seem to be playing a similar role to the large vacuole in the aleurone cell, releasing hydrolytic activity into the cytoplasm as PCD reaches its final stages. Again pH is postulated to play a part as acidification of isolated ricinosomes triggers the activation of the Cys‐EP protease (Schmid et al., 2001). In vivo this may occur as a result of cytoplasm acidification due to a change in tonoplast permeability late in PCD resulting in proton release from acidic vacuoles.
B. Cell Death in the Embryo and Suspensor Following endosperm degeneration, the next event in the life of the seed is the development of the embryo, and PCD plays a crucial role here too. Although shaping of the embryo also relies on PCD, this has been less well‐studied (Giuliani et al., 2002). Two major PCD events, however, have been studied in detail, the elimination of supernumerary embryos and the degeneration of the suspensor. In many plants, particularly gymnosperms, several embryos can develop from a single zygote, but usually only one develops to maturity (Bawa et al., 1989). The elimination of the excess embryos follows a precise pattern of cell elimination starting at the basal end of the embryo and moving toward the apex. Although little is known of the signals dictating which embryo survives and which die, more is known about the mechanisms of elimination. Thus, the PCD is accompanied by DNA fragmentation and increasing vacuolization (Filonova et al., 2002). The cytological features resemble those of autophagy in which the cytoplasm and organelles are engulfed and then lysed by the vacuoles. Suspensor degeneration is a more universal feature of plant embryogenesis. In the first zygotic division, the two daughter cells are diVerentiated into a densely cytoplasmic cell, which will divide to ultimately produce the embryo and a highly vacuolated cell, which undergoes fewer rounds of cell division to produce the suspensor. The suspensor acts as a conduit for nutrients to the developing embryo and degenerates once its function is
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complete. The signals limiting growth of the suspensor are thought to come from the embryo as mutants in which normal embryo development is impaired also have enlarged suspensor cells with features usually associated with embryonic cells (Schwartz et al., 1994). However, the nature of these signals remains unknown. PCD in the maize (Giuliani et al., 2002) and bean (Vicia faba; Wredle et al., 2001) suspensor is accompanied by DNA fragmentation and nuclear degeneration, which proceeds basipetally. In bean, evidence also exists of the formation of provacuoles and nuclear inclusions, which may indicate an autophagic type of PCD. However, the nuclear morphology of endosperm and suspensor cell PCD was compared in bean (Wredle et al., 2001) and diVerences were revealed in the arrangement of the chromatin, suggesting subtle diVerences in the PCD program in these two cell types. In plants it is also possible to stimulate embryo development in vitro by the appropriate use of plant‐growth regulators, and in these systems the embryonic and suspensor structures are clearly defined. This approach has been used for studying PCD in Norway spruce (Picea abies) embryos (Filonova et al., 2000; Smertenko et al., 2003). In this system as well, suspensor cell death was accompanied by DNA fragmentation initially into 50 kb fragments and later DNA laddering similar to events seen in animal apoptosis. Cytological studies showed the formation of large central vacuoles formed from smaller provacuoles (Filonova et al., 2000). The nucleus also fragmented, and finally the tonoplast ruptured and there was an almost complete lysis of cytoplasm, organelles, and nucleus. Thus, the PCD shows characteristics both of apoptosis and a form of autophagy. The cytoskeleton also rearranges during PCD in this system: microtubules are disrupted in the suspensor compared to the embryonic cells, and as PCD sets in, all microtubule fragments disappear completely (Smertenko et al., 2003). Conversely actin in the suspensor is organized into longitudinal and oblique cables. Disruption of these cables with actin‐depolymerizing drugs such as latrunculin results in aberrant suspensor development. The authors suggest that actin may be important in this system for autophagous PCD in these cells, as has been found in animal PCD (Aplin et al., 1992). The Norway spruce model was also used to study the role of caspase‐like enzymes in suspensor PCD (Bozhkov et al., 2004). Using synthetic peptide substrates and appropriate inhibitors, a caspase‐6‐like (VEIDase) activity was detected. The activity localized to suspensor and embryonic cells fated for PCD and inhibition of the VEIDase activity resulted in blocking suspensor PCD. This strongly suggests that this caspase‐like activity is required for PCD in these cells. However, how the caspase‐like activity functions here during PCD is unclear, although its early expression, coincident with microtubule disassembly and vacuole enlargement, might indicate a regulatory role. Cloning of the gene or genes responsible for the caspase‐like activity is clearly needed to help resolve its role.
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III. Leaf, Stem, and Root Development The major nonreproductive organs of the mature plant are the leaves, stem, and root system, and in all three of these PCD is important both in normal organ development and function and in shaping the organ in response to environmental cues. PCD is also the terminal event in the destruction of leaves in response to environmental and developmental cues known as senescence.
A. Leaf Sculpting One of the most dramatic examples of PCD is the remarkable sculpting of some leaves such as those of Monstera and the lace plant, Aponogeton madagascariensis (Fig. 1). Most complex leaf shapes are formed through an acceleration or repression of growth during morphogenesis (Dengler and Tsukaya, 2001), however, in Monstera and the lace plant, sculpting occurs through PCD. In Monstera tiny holes are formed early in development and increase in size with leaf expansion (Melville and Wrigley, 1969). These holes finally break through the leaf margin to form the characteristic lobed pattern. In the lace plant the program is diVerent. Here leaves form as solid shapes, and a subset of cells subsequently undergoes PCD to produce a pattern of holes (Gunawardena et al., 2004). The trigger for this PCD is unknown, but the mechanism seems to parallel other better‐studied systems with clear DNA degradation, chromatin condensation, and evidence of tonoplast rupture. However, DNA laddering is not detectable. Further work is clearly needed to understand how cells are targeted for PCD and how the tonoplast rupture is triggered.
Figure 1 Morphology of mature Lace plant leaf showing regular perforations arising from PCD remodeling of young expanding leaves. Bar ¼ 150 m. (Image kindly provided by Dr. Arunika Gunawardena.)
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B. Leaf Senescence Cell death has an additional role in plants compared to animals in that it is the terminal event of plant senescence. The process of senescence and cell death are clearly distinct at a physiological level because senescence, at least in leaves, can be a reversible process, whereas cell death is considered a terminal event (Thomas et al., 2003). Leaf senescence has been the focus of several reviews (Lim et al., 2003) and genomic‐wide approaches to identify regulatory networks (Buchanan‐Wollaston et al., 2003; Gepstein et al., 2003). This review will focus only on what is known about PCD at the cellular level, which is associated with leaf senescence. The signals initiating the overall process of senescence are common to other PCD events. Reduction of ethylene signaling (Grbic´ and Bleecker, 1995) and upregulation of cytokinin production (Gan and Amasino, 1995) delay senescence, indicating that the levels of these two PGRs are involved. Elevated cytoplasmic calcium is associated with leaf senescence in parsley (Huang et al., 1997), and calcium fluxes have also been implicated in two other leaf senescence systems (Chou and Kao, 1992; He and Jin, 1999). Thus, calcium signaling may play a role in leaf PCD, although further data are needed. Cytologically, PCD has been charted in rice leaves undergoing induced (by dark treatment) or natural senescence (Lee and Chen, 2002). In this system, features noted were cytoplasm depletion, organellar breakdown and expansion of the central vacuole, which at later stages contained inclusions possibly of chloroplast origin. Chromatin condensation and apoptotic bodies were not noted, and although cells became TUNEL‐positive and DNA became increasingly degraded, there was no evidence of DNA laddering. However, DNA laddering was detected in senescing leaves of other species such as wheat (Caccia et al., 2001), olive (Cao et al., 2003), and five other tree species (Yen and Yang, 1998). Chromatin condensation was also reported both in tobacco and in the monocot Ornithogalum virens (Simeonova et al., 2000). There was no clear reduction in mitochondrial membrane potentials in Pisum sativum mesophyll cells undergoing senescence (Simeonova et al., 2004), indicating that if mitochondria are associated with this PCD system, it is not in the same way as in animal apoptosis. Chloroplast disassembly seems to be an early sign of senescence, but whether this is part of the PCD mechanism remains uncertain (Thomas et al., 2003). C. Tracheary Element Differentiation A much better studied system is that of tracheary element diVerentiation. Tracheids are part of the xylem system, which in higher plants transports water and minerals from the roots to the rest of the plant. Tracheid cells, or
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tracheary elements (TEs), are dead at maturity, forming tapered hollow tubes with thickened cell walls. The formation of TEs requires targeted cell death of precursor cells derived from meristematic cells. An in vitro system has been developed in Zinnia to study this PCD in which mesophyll cells transdiVerentiate into TEs (Fukuda and Komamine, 1980). Evidence from the use of inhibitors implicates brassinosteroids, a type of steroid PGR, as a requirement for PCD progression (Yamamoto et al., 1997), and brassinosteroid biosynthesis genes increase in expression during the transition to PCD (Yamamoto et al., 2001) in this system. In Arabidopsis, mutants in brassinosteroid receptor genes also result in abnormal vascular diVerentiation (Cano‐Delgado et al., 2004), supporting the involvement of PGRs in this developmental process. AOS seems not to be involved in tracheary element PCD because inhibitors of the AOS generating enzyme NADPH oxidase did not inhibit TE PCD and levels of H2O2 were not elevated in cells following TE induction (Groover et al., 1997). In contrast, calcium/calmodulin signaling is involved. An increase in sequestered calcium occurs (Roberts and Haigler, 1989), and Ca2þ uptake is a requirement for PCD progression (Roberts and Haigler, 1990). Transiently elevated calmodulin levels are associated with PCD progression and calmodulin blockers also inhibit the process (Kobayashi and Fukuda, 1994). Calcium ionophores also induced DNA fragmentation (Groover and Jones, 1999). Furthermore, a link exists between calcium signaling and G proteins in this system. Thus, mastoparan, which is an activator of heterotrimeric G proteins, induced DNA degradation and this eVect was suppressed by inhibitors of Ca influx. Vacuolar collapse is the first visible sign of PCD in TE diVerentiation (Fukuda, 1996) followed by organelle degeneration. Organelle removal is a very rapid process, with nuclear degradation occurring within 10–20 min (Obara et al., 2001) and complete organellar destruction within 6 hr of vacuolar collapse (Groover et al., 1997). The vacuolar collapse may be triggered by changes in the organic ion permeability of the tonoplast and the collapse is preceded by loss of tonoplast integrity as the plasma membrane remains intact, but the tonoplast becomes unable to exclude fluorescein (Obara et al., 2001). Also treatment of diVerentiating TEs with an inhibitor of organic anion transport, probenecid, resulted in an acceleration of the vacuolar collapse (Kuriyama, 1999). In contrast to other systems, there is no evidence of nuclear condensation, fragmentation, or DNA laddering in TE PCD. DNA degradation was detectable using TUNEL staining, although this was a late event resulting from release of nucleases following vacuolar collapse (Mittler and Lam, 1995). In fact no DNA degradation is detected prior to vacuolar collapse (Obara et al., 2001). Various nuclease activities have been detected and genes encoding nucleases with very similar expression patterns are expressed
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during TE PCD, including an S1‐type nuclease (ZEN1), which is likely to be located in the vacuole (Sugiyama et al., 2000; Thelen and Northcote, 1989; Ye and Droste, 1996). ZEN1 may be of particular importance as it is specifically expressed in TEs, and both an anti‐ZEN1 antibody and antisense constructs of ZEN1 are able to block DNA degradation (Ito and Fukuda, 2002). Expression of these nuclease genes is just before the start of autolysis, indicating a role in DNA breakdown following vacuole rupture. Proteolysis appears to be required for TE PCD; in particular inhibition of both proteasome function and cysteine proteases delayed or inhibited the process (Fukuda, 2000). Two cysteine protease genes encoding papain‐like cysteine proteases are expressed as TEs move into PCD (Yamamoto et al., 1997; Ye and Varner, 1996) and cysteine protease activities of 20–30 kDa increase at a similar time (Beers and Freeman, 1997; Minami and Fukuda, 1995). These are probably located in the vacuole and may be involved in the autolysis of the cell following vacuolar rupture. A serine protease activity is also detected, but because it has a higher pH optimum, it is probably associated with the cytoplasm rather than the vacuole (Beers and Freeman, 1997). A protease activation cascade may also act as a possible trigger for vacuolar collapse, although the late timing of its activation makes this model less plausible (Fukuda, 2000; Groover and Jones, 1999). Clearly TE diVerentiation has proven an excellent system in which to study plant PCD, and some of the intercellular and intracellular regulatory pathways such as PGR and Ca2þ control have been elucidated. Transcriptomic approaches to identify genes expressed early after PCD induction in this system (Milioni et al., 2002) may also help to provide a more complete picture of how TE PCD is regulated, and whether such regulatory genes are shared with other PCD events during plant development.
D. PCD in Aerenchyma Formation Another plant tissue in which hollow tubes are formed via PCD is aerenchyma. This is a specialized root tissue common in wetland species, which can be induced in other plants including some major crops by water stress (Evans, 2004) (Fig. 2). It can be formed either by diVerential growth and cell separation or in the case of lysogenic aerenchyma by the selective death of cells in the mid cortex, spreading radially to form channels of air spaces, separated by living cells. The function of this architecture is to allow better gas exchange in waterlogged root systems (Armstrong, 1979). Induction of aerenchyma formation has been studied in more detail as the timing of events is easier to follow; however, the mechanisms are probably very similar in constitutive aerenchyma development. Low oxygen is the initial trigger for aerenchyma formation. This signal is transduced via ethylene: both
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Figure 2 Development of aerenchyma. Hematoxylin‐ and eosin‐stained maize root sections. (B) 2.5‐day‐old root treated with low oxygen levels (3%) to induce aerenchyma formation compared to (A) in 21% oxygen in which aerenchyma does not develop. Scale bar ¼ 100 m. Reproduced with permission from Gunawardena et al., 2001.
endogenously produced or exogenously elevated ethylene induces the formation of lysogenous aerenchyma (Drew et al., 1981). Presumably, the ethylene is released and then detected via receptors in the cells fated to die. A plausible system for distinguishing between cell fates would relate to the receptors as was seen for endosperm cells; however, this has not been shown directly for aerenchyma formation (Evans, 2004). Calcium signaling is again required: Ca2þ chelators and inhibitors of Ca2þ release from internal stores block aerenchyma formation (He et al., 1996). It has also been reported that cytosolic Ca2þ levels rise (Drew et al., 2000), and that this is linked to a signal transduction pathway involving inositol 1,4-5‐triphosphate, and protein phosphorylation, although details are as yet unpublished. It is also possible that AOS signaling may be active in aerenchyma PCD based on data showing an increase in AOS in cells beginning to degenerate (Bouranis et al., 2003). In induced aerenchyma formation in maize, the first cytological signs of cell death are plasma membrane (PM) invagination and the formation of small vesicles just beneath it (Gunawardena et al., 2001a). At a very similar time but probably afterwards, chromatin condensation is detected, followed by membrane‐enclosed organelles closely resembling animal apoptotic bodies (Gunawardena et al., 2001a). TUNEL staining is also detected in maize, although evidence for DNA laddering was less convincing (Gunawardena et al., 2001a). Changes in the cell wall are detected early in the PCD using specific antibodies, although they are not evident by
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electron microscopy until much later (Gunawardena, 2001b). In rice where aerenchyma is constitutive, a similar pattern is also observed with descriptions also of vacuolation (Webb and Jackson, 1986) and cytoplasm acidification probably resulting from loss of tonoplast integrity (Kawai et al., 1998). However in rice there was no TUNEL staining suggesting either a species‐specific diVerence in the PCD mechanism or perhaps reflecting a more rapid PCD progression in this system (Evans, 2004). What happens to the cell contents is an interesting question. Evidence from cryo‐SEM suggests that there may be rapid ionic reabsorption of cell contents by neighboring cells (van der Weele et al., 1996). Sadly, despite the wealth of knowledge about the cytological events in aerenchyma formation, and a relatively clear‐cut case for PGR triggering of PCD, there is very little biochemical data on aerenchyma PCD. Unlike the tracheary element work, no information is available to date on proteases, either as part of potential cascades or involved in the mopping up after vacuolar collapse, or on potential nuclease genes. This may be due to the inaccessibility of the aerenchyma tissue and the lack of a cell culture model (Evans, 2004).
E. Cell Death in the Root Cap A more exposed root tissue that undergoes PCD is the root cap. From early embryogenesis, root cap cells stain positively for TUNEL, indicating DNA degradation (Giuliani et al., 2002), whereas other areas of the root remain TUNEL‐negative. Root cap cells are derived continuously from the apical meristem and are sloughed oV at the surface of the root in a manner reminiscent of skin cells. Staining of mature onion roots reveals that only the outermost 2–3 cells of the root cap are TUNEL positive (Wang et al., 1996). In these cells nuclear fragmentation was also detected and the formation of apoptotic‐like bodies noted. Similar results were obtained with Arabidopsis roots (Møller and McPherson, 1998; Zhu and Rost, 2000). One study (Zhu and Rost, 2000) also showed chromatin condensation, vacuolation, and organelle destruction. Notably, the frequency of plasmodesmata fell in the outer root cap cells before DNA fragmentation was detected, suggesting an exclusion of the dying cells from the living root cap. Two Arabidopsis mutants give important insight into the determination of root cap cell fate. Mutants in TORNADO1 and TORNADO2 genes are defective in the early divisions of the epidermal/lateral root cap cells (Cnops et al., 2000). In these mutants, although root cap cells develop ectopically in the epidermal positions, they still die, suggesting that root cap cell death is a highly programmed event under genetic control. Signaling may involve AOS, as an Arabidopsis gene encoding an H2O2‐generating diamine oxidase
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(ATAO1) is expressed in lateral root cap cells (Møller and McPherson, 1998), which are undergoing vacuolation and are destined for PCD. Little is known about the machinery that executes PCD in these cells. However, two proteases from barley, aleurain and barley vacuolar aspartic proteinase (phytepsin), are expressed in root cap cells (Runeberg‐Roos and Saarma, 1998), suggesting a proteolytic mechanism, perhaps following the autophagic model seen in tracheary elements and aleurone cells.
IV. Flowering and Reproduction Plant reproduction requires the development of complex structures that interact with each other and that have an inherently limited life span. Thus, PCD is involved here both in shaping the sexual and nonsexual organs of the flower and in their removal once they are no longer needed. In addition, PCD also plays a crucial role in self‐incompatibility where genetically similar pollen is rejected by the female reproductive structures.
A. Sexual Organ Abortion While most flowering plants produce bisexual flowers containing both male and female organs, approximately 10% produce unisexual flowers either on the same or on diVerent individuals (Irish and Nelson, 1989). In these flowers sex determination is achieved by arrest or abolition of the sexual organ primordia and the role of PCD in this process has been reviewed (Wu and Cheung, 2000). Two types of plant growth regulator have been implicated in regulation of sexual organ abortion in maize. GA treatment partially abolished the phenotype of anther ear 1 and dwarf mutants, which produce male sexual organs (stamens) on the female flowers (Irish and Nelson, 1989), and which encode biosynthetic GA genes (Bensen et al., 1995; Winkler and Helentjaris, 1995). In another mutant ts2, the male tassels show varying degrees of feminization and the TS2 gene encodes a protein with homology to steroid dehydrogenases (DeLong et al., 1993). This enzyme may act on brassinosteroids, another class of PGRs mentioned previously (Wu and Cheung, 2000).
B. PCD in the Male Sexual Organs Pollen develops within locules of the anther, which are lined with a tissue known as the tapetum. This plays a nutritive role during pollen development, coating the pollen grains with proteins and lipids and forming the
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intricately sculpted outer layer of the pollen wall known as the exine. However, the tapetum degenerates once its function is completed. This degeneration is accompanied by cell shrinkage, vacuolation, and thinning of the cell walls (Bedinger, 1992). Chromatin condensation and ER inclusions have been reported in Lobivia rauschii and Tillandsia albida (Papini et al., 1999), as well as DNA fragmentation in barley anthers (Wang et al., 1999). In Brassica oleracea, B. napus, Digitalis purpurea and a cultivated form of Fuchsia, nuclear blebbing has also been reported in some cells (A. Stead, unpublished data). However, little seems to be known about the signaling or mechanism of tapetal PCD despite its importance in plant fertility. One study (Lesniewska et al., 2004) reported a depolarization of mitochondrial membranes in tapetal cells in advanced PCD which may indicate an involvement of the mitochondria in this process. A rice cysteine protease gene has been isolated, which is expressed in the tapetum and developing pollen grains, late during pollen development (Lee et al., 2004). Mutants in this gene are impaired in pollen development. It will be interesting to understand the function of this gene more fully; perhaps it may have relevance to the PCD mechanism operating in tapetal cells. Two distinct types of mutation, nuclear or cytoplasmic, result in an early onset of tapetal PCD. The best known of these is cytoplasmic male sterility, which has been described in over 150 species (Schnable and Wise, 1998). In these plants, mutations in the mitochondrial genome result in male sterility through aberrations in anther development, or more frequently, tapetal degeneration and pollen abortion soon after meiosis. Note that although the mitochondrial defect is present in all the cells of the plant, the major defect is in anther function. Because the mutation is known to be in the mitochondrial genome, it has long been assumed that the death of the anther cells is due to a failure of the mitochondria to sustain the large energy needs of these tissues (Levings, 1993). Thus, PCD in these cells may well be triggered by mitochondrial dysfunction. Evidence for the mechanism at play has come from studies of the sunflower CMS system, PET1‐CMS (Balk and Leaver, 2001). Here cell condensation, DNA laddering, and chromatin condensation are associated with the early death of the tapetum (Balk and Leaver, 2001). Mitochondria persist in the early PCD stages; however, release of cytochrome c into the cytosol is detected before the changes in cell morphology. This is followed by a loss of outer mitochondrial membrane integrity and a fall in the respiratory control ratio. Thus, in this system there appears to be a central role for the mitochondrion in PCD reminiscent of its role in mammalian PCD (Krishnamurthy et al., 2000). Numerous nuclear mutants exhibiting male sterility have also been described. In some cases these too owe their male sterility to a premature degeneration of the tapetum. Tapetal PCD was studied in one male sterile rice mutant (Ku et al., 2003) and was associated with cytoplasmic shrinkage,
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membrane blebbing, vacuolation, changes in mitochondrial morphology, and DNA fragmentation. The DNA fragmentation detected by TUNEL was seen premeiotically, which is earlier than in CMS. However, there is as yet no precise data on the mechanism of PCD in this system. Other anther tissues also undergo PCD: pollen is released from the anther locule by death of the stomium cells, and death of cells within the endothecium provides a mechanism for the ejection of the pollen at dehiscence. In the Arabidopsis mutant delayed dehiscence1, there is a delay in stomium degeneration resulting in male sterility (Sanders et al., 2000). The gene encodes an enzyme in the jasmonic acid biosynthetic pathway, and application of jasmonic acid restored wild‐type dehiscence. Thus, this plant growth regulator may be important directly or indirectly in the correct timing of death in these cells. In tobacco, ethylene may also be involved, as blocking ethylene perception delayed stomium degeneration (Rieu et al., 2003). Although the biochemical mechanisms of PCD in these cells are relatively under studied, a report of a cysteine protease expressed in the endothecium of brinjal (Xu and Chye, 1999) and a thiolendopeptidase expressed in tobacco stomium (Koltunow et al., 1990) may lead to further characterization of the processes involved.
C. PCD in the Female Gametophyte Numerous cells and tissues undergo PCD throughout female gametophyte development and function. The first step in the production of the female gametophyte is meiosis of the megaspore mother cell to yield four meiotic products. However, in most plant species, three of these degenerate, and the eight cells of the mature female gametophyte contained within the embryo sac derive from two rounds of mitosis of just one meiotic product. The three degenerating cells show some classical signs of PCD including cell shrinkage, cytoplasmic disorganization, and chromatin condensation (Bell, 1996). The next PCD event concerns four of the eight embryo sac cells. These are known as the synergids and the antipodal cells. The synergids are positioned close to the micropyle, the pore through which the pollen tube enters the ovule to eVect fertilization, whereas the antipodal cells are at the opposite end of the embryo sac. During or shortly before fertilization, one of the two synergids degenerates, and in some species the antipodal cells also degenerate after fertilization. Because the synergids in some species degenerate independently from pollination, it seems likely that the signals are intrinsic to their development (Drews and Yadegari, 2002). In Nicotiana, the degeneration of the synergids is characterized by a change in nuclear morphology, collapse of the vacuole, and disappearance of organelles (Huang and Russell, 1994). There is also a loss of membrane integrity (Huang and Russell, 1992) and
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the generation of ‘‘pinched‐oV ’’ cytoplasmic bodies (Huang et al., 1993). Likewise in wheat (Triticum aestivum) synergid PCD is accompanied by nuclear shrinkage and deformation, and chromatin condensation (An and You, 2004). Thus, the symptoms of PCD evident in synergid degeneration may fit with the autophagous model of PCD. However, an Arabidopsis female gametophyte mutant (gfa2) has been described in which one of the two synergids that normally degenerates following pollination, fails to die (Christensen et al., 2002). GFA2 encodes a member of the DnaJ protein family, which acts as a chaperone in the mitochondrial matrix. The authors suggest that GFA2 may be required for correct folding of mitochondrial proteins, which in turn are required for PCD in the synergid. Thus an involvement of mitochondrial function in synergid PCD at least in Arabidopsis seems to have been established. A comparison of antipodal and synergid PCD in wheat (An and You, 2004) indicates that the PCD mechanisms may diVer in these closely positioned cells. In particular, there are clear diVerences in chromatin and nuclear morphology, with only slight changes in nuclear volume in antipodal cells and only partial chromatin condensation. Also, in the gfa2 mutant previously described, antipodal PCD is not aVected. This may reflect the diVerent function of the cells and of course may vary in diVerent species, but would be an interesting area for further research.
D. PCD During Self‐Incompatibility Interactions One reproductive tissue that has received a lot of attention is the pollen tube and its interactions during pollination and fertilization. Over 50% of higher plants protect against self‐fertilization by self‐incompatibility (SI) mechanisms that are thought to have evolved independently in diVerent plant families several times. SI involves highly specific interactions between the pollen and the pistil. In incompatible reactions fertilization is prevented and in some systems is associated with PCD of the pollen tube. Research (Thomas and Frankin‐Tong, 2004) has shown that SI in Papaver is associated with DNA fragmentation and caspase‐3‐like activity. An inhibitor of caspase‐3, the peptide Ac‐DEVD‐CHO, inhibited DNA fragmentation and pollen tube growth. Poly(ADP) ribose polymerase (PARP) is involved in nuclear DNA repair (Smith, 2001) and is a classic substrate for caspase‐3 enzymes. This study (Thomas and Franklin‐Tong, 2004) showed that the caspase‐3‐like activity elicited by SI in the Papaver pollen tubes was able to cleave PARP, and that detection of this activity preceded DNA fragmentation. SI also stimulated the release of cytochrome c into the cytosol. Papaver SI results in increases in cytosolic [Ca2þ] (Franklin‐Tong et al., 1993), and elevating cytosolic [Ca2þ] in the pollen tubes using Mastoparan resulted in the release
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of cytochrome c, providing a link between SI, Ca signaling, and cytochrome c release. Mastoparan treatment also stimulated caspase‐3‐like activity linking the DNA degradation and cytochrome c release to increases in cytosolic [Ca2þ]. Thus, at least in this SI system there is compelling evidence for a PCD mechanism with several of the hallmarks of animal PCD.
E. Petal Senescence and Cell Death Petal senescence, unlike leaf senescence, inevitably ends with PCD; however again, whether the two processes should be treated as one remains hotly debated (Thomas et al., 2003; van Doorn and Woltering, 2004). Global genomic approaches are being used in addition to studies of individual processes to identify genes involved in petal senescence (e.g., Breeze et al., 2004), and the expression of several of the known homologues to animal PCD‐related genes— namely Beclin, Bax inhibitor I, and dad‐1, (Hu¨ckelhoven, 2004; Orza`ez and Granell, 1997; WagstaV et al., 2003; C. WagstaV, unpublished results)—has been revealed, although the role of these genes in regulating petal PCD remain unresolved. Evidence from electron microscopy across Alstroemeria senescent petals (WagstaV et al., 2003) clearly shows that PCD is already occurring at relatively early stages of petal senescence. In many flowers including Arabidopsis, tobacco, and many others, senescence and thus PCD is triggered by ethylene (Stead and van Doorn, 1994) sometimes associated with pollination (O’Neill, 1997). However, in another group, including lilies such as Alstroemeria, the role of ethylene is less clear‐cut. Given the very tightly regulated life span of many flowers (van Doorn and Stead, 1997), endogenous cellular factors may determine onset of PCD, but this has not been clearly demonstrated. Once the trigger for PCD has been perceived by the cell, the intracellular mechanisms operating in petal PCD closely resemble those of the other systems described. Calcium signaling and GTP‐binding proteins have been implicated in some species (Porat et al., 1994). ROS accumulation has also been reported both in ethylene‐induced (Bartoli et al., 1996) and in ethylene‐independent (Panavas and Rubinstein, 1998) petal senescence, together with a drop in antioxidants (Bartoli et al., 1997). There is often evidence of tonoplast invagination (Matile and Winkenbach, 1971; Phillips and Kende, 1980) or the formation of vesicles (Smith et al., 1992), and in the final stages only a thin layer of cytoplasm remains (Stead and van Doorn, 1994; WagstaV et al., 2003) and organelles disappear (Stead and van Doorn, 1994). Changes in membrane composition, fluidity, and peroxidation occur in several species (Rubinstein, 2000). In some cases, such as carnation (Dianthus caryophyllus; Fobel et al., 1987), daylily (Hemerocallis hybrid; Panavas and Rubinstein, 1998), and rose (Rosa hybrid; Fukuchi‐Mizutani et al., 2000),
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lipid peroxidation (e.g., through the action of lipoxygenases [LOX]) may be one of the key factors eVecting loss of membrane integrity. However, in other species like Alstroemeria (Leverentz et al., 2002), loss of membrane semipermeability was chronologically separated from LOX activity that had declined by over 80% by the onset of electrolyte leakage. Thus, in this species loss of membrane function was not related to LOX activity or accumulation of lipid hydroperoxides per se representing a novel pattern of flower senescence. DNA laddering has been found in some petals such as Alstroemeria (WagstaV et al., 2003) and pea (Orza`ez and Granell, 1997), and both nucleases (Breeze et al., 2004; Langston et al., 2005; Panavas et al., 1999) and proteases (Stephenson and Rubinstein, 1998; WagstaV et al., 2002) are upregulated during petal senescence in several species. Significantly in Sandersonia aurantiaca, a member of a class of cysteine proteases carrying the KDEL C‐terminal motif is expressed during senescence (Eason et al., 2002), which shows homology to a protease from Ricinus communis implicated in ricinosome–mediated endosperm PCD (Gietl et al., 1997) (see earlier discussion). This suggests a mechanism for petal cell PCD, at least in this species, which may parallel the vacuole‐driven autophagous model previously described in the Ricinus endosperm.
V. Conclusions and Future Directions The examples of developmental PCD in the plants reviewed here illustrate a diversity of data sets but also some fundamental diVerences between the diVerent cell fates and PCD mechanisms. It is perhaps useful to compare the data available for each of the systems reviewed (Table I) to look for commonalities. It can be asked how useful is the animal model of apoptosis or autophagy in explaining plant PCD mechanisms? In common with most animal PCD systems, a signal extrinsic to the cell, often hormonal, is usually involved. However there is no one PGR that induces PCD, although ethylene, GA, and brassinosteroids appear in more than one system. There are few clear‐cut examples of completely autonomous PCD, although perhaps the root cap comes close. In this case the evidence from mutant studies (Cnops et al., 2000) supports a direct genetic control of PCD for these cells, and more data from mutants in other systems where extrinsic signals have not been clearly defined, such as PCD during ethylene‐independent petal senescence, may reveal the degree of PCD autonomy. Another interesting issue is how cells adjacent to cells undergoing PCD protect themselves from the ‘‘death signals.’’ Data from ethylene signaling in the endosperm (Gallie and Young, 2004) indicate that at least in some systems this may be achieved by a diVerential in sensitivity. However, in the TE system another mechanism seems to be operating. Here there is evidence for an inhibitor of
Table I
Comparison of PCD Characteristics from DiVerent Plant Tissues Possible Signaling
Plant Tissue
PGR Control
Aleurone cells
GA (þ); ABA ( )
Starchy endosperm
Ethylene (þ); ABA ( )
Ricinus endosperm
Germination
Suspensor
Signals from embryo
Supernumary embryos
Signals from Female gametophyte nutrient depletion Unknown
Leaf sculpting
Ca2þ/Phosphatase Signaling Ca2þ increases inhibited by protein phosphatase and protein kinase inhibitors NO( )? AOS (þ) AOS(þ)?
Possible Mechanism
Apoptosis? No DNA laddering, or PM blebbing
Autophagy? Possible proteolytic cascade, lytic vacuoles
DNA laddering nucleases
DNA laddering Caspase‐6 activity
DNA fragmentation but no laddering
Fate of Corpse
Some Key References
Disappears
Fath et al., 1999; Kuo et al., 1996; Lam et al., 2000
Remains
Stacy et al., 1996; Young and Gallie, 2000a; Young et al., 1997 Schmid et al., 1999, 2001
Lytic ricinosomes, from ER, proteolytic cascade Vacuolarization, autophagic vacuole
Disappears
Vacuolarisation, vacuole rupture TUNEL positive
Disappears
Vacuole rupture,
Disappears
Disappears
Bozhkov et al., 2004; Giuliani et al., 2002; Wredle et al., 2001 Filonova et al., 2002
Gunawardena et al., 2004
Leaf senescence
Ethylene (þ) and cytokinin ( )
Ca2þ increases
Tracheary elements
Brassinosteroids
Ca2þ increases, calmodulin, G proteins
Aerenchyma
Ethylene
Ca2þ increases, protein phosphorylation
Apoptotic bodies, DNA laddering (?)
Root cap
Genetic control
Possible AOS
Apoptotic bodies, chromatin condensation
Sex organ abortion Tapetum
GA and brassinosteroids
CMS
Mitochondrial disfunction
Cytochrome c release
DNA laddering in some species
Chromatin condensation DNA laddering, nuclear blebbing; mitochondrial membrane depolarization DNA laddering, chromatin condensation
Vacuolation
Disappear
Vacuolar collapse, rapid organelle degradation, nucleases No DNA laddering Vacuolation, loss of tonoplast integrity
Cytoplasm disappears cell wall structural
Vacuolation, organelle destruction
Vacuolation
Cytoplasm disappears cell wall structural Sloughed oV
Disappear, cell contents taken up by microspores Disappear
Caccia et al., 2001; Gan and Amasino, 1995; Grbic´ and Bleecker, 1995; Huang et al., 1997; Lee and Chen, 2002 Roberts and Haigler, 1989, 1990
Drew et al., 2000; Gunawardena et al., 2001a,b Møller and McPherson, 1998; Wang et al., 1996; Zhu and Rost, 2000 Wu and Cheung, 2000 Lesniewska et al., 2004; Papini et al., 1999; Wang et al., 1999 Balk and Leaver, 2001 (Continued )
Table I Continued Possible Signaling
Plant Tissue
PGR Control
Ca2þ/Phosphatase Signaling
Possible Mechanism
Apoptosis?
Endothecium
Jasmonic acid, ethylene
Megaspores
Embryo sac expansion
Synergids
Pollination in some species
Petal senescence
Ethylene in some species
Ca2þ/phosphatase signalling, ROS increases
DNA laddering
Pollen tube
SI interactions
Cytochrome c release, increased Ca2þ
DNA fragmentation Caspase‐3 activity
Autophagy?
Fate of Corpse Disappear
Chromatin condensation; requirement for mitochondrial chaperone
Cell shrinkage, chromatin condensation Vacuolar collapse
Disappear
Tonoplast invagination formation of vesicles
Mesophyll cells disappear
Burst releasing contents
Burst
Some Key References Rieu et al., 2003; Sanders et al., 2000 Bell, 1996
An and You, 2004; Christensen et al., 2002; Huang and Russell, 1992, 1994 Bartoli et al., 1996; Orza`ez and Granell, 1997; Panavas and Rubinstein, 1998; Phillips and Kende, 1980; Porat et al., 1994; WagstaV et al., 2003 Thomas and Franklin‐Tong, 2004
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proteasome‐mediated protein degradation, which is released into the apoplastic space (Endo et al., 2001) and which protects living cells from the hydrolases released during TE PCD. A more thorough comparison of gene expression and biochemistry between cells undergoing PCD and their living neighbors may be helpful to understand more fully the complexity of extrinsic cellular signaling leading to PCD. Intracellular calcium signaling seems almost universal where it has been assayed and may well be a common feature in all plant PCD. Involvement of protein kinase and phosphatase control also seems an area worthy of further study. Release of cytochrome c has only been reported in two PCD systems, CMS and SI, both of which might be considered special cases of PCD. Whether cytochrome c release is more widespread is not clear at present, but it would be very useful to resolve the extent to which it occurs in any of the other systems, as this may shed light on the involvement of the mitochondria in the plant PCD systems. DNA laddering appears in many of the PCD systems but is not universal, and may not ultimately be very useful in understanding the PCD mechanism. Laddering clearly suggests activation of nucleases, which cut the DNA in a specific manner but does not explain much about how the activation occurs. The cytological evidence is perhaps more useful. Vacuolation accompanies PCD in more than half of the examples reviewed, and in general rupture of the vacuole coincides with release of hydrolases into the cytoplasm. This suggests a possible model that would account for at least some of the systems reviewed (Fig. 3). In this model an external signal activates increases in cytoplasmic calcium, which in turn stimulates the coalescence of small vacuoles derived from either the ER or the Golgi to form a large vacuole. This vacuole accumulates hydrolytic enzymes. Its collapse, resulting either from ROS accumulation or activation of a proteolytic cascade, releases hydrolases into the cytoplasm. The hydrolase release results in organellar breakdown and macromolecule degradation ending in cell death. This model has clear parallels to animal autophagy, although the vacuole rather than the lysosome is the primary organelle involved. However, this model is clearly not universal, and some PCD systems such as CMS, SI, and PCD during leaf senescence may follow a very diVerent pattern, perhaps more similar to animal apoptosis. An important distinction in the role of cell death in organ development is also the fate of the dead cell. Thus, in some cases such as aerenchyma formation, the corpse of the cell is removed leaving a space; however, in other cases the dead cell has a structural (e.g., TE formation) or nutritive (e.g., the starchy endosperm) role. However, from the evidence reviewed here, the fate of the cell and the mechanism of its destruction do not seem to be closely linked; thus, both TE formation and endosperm destruction in Ricinus follow quite closely the model outlined, although the fate of the cell
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Figure 3 Model showing major signals (black arrows) and cytological/biochemical events (open arrows) during autophagic‐type PCD in plants. Cell types/tissue in which these features have been reported are indicated as: A, aleurone cells; AE, aerenchyma; L, leaf sculpting; LS, leaf senescence; P, petal senescence; R, Ricinus endosperm; RC, root cap; S, starchy endoasperm; SE, supernumerary embryos; SI, pollen tube during SI interaction; SU, suspensor; SY, synergids; T, tapetum; TE, tracheary elements.
is very diVerent. There may thus be parallel pathways that diverge at the point of activation of cell wall‐degrading enzymes. This review has focused largely on the mechanisms of PCD seen in diVerent stages of plant development. This mechanistic approach seemed important in an attempt to understand the diversity of developmental PCD
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in plants. The future challenge lies in filling in biochemical and molecular gaps, which will explain not only how plant cells die during development but also what is regulating their demise.
Acknowledgments I would like to thank Dr. Tony Stead and Dr. Carol WagstaV for helpful comments on the manuscript and Dr. David Evans and Dr. Arunika Gunawardena for supplying the images in this review.
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The Blood‐Testis Barrier: Its Biology, Regulation, and Physiological Role in Spermatogenesis Ching‐Hang Wong and C. Yan Cheng Population Council, New York, New York 10021
I. Introduction II. The Molecular Architecture of the BTB III. Models to Study BTB Dynamics A. The Cadmium Model B. Other Potential Models IV. Regulation of BTB Dynamics A. Does the BTB Restructuring Share Similar Features Pertinent to Cellular Movement with Cell‐Matrix Junction Dynamics? B. Roles of Cytokines and their Downstream Signaling Pathways in Regulating BTB Dynamics C. Interplay of Proteases and Protease Inhibitors D. Unique Physiological Features of the BTB Pertinent to its Regulation During Spermatogenesis E. An Integrated Model of BTB Regulation During Spermatogenesis V. Concluding Remarks Acknowledgments References
The blood‐testis barrier (BTB) in mammals, such as rats, is composed of the tight junction (TJ), the basal ectoplasmic specialization (basal ES), the basal tubulobulbar complex (basal TBC) (both are testis‐specific actin‐based adherens junction [AJ] types), and the desmosome‐like junction that are present side‐by‐side in the seminiferous epithelium. The BTB physically divides the seminiferous epithelium into basal and apical (or adluminal) compartments, and is pivotal to spermatogenesis. Besides its function as an immunological barrier to segregate the postmeiotic germ‐cell antigens from the systemic circulation, it creates a unique microenvironment for germ‐cell development and confers cell polarity. During spermatogenesis, the BTB in rodents must physically disassemble to permit the passage of preleptotene and leptotene spermatocytes. This occurs at late stage VII through early stage VIII of the epithelial cycle. Studies have shown that this dynamic BTB restructuring to facilitate germ‐cell migration is regulated by two cytokines, namely transforming growth factor‐ 3 (TGF‐ 3) and tumor necrosis Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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factor‐ (TNF ), via downstream mitogen‐activated protein kinases. These cytokines determine the homeostasis of TJ‐ and basal ES‐structural proteins, proteases, protease inhibitors, and other extracellular matrix (ECM) proteins (e.g., collagen) in the seminiferous epithelium. Some of these molecules are known regulators of focal contacts between the ECM and other actively migrating cells, such as macrophages, fibroblasts, or malignant cells. These findings also illustrate that cell–cell junction restructuring at the BTB is regulated by mechanisms involved in the junction turnover at the cell‐matrix interface. This review critically discusses these latest findings in the field in light of their significance in the biology and regulation of the BTB pertinent to spermatogenesis. ß 2005, Elsevier Inc.
I. Introduction In the seminiferous tubule of the mammalian (such as rats and mice) testis, somatic Sertoli cells are crucial to postmeiotic germ‐cell development during spermatogenesis by providing the nourishment and mechanical support to developing germ cells until they become mature spermatids (i.e., spermatozoa). Sertoli cells also create the BTB in the seminiferous epithelium that physically divides the seminiferous epithelium into the basal (outside the barrier) and the adluminal (behind the barrier) compartments (for reviews, see Dym and Fawcett, 1970; Setchell, 1980). This almost impermeable barrier is of great physiological importance, and it is critically selective to molecules that can enter the adluminal compartment (for reviews, see Bart et al., 2002; Cheng and Mruk, 2002; Griswold, 1998; Mruk and Cheng, 2004b). Thus, when the BTB is dysfunctional, germ‐cell diVerentiation and development are arrested (for a review, see Toyama et al., 2003). Under the electron microscope, the BTB appears to be a continuous strand of electron‐dense material sandwiched between two apposing Sertoli cells, where tight junctions (TJs) are found (Fig. 1). Adjacent to the TJ strands, some actin filament bundles can be identified between the Sertoli cell plasma membrane and the subsurface cistern of the endoplasmic reticulum. This is the typical structural feature of the basal ectoplasmic specialization (ES), a type of actin‐based adherens junction (AJ) specifically found in the testis (for reviews, see Mruk and Cheng, 2004a; Russell, 1977b; Toyama et al., 2003; Vogl et al., 2000) (Fig. 1). ES is not restricted to the BTB but is also found at the site where developing (step 8 and beyond in rats) spermatids attach to the Sertoli cell in the adluminal compartment, although the structure is only restricted to the Sertoli cell side and is known as apical ES (for reviews, see Mruk and Cheng, 2004a; Toyama et al., 2003; Vogl et al., 2000). The coexistence of TJ and ES structures, together with the desmosome‐like junction and the tubulobulbar complex (TBC) at the BTB,
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Figure 1 Schematic drawing illustrating the relative locations of diVerent junction types between Sertoli cells, as well as between Sertoli and germ cells at diVerent stages of germ‐cell development in the seminiferous epithelium of the rat testis. At present, it is not known whether focal adhesion complex, a cell‐matrix actin‐based anchoring junction type, is present in the rodent testis. Other anchoring junction types present in the seminiferous epithelium are: (1) cell–cell actin‐based AJ (e.g., basal and apical ES, basal and apical TBC); (2) cell–cell intermediate filament‐based desmosome‐like junctions (having the properties of both desmosomes and gap junctions uniquely found in the testis); and (3) cell‐matrix intermediate filament‐based hemidesmosomes. The electron micrographs are cross‐sections of the seminiferous epithelium from an adult rat testis. The micrograph in the left panel shows the ultrastructure of the blood‐testis barrier (BTB) between two apposing Sertoli cells, which is composed of both tight junctions (TJs) and basal ectoplasmic specialization (basal ES) that are present side‐by‐side. The BTB morphologically divides the epithelium into the basal and apical (adluminal) compartments. The basal ES is typified by the presence of actin filament bundles sandwiched between the Sertoli cell membrane and the cisternae of the endoplasmic reticulum (ER), which is found on both sides of the Sertoli cells, whereas TJs are found between the basal ES of the two adjacent Sertoli cells. These junctions in turn constitute the functional BTB. The micrograph in the right panel shows the apical ES between a Sertoli cell and an elongate spermatid. The apical ES, similar to basal ES, is typified by the presence of actin filament bundles sandwiched between the Sertoli cell membrane and the ER. However, this ultrastructural feature is restricted only to the Sertoli cell side and is absent in the spermatid. Also, TJ is not present in apical ES. Ac, Acrosome. Bar ¼ 0.4 m on the left panel and 0.2 m on the right panel.
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Figure 2 A simplified schematic diagram illustrating the morphological diVerences between the blood‐testis barrier (BTB) and other barriers in the mammalian body (e.g., the blood‐brain [BBB] and the blood‐retina barrier [BRB]). (A) Tight junction (TJ) coexists with basal ES, basal TBC, and desmosome‐like junction that constitute the functional BTB. (B) In other barriers (e.g., BBB), TJ is restricted to the apical surface of the epithelium, sealing the intercellular space, whereas AJ is located right behind the TJ fibrils.
is not found in other barriers such as the blood‐brain barrier (BBB) and the blood‐retinal barrier (BRB), where AJs and desmosomes are distinctly separated from TJs, lying behind the TJ fibrils as individual entities (Fig. 2) (for reviews, see Denker and Nigam, 1998; Petty and Lo, 2002; Rubin and Staddon, 1999). Although an intact BTB is essential for spermatogenesis, spermatogonia and preleptotene/leptotene spermatocytes reside outside the BTB in the basal compartment of the seminiferous epithelium. During spermatogenesis, preleptotene/leptotene spermatocytes must pass through this barrier to gain entry to the adluminal compartment where meiosis can be completed, which occurred at late stage VII and early stage VIII of the epithelial cycle (Russell, 1977a) (for reviews, see Cheng and Mruk, 2002; Dym and Fawcett, 1970; Mruk and Cheng, 2004b). Therefore, the BTB is a dynamic structure that undergoes cycles of ‘‘opening’’ and ‘‘closing’’ during the epithelial cycle to facilitate germ‐cell migration, yet its integrity must not be compromised so that the microenvironment behind it can be maintained. These events obviously have to be intricately regulated and is likely involving a complicated network of signaling cascades and rapid turnover of junction‐associated molecules. In this review, some findings based on studies using diVerent models that mimic the disassembly (opening?) and reassembly (closing?) of the BTB are discussed. It is increasingly clear that the mechanism(s) regulating BTB dynamics is the reminiscence of that being utilized to regulate junction turnovers at the cell–extracellular matrix (ECM) interface. The discussion here is focused primarily on newer findings, and how they shed light on the role of the BTB in spermatogenesis. Earlier studies have been discussed in several excellent reviews (see Byers et al., 1993; Dym and Cavicchia, 1997; Pelletier and Byers, 1992).
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II. The Molecular Architecture of the BTB At the molecular level, the BTB is currently known to be composed of three major classes of proteins: integral membrane proteins, peripheral adaptors and their associated signaling molecules, and cytoskeletal proteins. The extracellular domains of integral membrane proteins, which are present in adjacent Sertoli cells near the basolateral region of the epithelium, seal up the intercellular space, forming an interlocking structure usually via homophilic protein–protein interactions (e.g., occludin–occludin). The cytoplasmic domains of the transmembrane proteins are connected to the underlying cytoskeletal network via adaptors, whereas their interactions are regulated via signaling molecules, such as protein and lipid kinases and phosphatases. To date, three diVerent types of TJ‐associated transmembrane proteins: occludins (Furuse et al., 1993; Moroi et al., 1998), claudins (Gow et al., 1999) and junctional adhesion molecules (JAMs) (Gilki et al., 2004), and two types of AJ‐transmembrane proteins: classic cadherins (Lee et al., 2003; Wine and Chapin, 1999) and nectin‐2 (Mueller et al., 2003; Ozaki‐Kuroda et al., 2002), are known to constitute the BTB in rat or mouse testes. These proteins are linked to actin microfilaments via adaptors. For example, zonula occludens (ZO)‐1/‐2/‐3 are the adaptors to which occludin (Furuse et al., 1994; Itoh et al., 1999b), claudins (Itoh et al., 1999a), and JAMs (Bazzoni et al., 2000; Ebnet et al., 2000) attach; ‐ and ‐catenins are the binding partners of cadherins (Hulsken et al., 1994; Sacco et al., 1995); whereas afadin is the adaptor of nectin‐2 (Ozaki‐Kuroda et al., 2002; Takahashi et al., 1999). Because these proteins have been extensively reviewed elsewhere (Balda and Matter, 2000; Bazzoni and Dejana, 2004; Cheng and Mruk, 2002; Irie et al., 2004; Lapierre, 2000; Lee and Cheng, 2004; Mitic and Anderson, 1998; Mruk and Cheng, 2004b; Takai and Nakanishi, 2003), they are not being discussed in detail herein. However, it is of interest to note that these structural components are important markers for studying BTB dynamics. The modulation of their production, degradation, internalization, recycling, and most importantly the association and dissociation between them are keys to the BTB restructuring. Furthermore, the regulatory signaling pathways that are to be discussed herein can lead to changes in one or more of these parameters, resulting in the delicate restructuring of the BTB.
III. Models to Study BTB Dynamics The regulation of BTB dynamics cannot be fully understood without the use of suitable animal models. Although the in vitro model using primary Sertoli cell cultures has long been applied to study inter‐Sertoli cell TJ‐dynamics
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(Byers et al., 1986; Janecki and Steinberger, 1986), it can never fully resemble the complexity of restructuring events in vivo. Therefore, the establishment of in vivo models for studying BTB dynamics has become increasingly important in the field. The models that have been used thus far involve the application of diVerent chemicals to host animals via gavage, intraperitoneal injections, or intratesticular injections, which subsequently target the BTB and perturb its functionality. These models thus allow investigators to dissect changes in the seminiferous epithelium during BTB restructuring. Unfortunately, most of the BTB disruptions caused by these substances are irreversible, making it diYcult to study the event of BTB reassembly. Nonetheless, these studies are still very important for investigating the biology of BTB restructuring, and are also therapeutically important. For example, if the permeability of the BTB can be precisely manipulated, therapeutic drugs can be delivered (e.g., for chemotherapy and inflammatory disorders) to the microenvironment behind the barrier. It can also be applied to the development of male contraceptives, because BTB disassembly is usually associated with germ‐cell depletion from the epithelium.
A. The Cadmium Model Cadmium is a major industrial pollutant, an environmental toxicant, and an endocrine disruptor that can cause significant damage to many organs, including the lung, liver, kidney, and testis (for reviews, see Lafuente et al., 2001; Prozialeck, 2000). Its presence in daily‐consumed items is widespread, including water, food, beverages, and cigarettes, and it has a long half‐life of 15 years in humans, making its toxicity accumulative (Kjellstrom and Nordberg, 1978). Its damaging eVects to the testis have been known for decades, including disruption of the Sertoli cell TJ‐barrier function and the vascular system, failure of spermiation, germ‐cell loss from the seminiferous epithelium, tissue necrosis, and apoptosis (Aoki and HoVer, 1978; Gunn and Gould, 1970; Hew et al., 1993a; Mason et al., 1964; Xu et al., 1996), some of which can be prevented by zinc (Parizek, 1957). In the testis, cadmium primarily targets Sertoli cells, perturbing the TJ‐barrier function in vitro (Chung and Cheng, 2001; Janecki et al., 1992) and the BTB and apical ES integrity in vivo (Hew et al., 1993b; Lui et al., 2003d; Setchell and Waites, 1970). As such, the use of cadmium administration in vivo to induce BTB damage has been widely used to study BTB dynamics. This method is convenient because a single low dose of cadmium salt, usually CdCl2 administered via intraperitoneal (IP) injection at 1–3 mg/kg body weight, is enough to induce irreversible damages to the BTB (Hew et al., 1993a,b; Lui et al., 2003d; Wong et al., 2004, 2005a). Indeed, results
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from these studies and others using diVerent environmental toxicants (see following) have consistently shown that the BTB is exceedingly sensitive to toxicants versus the endothelial TJ‐barrier found in blood vessels. The fact that the BTB is composed of coexisting TJ and AJ (and desmosome‐like junctions), which are likely being used to safeguard the barrier function during extensive junction restructuring that facilitates germ‐cell migration across the BTB (see Section I), may be the very reason that makes it so vulnerable to TJ/AJ disruptors, such as cadmium. For instance, in the microvascular endothelial TJ‐barrier, AJ plaques that are composed of E‐ cadherin are the primary target of cadmium (for a review, see Prozialeck, 2000). However, they are being sealed from the environmental toxicants because they are morphologically located behind the TJ barrier, making them less accessible to the cadmium. But the BTB, unlike the endothelial TJ‐barrier in the microvessel, is morphologically closest to the basement membrane, a modified form of ECM (for reviews, see Dym, 1994; Siu and Cheng, 2004). As such, E‐cadherin and N‐cadherin that are intermingled with TJ proteins at the BTB are readily exposed to cadmium, which conveniently induces AJ (as well as TJ) disruption. However, the mechanism that transmits signals from the BTB to the adluminal compartment to induce the disruption of the apical ES remains to be identified. Nonetheless, this model has been used in studies to tackle the signaling pathways that regulate many of the proteins (e.g., cytokines, proteases, protease inhibitors, and TJ‐ and AJ‐associated proteins) crucial to BTB restructuring during spermatogenesis (Wong et al., 2004, 2005a).
B. Other Potential Models Besides cadmium, other reagents are also known to target the BTB. For instance, the local injection of aqueous glycerol solution to the testis has been shown to reduce testicular weight and suppress spermatogenesis in rats (Igdoura and Wiebe, 1994; Weinbauer et al., 1987; Wiebe and Barr, 1984), which is possibly mediated via a disruption of the BTB (Eng et al., 1994). These eVects apparently are caused by a disruption of the TJ‐associated microfilaments and microtubules, altering occludin distribution in the seminiferous epithelium in vivo (Wiebe et al., 2000). Another model to study BTB dynamics in vivo is to administer a synthetic peptide corresponding to the second extracellular loop of the occludin molecule to perturb the BTB integrity (Chung et al., 2001; Wong and Gumbiner, 1997). It is believed that the interaction between the peptide and the ECD of endogenous occludins possibly releases the interlock structure that is essential for TJ assembly. For instance, when this peptide was administered to rats via intratesticular
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injection, the BTB was disrupted, as demonstrated by micropuncture technique. Unlike other models, the occludin peptide‐mediated BTB disruption is reversible, making this model extremely useful to study BTB dynamics (Chung et al., 2001). Other studies have also shown that a deficiency in vitamin A can induce Sertoli cell TJ disruption in rats (Huang et al., 1988; Ismail and Morales, 1992; Morales and Cavicchia, 2002). Furthermore, vitamin A‐deficient diet can lead to a depletion of virtually all stages of germ cells, except spermatogonia, from the seminiferous epithelium (for a review, see Kim and Wang, 1993). The BTB in vitamin A–deficient rat was leaky to intercellular tracers such as lanthanum when examined by electron microscopy (Huang et al., 1988; Morales and Cavicchia, 2002). Interestingly, this BTB disruption and spermatogenic arrest can be reversed by vitamin A replacement with full recovery within 16 wk (Morales and Cavicchia, 2002). Another means to induce BTB restructuring is via local administration of cytochalasin D, a fungal metabolite that aVects actin polymerization (for a review, see Cooper, 1987). For instance, an intratesticular injection of cytochalasin D to adult rats disrupted basal ES when examined by electron microscopy, and tracer experiments illustrated a leaky BTB (Weber et al., 1988). BTB can also be disrupted by flutamide, an antiandrogen that interferes with the binding of androgen to its receptors. When administrated to rats orally, flutamide reduced the expression of occludin in the testis and perturbed TJ integrity (Gye and Ohsako, 2003), possibly because of a loss of building blocks that construct TJs. Furthermore, Bisphenol A and tert‐octylphenol (two environmental toxicants with estrogen‐like activity) were shown to inhibit the expression of occludin and N‐cadherin in Sertoli cells in vitro (Fiorini et al., 2004), illustrating that both toxicants may induce BTB disruption in vivo. Undoubtedly, the potential of these reagents to be developed into useful models for studying BTB dynamics remains to be explored.
IV. Regulation of BTB Dynamics A. Does the BTB Restructuring Share Similar Features Pertinent to Cellular Movement with Cell‐Matrix Junction Dynamics? In the seminiferous epithelium, TJs are present proximally to the basolateral region of adjacent Sertoli cells closest to the ECM (i.e., basement membrane), rather than apically, like the case in most epithelia (for reviews, see Cheng and Mruk, 2002; Dym and Fawcett, 1970) (Fig. 1). Furthermore, the basement membrane is crucial to the integrity of the Sertoli cell TJ‐barrier
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function. For instance, Sertoli cells appear morphologically normal and can maintain proper TJ‐barrier function in vitro only when they were cultured on an artificially reconstituted basement membrane (Matrigel) (Hadley et al., 1985). Studies have also shown that a disruption of the ECM proteins, such as the use of an antibody against collagen in vitro (Siu et al., 2003), can reversibly perturb the Sertoli cell TJ functionality. Such tight morphological association between the BTB and the basement membrane, and findings that illustrate the significance of ECM proteins to Sertoli cell TJ‐barrier function, seemingly suggest that the Sertoli cell–ECM interactions play a pivotal role to BTB dynamics. Interestingly, extensive restructuring of the BTB that facilitates germ‐ cell movement during spermatogenesis, also reminiscent of the events of cell migration at the cell‐matrix interface. The classic examples of cell migration involving ECM turnover include the transmigration of leukocytes from microvessels to the site of injury across the endothelial TJ‐barrier during an inflammatory response, and the metastasis of malignant cells during tumor invasion. Under these conditions, ECM undergoes extensive restructuring under the influence of extracellular factors, such as cytokines and proteases. These factors are either produced by the migrating cells or released from the ECM, which in turn modifies the status of the ECM to facilitate cell movement (for reviews, see Ahmad et al., 2002; Lazar‐Molnar et al., 2000; Vaday and Lider, 2000). While developing germ cells are not migrating cells per se (e.g., they do not change cell shape as they translocate across the seminiferous epithelium during spermatogenesis, and they do not display distinctive lamellipodia similar to fibroblasts, leukocytes, and macrophages), they do possess many of the mechanochemical molecules uniquely found in migrating cells (e.g., Rho GTPases, GTPase‐activating proteins, and the downstream eVectors such as Rho kinases, LIM kinases, and cofilin [Guttman et al., 2004; Lui et al., 2005] [for reviews, see Lui et al., 2003a; Takahashi et al., 2003]). It is likely that there are cross‐talks between Sertoli and germ cells in the epithelium in which Sertoli cells provide the mechanical ‘‘force’’ to facilitate germ cell movement, utilizing the unique Sertoli–germ‐ cell anchoring devices (for a review, see Mruk and Cheng, 2004b). Sertoli– germ‐cell interactions also regulate BTB dynamics, in which signals (e.g., cytokines released from germ cells) are sent to Sertoli cells to facilitate germ‐ cell migration (for reviews, see Cheng and Mruk, 2002; Mruk and Cheng, 2004b). Such interactions are likely to be mediated via similar mechanisms and/or molecules employed at the cell–ECM interface. This is obviously a research area of high priority because a thorough understanding of this event can lead to new approaches to compromise germ‐cell movement in the epithelium, which disrupt spermatogenesis and induce infertility as a means of male contraception.
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B. Roles of Cytokines and their Downstream Signaling Pathways in Regulating BTB Dynamics Cytokines are secretory products of Sertoli and germ cells in the seminiferous epithelium (for reviews, see Griswold, 1988; Lui et al., 2003b; Skinner, 1993). They play versatile roles in spermatogenesis, such as regulating protein synthesis, modulating cell division and diVerentiation, and promoting cell survival. Studies have shown that cytokines in the seminiferous epithelium are uniquely important for regulating BTB dynamics via their eVects on the levels of TJ‐ and AJ‐integral membrane proteins, proteases, protease inhibitors, and ECM proteins in the seminiferous epithelium. Indeed, the modulation of in vivo barrier functions in mammalian epithelia by cytokines has been reported under pathophysiological conditions. For instance, the BBB of juvenile rats can be compromised by interleukin (IL)‐1 , possibly a result of inflammatory response (Anthony et al., 1997). The BRB in rabbits is also disrupted by treatments with IL‐1 or tumor necrosis factor (TNF)‐ (Claudio et al., 1994), whereas another cytokine namely transforming growth factor (TGF)‐ can perturb retinal endothelial barrier in vitro (Behzadian et al., 2001). Furthermore, the breakdown of the BRB caused by diabetes is initiated by vascular endothelial growth factor (VEGF) (Qaum et al., 2001). Interestingly, while cytokines were shown to aVect junction integrity in diVerent cell types and tissues (for a review, see Walsh et al., 2000), virtually no reports were found in the literature that investigated the action of cytokines in the testis regarding the BTB regulation until very recently. The most extensively studied cytokines that are known to be involved in BTB dynamics thus far are TNF‐ and TGF‐ 3. 1. TNF‐a TNF‐ is a proinflammatory cytokine secreted by germ cells, predominantly pachytene spermatocytes and round spermatids, and Sertoli cells (De et al., 1993; Siu et al., 2003), whereas its receptors are restricted to Sertoli cells (De et al., 1993). During the assembly of Sertoli cell TJ‐barrier in vitro, the amount of TNF‐ produced by Sertoli cells declines significantly, indicating that it can downregulate the Sertoli cell TJ‐barrier function (Siu et al., 2003). Indeed, the inclusion of recombinant TNF‐ in rat Sertoli cell cultures can perturb the integrity of the TJ barrier, as shown in experiments quantifying the transepithelial electrical resistance (TER) across the Sertoli cell monolayer (Siu et al., 2003). This can be caused by the direct eVect of TNF‐ on the TJ‐associated structural protein complexes, because the level of occludin is significantly reduced after TNF‐ treatment (Siu et al., 2003). Besides, it is shown that TNF‐ can inhibit claudin‐11 expression in cultured mouse Sertoli cells (Hellani et al., 2000). Another possible route by which TNF‐
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aVects TJ‐barrier function is through its eVect on follicle‐stimulating hormone (FSH). Because FSH can enhance the assembly of Sertoli cell TJ‐ barrier (Janecki et al., 1991), TNF‐ , a known antagonist of FSH (Mauduit et al., 1993), may block the action of FSH on Sertoli cells and lead to the loss of the barrier integrity. Furthermore, subsequent studies have shown that TNF‐ can mediate its eVects via mitogen‐activated protein kinase (MAPK) pathways and the integrin‐linked kinase (ILK)/glycogen synthase kinase‐3 (GSK‐3 ) pathway (De Cesaris et al., 1998; Siu et al., 2003). This in turn aVects the homeostasis of proteases and protease inhibitors and subsequently the nature of the ECM adjacent to the BTB (see later). Although the precise mechanism by which TNF‐ aVects BTB dynamics in vivo remains to be elucidated, it has been reported that the intraperitoneal injection of CdCl2 in rats can elevate the TNF‐ level in the testis (Chen et al., 2003), further strengthening the postulate that this cytokine is involved in BTB restructuring in vivo. Studies have illustrated that TNF‐ may mediate its eVect via aVecting the homeostasis of proteases and protease inhibitors in the seminiferous epithelium, which in turn modulates BTB integrity in vivo (Wong et al., 2005a). 2. TGF‐b3 Another cytokine that exhibits a similar regulatory eVect, yet via diVerent signaling pathways, on the Sertoli cell TJ‐barrier as TNF‐ is TGF‐ 3. TGF‐ 3 is the most abundant form of TGF‐ in the testis, which expression is the highest at the onset of puberty in rats (Mullaney and Skinner, 1993). It is produced by Sertoli cells (Watrin et al., 1991), and its level declines when Sertoli cell TJ‐barrier is being assembled in vitro (Lui et al., 2001). It is also a product of premeiotic germ cells, such as spermatogonia and early spermatocytes, in adult boar and rat testes (Caussanel et al., 1997; Lui et al., 2003c). The receptors (type I and type II) for TGF‐ 3 are found predominantly in Sertoli cells in the rat testis (Le Magueresse‐Battistoni et al., 1995). Earlier studies have shown that an inclusion of recombinant TGF‐ 3 in Sertoli cells cultured in vitro can lead to a significant drop of TER across the cell monolayer (Lui et al., 2001). This drop in TER, an indication of a disruption of Sertoli cell TJ, is accompanied by a decline in the protein levels of several TJ‐associated proteins, including occludin, ZO‐1, and claudin‐11 in Sertoli cells (Lui et al., 2001). Subsequent studies have shown that TGF‐ 3 mediates its eVects on Sertoli cell TJs via the p38 MAPK pathway (Lui et al., 2003c). The use of SB202190, a specific p38 MAPK inhibitor, can block the disruptive eVects of TGF‐ 3 on the Sertoli cell TJ‐barrier function, confirming the significance of the p38 MAPK in the regulation of Sertoli cell TJ dynamics (Lui et al., 2003c). More importantly, these observations in vitro have been validated in vivo using the CdCl2‐treated rats as a model. For instance, it was shown that TGF‐ 3 was significantly induced in the testis during
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CdCl2‐induced BTB damage, concomitant with a reduction in the levels of occludin and ZO‐1 at the site of the BTB (Lui et al., 2003d; Wong et al., 2004). Perhaps the most important of all, by blocking the p38 MAPK activity locally in the testis via an intratesticular injection of SB202190 prior to CdCl2 administration, the loss of occludin, ZO‐1, and other BTB‐associated adhesion molecules during the CdCl2‐induced BTB damage was indeed significantly reduced (Lui et al., 2003d; Wong et al., 2004). Collectively, these data suggest that TGF‐ 3 regulates BTB dynamics in vivo via the p38 MAPK pathway, modulating the steady‐state protein levels of TJ‐ and AJ‐associated proteins. It is therefore important that studies of these two cytokines regarding their role in BTB regulation be expanded in the future, which should include a detailed examination of their corresponding signaling cascades such as the transcriptional regulation of the BTB‐associated proteins. Furthermore, these two cytokines may have cross‐talks and/or converged at specific point(s), which should be resolved in future studies.
C. Interplay of Proteases and Protease Inhibitors Extracellular proteolysis is intimately associated with the events of tissue remodeling and cell migration, but it usually is restricted to the cell‐matrix interface in tissues and epithelia except under pathophysiological conditions like inflammation (for reviews, see DeClerck et al., 2004; Murphy and Gavrilovic, 1999; Stamenkovic, 2003). When the homeostasis between proteases and protease inhibitors is disturbed, the biochemical composition of ECM (i.e., the basement membrane) changes. This leads to the opening of the cell‐matrix interface, facilitating cell movement. For instance, the BBB can be disrupted by gelatinases (also known as type IV collagenases or matrix metalloproteinase [MMP]‐2/9) in patients suVering from acute multiple sclerosis (MS), stroke, and head trauma, possibly by disrupting basal lamina around cerebral blood capillaries (for a review, see Avolio et al., 2003; Rosenberg et al., 1992, 1996). Another study has shown that tissue‐type plasminogen activator, a serine protease, caused the opening of the BBB in mice suVering from experimentally induced cerebral ischemia (Yepes et al., 2003). The BRB is also susceptible to proteases that degrade ECM (Behzadian et al., 2001; Pino, 1987). Occasionally, extracellular proteolysis can occur at the cell–cell interface that helps to degrade junction components to facilitate the barrier’s disassembly. For example, when neutrophils transmigrate across the vascular endothelial TJ barrier during inflammation, they release proteases to cleave cadherins at the cell–cell interface (Hermant et al., 2003). Also, an activation of gelatinase B can be detected at the BBB when mice suVer from experimentally induced cerebral injury, which cleaves ZO‐1 and leads to BBB disruption (Asahi et al., 2001).
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However, while most of these cases found in other epithelia take place under pathophysiological conditions, extracellular proteolysis is essential for maintaining normal spermatogenesis in the seminiferous epithelium of the testis. A large variety of proteases and protease inhibitors are present, with some of them predominantly or even exclusively in the testis (Table I). This also illustrates their critical roles in the restructuring event that occurs in the seminiferous epithelium during spermatogenesis, which include the timely ‘‘opening’’ and ‘‘closing’’ of the BTB to facilitate germ‐cell movement (for reviews, see Cheng and Mruk, 2002; Fritz et al., 1993; Monsees et al., 1997). Predictably, proteases can either be directly involved in the removal of junction components during BTB disassembly to facilitate preleptotene/ leptotene spermatocyte migration, or they can indirectly regulate junction turnover by activating other biologically active molecules, such as growth factors, cytokines, and other ECM components (e.g., collagens). At the same time, protease inhibitors limit the activity of proteases so that proteolysis can be confined to a specific microenvironment within the epithelium. Studies have implicated the participation of several proteases and protease inhibitors in the regulation of BTB dynamics, including cathepsins (cysteine proteases), plasminogen activators (PAs, serine proteases), gelatinases (metalloproteinases), cystatin C (a cysteine protease inhibitor), protein C inhibitor (PCI, a serine protease inhibitor), tissue inhibitors of metalloproteinases (TIMPs), and 2‐macroglobulin ( 2‐MG, a nonspecific protease inhibitor), which are to be discussed herein. 1. Cathepsins and Cystatin C Cathepsins are lysosomal proteases. They are ubiquitously expressed cysteine proteases, except for cathepsin A and D, which are serine and aspartic proteases, respectively (for reviews, see Berdowska, 2004; Turk et al., 2001). In the testis, cathepsins that have been positively identified include cathepsin A (Luedtke et al., 2000), cathepsin B (Scott et al., 1987), cathepsin C (Chung et al., 1998), cathepsin D (Igdoura et al., 1995), cathepsin F (Wang et al., 1998), cathepsin K (Anway et al., 2004), cathepsin L (Erickson‐Lawrence et al., 1991), cathepsin S (Chung et al., 1998), and cathepsin V (Bromme et al., 1999). By far, cathepsin L is the best‐studied cathepsin in the rat testis. Although cathepsin L is a lysosomal protease, it is also a secretory protease that is produced by Sertoli cells in the form of an inactive precursor previously called CP‐2 (Erickson‐Lawrence et al., 1991; ZabludoV et al., 1990) or CMB‐2 (Lee et al., 1986) in the seminiferous epithelium. Its mRNA can also be found in spermatocytes and spermatogonia (Chung et al., 1998). The production of this enzyme by Sertoli cells is regulated by germ cells and vice versa (Chung et al., 1998; ZabludoV et al., 2001), but it is not expressed in testes with Sertoli‐cell‐only syndrome (Gye and Kim, 2004). A study has
Table I
Proteases and Protease Inhibitors Found in the Seminiferous Epithelium and their Cellular Association in Mammalian Testes*
Classes of proteases and protease inhibitors Cysteine‐proteases
Members identified in the testis Cathepsins –Cathepsin L –Cathepsin K Calpain‐1 and ‐2 Caspases –procaspase 3 and 6
Cysteine‐protease inhibitors Serine‐proteases
Serine‐protease inhibitors Aspartic‐proteases
Cystatin C Testis‐specific calpastatin Plasminogen activators (PAs) –tPA –uPA Cathepsin A Tissue Kallikreins Testisin Testis‐specific seine protease (TESSP)‐1 Plasminogen activator inhibitor (PAI)‐1 Protein C inhibitor (PCI) Cathepsin D
Sources and localizations in the seminiferous epithelium
Sertoli cells, spermatocytes, spermatogonia Sertoli cells Sertoli cells Sertoli cells and elongated spermatids Pachytene spermatocytes and spermatids Sertoli cells, germ cells Postmeiotic germ cells Sertoli cells Sertoli cells Sertoli cells Round and elongate spermatids Elongated spermatids Spermatogonia and spermatocytes Sertoli cells Early and elongated spermatids Sertoli cells and spermatids
References See section IV.C.1. Erickson‐Lawrence et al., 1991 Anway et al., 2004 Sultana et al., 2003 Tesarik et al., 2002 Omezzine et al., 2003 Esnard et al., 1992; Tsuruta et al., 1993 Somwaru et al., 2004 See Section IV.C.2. Hettle et al., 1986 Hettle et al., 1986 Luedtke et al., 2000 Monsees et al., 1999 Nakamura et al., 2003 Takano et al., 2005 Le Magueresse‐Battistoni et al., 1998 Odet et al., 2004 Igdoura et al., 1995
Metalloproteases
Matrix metalloproteinases (MMPs) –Gelatinase A (MMP‐2) –Gelatinas B (MMP‐9) –Stromelysin‐1 (MMP‐3)
Metalloprotease inhibitors Other protease inhibitors
*
Sertoli cells Sertoli cells, spermatocytes, round spermatids Sertoli cells
–Membrane‐type‐1 MMP (MT1‐MMP) –Matrilysin (MMP‐7) A disintegrin and metalloprotease (ADAM) –Fertilin (ADAM‐2)
Elongating and elongated spermatids
–Cyritestin (ADAM‐3)
Spermatids
Tissue inhibitors of MMP (TIMPs) –TIMP‐1 –TIMP‐2 –TIMP‐4 2‐Macroglobulin Cystatin‐related epididymal spermatogenic (CRES) subgroup of family 2 cystatins –CRES –Testatin –Cres 3
Sertoli cells, germ cells Spermatocytes
Sertoli Sertoli Sertoli Sertoli
cells, germ cells cells cells, germ cells cells
Round and elongating spermatids Sertoli cells Sertoli cells
See Section IV.C.3. Ailenberg et al., 1991 Hoeben et al., 1996; Siu et al., 2003 Longin and Le Magueresse‐ Battistoni, 2002 Longin et al., 2001 Rudolph‐Owen et al., 1998
For a review, see PrimakoV and Myles, 2000 McLaughlin et al., 1997 Forsbach and Heinlein, 1998 See Section IV.C.3. Mruk et al., 2003; Ulisse et al., 1994 Grima et al., 1996; Ulisse et al., 1994 Robinson et al., 2001 Cheng et al., 1990 For a review, see Cornwall and Hsia, 2003
Cornwall and Hann, 1995 Tohonen et al., 1998 Hsia and Cornwall, 2003
This list is not intended to be exhaustive, and readers are encouraged to find additional information in the cited reviews and reports.
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reported that mice expressing inactive cathepsin L (the furless strain), although fertile, exhibited abnormal spermatogenesis (Wright et al., 2003). The development of germ cells beyond pachytene spermatocytes in furless mice is significantly disturbed, which can possibly be an indication of BTB damage. This possibility should be investigated in future studies. Also, a disruption of BTB by cadmium in vivo is accompanied by an induction in cathepsin L, suggesting that its activity is needed to regulate BTB disassembly (Wong et al., 2004). This cadmium‐induced cathepsin L production in the epithelium apparently is mediated by TGF‐ 3 via the p38 MAPK pathway, given that a blockade of this pathway by SB202190 can alleviate the CdCl2‐induced cathepsin L production in the testis (Wong et al., 2004). Another cathepsin that shares a similar expression profile and localization pattern in the seminiferous epithelium of rat testes as cathepsin L is cathepsin K (Anway et al., 2004). However, whether it plays any role in junction dynamics in the testis remains unknown. While cathepsins are abundantly expressed in the testis, one of their specific inhibitors, namely cystatin C, is also present there. Cystatin C is a product of Sertoli cells (Esnard et al., 1992), but its truncated transcript has also been identified in germ cells (Tsuruta et al., 1993). In the male genital tract, cystatin C is the most potent inhibitor of cathepsin L (Peloille et al., 1997). Indeed, when cathepsin L was induced in the cadmium model, it was followed by an increase in the level of cystatin C, illustrating that cystatin C may be involved in the control of proteolysis contributed by cathepsin L during CdCl2‐induced BTB disruption (Wong et al., 2004). 2. PAs and PCI Tissue‐type PA (tPA) and urokinase‐type PA (uPA) are two of the most well‐studied serine proteases in the testis. They are not only active proteases per se, but also function as activators of other proteases and growth factors. Both proteins are secretory products of Sertoli cells, and their expression profile, localization, and hormonal regulation in the seminiferous epithelium have been well characterized (for a review, see Fritz et al., 1993). Under normal physiological conditions, Sertoli cells predominantly produce uPA, whereas tPA is the dominant type in FSH‐stimulated Sertoli cells (Hettle et al., 1986). Moreover, the activity of uPA, but not tPA, is largely regulated by germ cells in the seminiferous epithelium (Mruk et al., 1997; Penttila et al., 1994). This is possibly because uPA works best when it binds to its receptor (u‐PAR), which is exclusively associated with spermatocytes and spermatids in the seminiferous epithelium (Odet et al., 2004). These diVerences illustrate the diverse roles of the two PAs in spermatogenesis. Nevertheless, both PAs are important for maintaining normal reproduction, because mice with double deletion of tPA and uPA produce significantly
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fewer pups than normal mice (Carmeliet et al., 1994). The first piece of evidence supporting that PAs might be involved in BTB dynamics is derived from studies using Sertoli cells cultured on bicameral units with peritubular myoid cells. It was observed that when the level of PAs (most likely tPA) produced from Sertoli cells was induced, the integrity of the Sertoli cell TJ barrier was perturbed (Ailenberg and Fritz, 1989). Subsequent studies using CdCl2 to perturb inter‐Sertoli cell TJ in vitro have demonstrated that uPA was induced transiently at the time when the TJ barrier was disrupted and later when it reassembled, suggesting a complex role of uPA during junction restructuring (Chung and Cheng, 2001). It is obvious that in order to counterbalance the activity of PAs, their inhibitors are likely present in the seminiferous epithelium. Indeed, PCI, a serine protease inhibitor that can inhibit the activity of PA, is found in the epithelium (Potempa et al., 1994). PCI shares a similar expression profile to uPA and uPAR in developing mouse testes, and while the levels of both PCI and uPAR increase in the testis during aging in mice, no induction in uPA activity is detected (Odet et al., 2004). It is therefore highly possible that uPA is a target protease of PCI (Odet et al., 2004). More important, male PCI / mice were infertile with abnormal spermatogenesis and a disrupted BTB (Uhrin et al., 2000). It was suggested that the loss of BTB integrity in PCI / mice is a result of uncontrolled proteolysis at the site, because the absence of PCI was accompanied by an increase in the activity of proteases, including urokinase and plasmin (Uhrin et al., 2000). This further supports the fact that PAs are involved in BTB regulation. Besides PCI, PA inhibitor‐1, the primary inhibitor of the two PAs, has also been identified in the testis and is a product of Sertoli cells (Le Magueresse‐Battistoni et al., 1998), although much research is needed to dissect its significance in BTB dynamics. 3. Gelatinases and TIMPs Gelatinases, or type IV collagenases, refer to two members of the zinc‐ dependent MMP family: gelatinase A (MMP‐2) and gelatinase B (MMP‐9). Although almost every known MMP has been found in the testis (for a review, see Nuttall et al., 2004; Siu and Cheng, 2004), gelatinases are among some of the best characterized MMPs. Gelatinases are known to regulate junction restructuring at other barriers such as the BBB as mentioned previously. In the testis, both gelatinases are secreted by Sertoli cells as proenzymes, with gelatinase B also associating with germ cells including spermatocytes and round spermatids (Ailenberg et al., 1991; Longin et al., 2001; Robinson et al., 2001; Siu et al., 2003). These proenzymes require other proteases, such as membrane‐type 1‐MMP (MT1‐MMP) and stromelysin‐1 (MMP‐3), for activation to become fully functional (Longin et al., 2001; Ogata et al., 1992). An in vitro study has shown that both gelatinases are
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involved in the turnover of Sertoli cell AJs and TJs (Siu et al., 2003). For instance, both gelatinases are induced during the assembly of Sertoli cell TJ barrier in vitro and gelatinase B is also induced by adding TNF‐ , a Sertoli cell TJ disruptor (see Section IV.B) to the cultures, suggesting their roles in junction restructuring at the barrier. It is suggested that the TNF‐ ‐induced junction disruption is caused in part by the cleavage of type IV collagens and other ECM proteins at the basement membrane by activated gelatinase B (Siu et al., 2003). This in turn releases active fragments from ECM proteins (e.g., NC1 domain from type IV collagen), which further promote the disruption of junctions. On the other hand, the level of gelatinase A in the testis can be significantly induced during the in vivo CdCl2‐induced BTB disruption, suggesting its participation in eliciting barrier damage in the seminiferous epithelium (Wong et al., 2004). Similar to other MMPs, inhibitors for gelatinases are members of a family of protease inhibitors named TIMPs. There are four TIMPs known to date, which are all present in mammalian testes. Among them, TIMP‐1, TIMP‐2, and TIMP‐4 have been identified within the seminiferous epithelium behind the BTB (Grima et al., 1996; Gronning et al., 2000; Robinson et al., 2001; Siu et al., 2003; Ulisse et al., 1994). Both TIMP‐1 and TIMP‐2 can be induced when Sertoli cell TJ barrier is being assembled in vitro, similar to the expression pattern of gelatinases (Longin et al., 2001; Siu et al., 2003). Besides, the production of both TIMPs and gelatinases by Sertoli cells can be induced by follicle‐stimulating hormone (Gronning et al., 2000; Longin and Le Magueresse‐Battistoni, 2002; Ulisse et al., 1994). Moreover, TIMP‐1 can be induced by TNF‐ in Sertoli cell cultures (Siu et al., 2003), as well as by interleukin‐1 , another cytokine released by Sertoli cells and possibly germ cells (Gronning et al., 2000). These findings thus support the notion that TIMPs are involved in BTB restructuring, with gelatinases as their potential target substrates. The physiological importance of these works can be significantly enhanced by using cell‐specific knockout mice, which should be considered in future studies. 4. a2‐MG 2‐MG is one of the earliest discovered and most studied protease inhibitors in the testis (for a review, see Fritz et al., 1993). It inhibits all classes of proteases via an entrapment mechanism (for a review, see Baker et al., 2002; Sottrup‐Jensen, 1989). It also binds to TGF‐ s (O’Connor‐McCourt and Wakefield, 1987) and other growth factors and hormones with high aYnity, possibly regulating their bioavailability (for reviews, see Armstrong and Quigley, 1999; Feige et al., 1996; James, 1990). In the seminiferous epithelium behind the BTB, 2‐MG is a secretory product of Sertoli cells (Cheng et al., 1990; Zhu et al., 1994), where its expression is regulated diVerently
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from that of the same protein produced by hepatocytes in the liver (Cheng et al., 1990; Stahler et al., 1991). Although 2‐MG is an acute‐phase protein in the circulation, it does not respond to induced inflammation in the testis (Braghiroli et al., 1998; Stahler et al., 1991). However, the basal physiological level of 2‐MG in the seminiferous epithelium is significantly higher than that of the liver unless the animal is subjected to experimentally induced inflammation (Braghiroli et al., 1998; Li et al., 1994), suggesting that 2‐MG in the testis is constantly at an ‘‘elevated’’ level, possibly for protecting the epithelium and the BTB from undesired proteolysis. Studies by immunohistochemistry have localized 2‐MG to the BTB and apical ES sites in the seminiferous epithelium, indicating its likely participation in junction dynamics at these two sites (Zhu et al., 1994). The involvement of 2‐MG in junction restructuring particularly at the BTB has been demonstrated in an in vivo study using cadmium to induce BTB restructuring (Wong et al., 2004, 2005a). 2‐MG is significantly induced, particularly at the site of the BTB, during the CdCl2‐induced BTB disruption (Wong et al., 2004). Furthermore, it is structurally associated with the nectin/afadin protein complex, but not the cadherin/catenin complex, in the testis, indicating that it is intimately involved in the nectin‐based cell adhesion function. 2‐MG is also structurally associated with TGF‐ 3 in the testis, confirming its ability to bind to cytokines and its possible role in regulating the pool of bioavailable cytokines in the seminiferous epithelium (Wong et al., 2004). Subsequent studies have shown that the induction of this protease inhibitor by cadmium was regulated through c‐Jun N‐terminal protein kinase (JNK), another MAPK, instead of p38 MAPK (Wong et al., 2005a). Studies using dimethylaminopurine (DMAP), a specific inhibitor of JNK, have shown that the CdCl2‐induced 2‐MG could indeed be suppressed by blocking endogenous JNK activity, while at the same time the BTB became more susceptible to the CdCl2‐mediated damage (Wong et al., 2005a). Taken collectively, these results illustrate that 2‐MG is a crucial component in the seminiferous epithelium that protects the BTB from cellular fall‐outs pertinent to junction restructuring during spermatogenesis. For instance, it may function as a prominent protease inhibitor to limit excessive proteolysis during junction restructuring in the epithelium and/or act as a regulator to determine the bioavailability of cytokines in the epithelium that can elicit junction disassembly (e.g., TGF‐ 3 and TNF‐ ). Indeed, 2‐MG can partially abolish the disruptive eVect of TGF‐ 3 on the Sertoli cell TJ‐barrier integrity in vitro when the TJ‐barrier function is quantified by TER (Wong and Cheng, unpublished observations). Although all these observations strongly support the notion that 2‐MG is an important regulator of the BTB, it is noteworthy that 2‐MG / mice are fertile (Umans et al., 1995, 1999). This suggests that the role of 2‐MG can be substituted by other protease inhibitor(s) in the seminiferous epithelium. However, 2‐MG is apparently important
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for protecting the seminiferous epithelium from assaults of environmental stresses, such as cadmium toxicity. This possibility is strengthened by the observation that 2‐MG / mice are highly susceptible to diet‐induced pancreatitis even though the mutation per se did not aVect vitality (Umans et al., 1999). D. Unique Physiological Features of the BTB Pertinent to its Regulation During Spermatogenesis In short, the preceding sections summarize some of the latest findings in the field regarding the regulation of BTB dynamics. It is increasingly clear that because of the unusual morphological features of the BTB versus other barriers (e.g., BBB, BRB) (Fig. 2), some of the generally accepted cell physiological phenomena that occur in other epithelia do not apply to the BTB. For instance, cytokines and proteases/protease inhibitors as reviewed herein play a rather significant role in mediating BTB dynamics in the testis, but these molecules by and large are being restrictedly used in cell‐matrix restructuring under pathophysiological conditions such as inflammation and tumor metastasis. Furthermore, a disruption of AJ in other epithelia can usually lead to a loss of TJ integrity (Gassler et al., 2001; Guo et al., 2003; Venkiteswaran et al., 2002), yet AJ restructuring in the seminiferous epithelium pertinent to spermatogenesis does not disrupt the BTB integrity. Indeed, even when the epithelium is under the assault of chemicals, such as AF‐2364 (1‐[2,4‐dichlorobenzyl]‐indazole‐3‐carbohydrazide), that induce germ cell loss from the epithelium by perturbing Sertoli–germ cell AJ, the BTB integrity remains relatively intact (for a review, see Mruk and Cheng, 2004b). This observation has been confirmed using another model of AJ restructuring where Sertoli–germ cell AJ is perturbed (Wong et al., 2005b; Zhang et al., 2005) (for a review, see McLachlan et al., 2002) without compromising the BTB integrity (Xia et al., 2005) via a suppression of intratesticular androgen level. It is obvious that much research is needed to identify all the crucial players that modulate BTB function so that functional experiments can be designed to investigate BTB regulation.
E. An Integrated Model of BTB Regulation During Spermatogenesis As it is reviewed previously, cytokines and proteases are known to regulate cell‐matrix restructuring events to facilitate cellular movement. Because many of these molecules are found in the seminiferous epithelium, it is reasonable to postulate that the mechanism(s) utilized by the testis to regulate the ‘‘opening’’ and ‘‘closing’’ of cell–cell junctions at the BTB is similar
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to that at the cell‐ECM interface. When migrating cells, such as leukocytes and macrophages, traverse the ECM, they release signals (e.g., cytokines) to induce changes at the front‐ and rear‐ends of the cell via molecular switches (e.g., Rho‐GTPases) that alter the G‐actin and F‐actin ratio in the cytoplasm, creating the necessary protrusive force for cell migration. It is likely that preleptotene and leptotene spermatocytes can release TGF‐ 3 and/or TNF‐ to induce changes of the Rho‐GTPase signaling function in Sertoli cells and/or germ cells to alter the actin‐based cytoskeleton (for a review, see Lui et al., 2003a). Alternatively, cellular movement can be facilitated via degradation of ECM components by proteases released from either the basement membrane or the passing cells, whereas in the case of BTB restructuring, the release of proteases from Sertoli cells can be induced by germ cells. Proteases may also activate other biologically active molecules, including growth factors, cytokines, cell‐associated receptors, and even other proteases, causing further disruption of the ECM (for reviews, see DeClerck et al., 2004; Lauwaet et al., 2000). After crossing the ECM, migrating cells (e. g., during tumor cell metastasis) may continue to invade the endothelial barrier by producing cytokines to facilitate junction disruption or provoke proteolysis at the cell–cell interface (for a review, see Edens and Parkos, 2000). Similar to these other epithelia, it is possible that germ cells release signals via the secretion of cytokines, which in turn aVects the homeostasis of proteases and protease inhibitors in Sertoli cells and in the basement membrane that leads to BTB disassembly (Fig. 3). This model is best demonstrated in studies using cadmium to induce BTB disruption (Wong et al., 2004, 2005a). When the BTB is under the assault of cadmium toxicity, cytokines, proteases, and protease inhibitors in the seminiferous epithelium are significantly induced. For instance, the cadmium‐induced TGF‐ 3 production elicits an inhibition on the steady‐state level of junction‐associated proteins, which are necessary for maintaining BTB integrity. TGF‐ 3 also induces proteolysis by increasing the level of proteases in the seminiferous epithelium, such as cathepsin L, via the p38 MAPK pathway, leading to junction disassembly at the BTB (Fig. 3). TNF‐ , another disruptor of Sertoli cell TJ barrier and an activator of p38 MAPK and the ILK/GSK‐3 signaling pathway in Sertoli cells in vitro (De Cesaris et al., 1998; Siu et al., 2003), may also contribute to the BTB disruption in vivo via these pathways (Fig. 3). At the time when the BTB ‘‘opens,’’ protease inhibitor 2‐MG is being induced, likely to be used to limit excessive proteolysis and regulate the bioavailability of TJ‐disrupting cytokines. This induction of 2‐MG, however, is mediated via the JNK, which is activated by cytokines other than TGF‐ 3 (Wong et al., 2004, 2005a). A possible candidate for this activation is TNF‐ , which induces the production of 2‐MG in Sertoli cells cultured in vitro (Wong and Cheng, unpublished observations). In fact, TNF‐ is known to activate JNKs in cultured Sertoli cells (De Cesaris et al., 1999).
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Although the possibility that TNF‐ can induce a molecule that serves a protective function at the BTB seems unlikely, this may well be a negative feedback mechanism elicited by TNF‐ to limit its damaging eVect on the BTB (Fig. 3). Undoubtedly, this hypothesis requires vigorous investigation in future studies, but it has certainly provided a new framework to study cross‐talks between Sertoli and germ cells when germ cells traverse the BTB during spermatogenesis.
V. Concluding Remarks Findings as reviewed herein have shown that the BTB is composed of intermingling TJs and AJs and has the characteristics of both cell–cell and cell‐ECM junctions. Its restructuring is delicately regulated by intricate interactions between Sertoli and germ cells by modulating the homeostasis of cytokines, junction‐associated protein complexes, proteases, protease inhibitors, and basement membrane components. This in turn determines whether junctions at the BTB should be ‘‘closed’’ or ‘‘opened.’’ Based on available information, future functional studies can be conducted to discover diVerent approaches to manipulate the restructuring of the BTB. For instance, this can be achieved by interfering with the binding of cytokines to their receptors, by disturbing the balance between proteases and protease inhibitors, or by blocking any critical signaling molecules that aVect these events. If the disassembly and reassembly of the BTB can be deliberately manipulated, it is undoubtedly a valuable tool for delivering therapeutic Figure 3 Schematic drawing illustrating the most current model of BTB regulation in the seminiferous epithelium of the rat testis, which permits the timely ‘‘opening’’ (left panel) and ‘‘closing’’ (right panel) of the BTB to facilitate germ‐cell migration. The restructuring of TJs and AJs (e.g., basal ES) are likely determined by diVerent cytokines released from Sertoli and/or germ cells. This in turn activates the corresponding downstream signaling pathways, aVecting the homeostasis of junction‐associated proteins, proteases, and protease inhibitors, and determining the status of the barrier. When preleptotene/leptotene spermatocytes traverse the BTB at late stage VII through early stage VIII of the epithelial cycle, they transmit signals to Sertoli cells, possibly via the production of TGF‐ 3 and TNF‐ , since both cytokines are products of Sertoli and germ cells. When these cytokines bind to their corresponding receptors present on Sertoli cells (note: receptors of TGF‐ 3 and TNF‐ are restricted to Sertoli cells), their downstream signaling molecules, such as the p38 MAPK and ILK/GSK‐3 as shown here, are activated. This reduces the levels of TJ‐ and AJ‐associated proteins (e.g., occludins and cadherins), while promoting protease production (e.g., cathepsin L) in the seminiferous epithelium, resulting in the disassembly (‘‘open’’) of the BTB to facilitate the migration of spermatocytes. To limit the extent of junction disassembly, 2‐MG, the production of which is positively induced by TNF‐ and JNK, binds excessive cytokines, rendering them inactive. 2‐ MG also inhibits excessive proteolysis to prevent further damage to the TJ and AJ proteins at the cell–cell interface in the seminiferous epithelium. This in turn favors the reassembly of the junctions, ‘‘closing’’ the BTB.
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agents across the barrier. It may also be developed into a novel method for male contraception, because a malfunctioned BTB can almost certainly lead to an arrest in spermatogenesis. Reversible infertility can be achieved by either ‘‘sealing’’ the BTB to prevent germ‐cell migration during spermatogenesis, or creating a ‘‘leaky’’ BTB that promotes immature depletion of germ cells. Nevertheless, this cannot be done unless some critical questions can be answered. For example, what junction proteins are substrates of each specific protease pertinent to junction disassembly? What other factor(s) controls the cytokine‐induced losses of proteins at the BTB? What are the changes at the transcription level that lead to BTB restructuring? How is the homeostasis of proteases and protease inhibitors being fine‐tuned to maintain the ever‐changing status of the BTB? More investigations along these lines must be performed in future studies.
Acknowledgments This work was supported in part by grants from the National Institutes of Health (5 U01 HD045908 to C.Y.C. and 5 U54 HD029990, Project 3 to C.Y.C.) and the CONRAD Program (CICCR CIG 01–72). C.H.W. was supported by a postgraduate scholarship from the University of Hong Kong.
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Angiogenic Factors in the Pathogenesis of Preeclampsia Hai‐Tao Yuan,* David Haig,{ and S. Ananth Karumanchi* *Renal, Molecular, and Vascular Medicine Division, Departments of Medicine, Obstetrics and Gynecology, Beth Israel Deaconess Medical Center and Harvard Medical School, Boston, Massachusetts { Department of Organismic and Evolutionary Biology, Harvard University Cambridge, Massachusetts
I. Introduction II. Abnormal Placentation and Placental Ischemia (Stage 1) III. Systemic Endothelial Dysfunction (Stage 2) A. Excess Circulating sFlt1, Impaired VEGF Signaling, and Antiangiogenic State B. sFlt1 and Maternal–Fetal Conflict C. Speculations About the Mechanisms of Preeclampsia D. Unanswered Questions IV. Conclusions Acknowledgments References
Preeclampsia aVects 5–10% of pregnancies and is responsible for substantial maternal and neonatal morbidity and mortality. It is believed to be a two‐stage disease with an initial placental trigger with no maternal symptoms followed by a maternal syndrome characterized by hypertension, proteinuria, and endothelial dysfunction. The first stage is thought to be due to shallow cytotrophoblast invasion of maternal spiral arterioles leading to placental insuYciency. The diseased placenta in turn releases soluble angiogenic factors that induce systemic endothelial dysfunction and clinical preeclampsia during the second stage. This review will discuss the role of circulating angiogenic factors of placental origin as potential mediators of the systemic endothelial dysfunction and the clinical syndrome of preeclampsia and provide an evolutionary explanation for this phenomenon. ß 2005, Elsevier Inc.
I. Introduction Preeclampsia is characterized by the new onset of hypertension and proteinuria after 20 weeks of gestation (Roberts, 2000; Roberts and Cooper, 2001; Walker, 2000). Preeclampsia is also frequently associated with edema and Current Topics in Developmental Biology, Vol. 71 Copyright 2005, Elsevier Inc. All rights reserved.
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hyperuricemia and it usually remits when the placenta is delivered. The placenta in preeclampsia is often abnormal, with evidence of hypoperfusion and ischemia. Vascular endothelial dysfunction and microangiopathy are present in the mother, but not in the fetus. Severe complications of preeclampsia can include acute renal failure, cerebral edema, cerebral hemorrhage, seizures (eclampsia), pulmonary edema, thrombocytopenia, hemolytic anemia, coagulopathy, and liver injury—including HELLP, the syndrome of Hemolysis, Elevated Liver enzymes, and Low Platelets (Sibai et al., 2005). When preeclampsia threatens to lead to severe maternal complications, urgent delivery of the fetus and placenta is often undertaken to preserve maternal health. Although preeclampsia has been traditionally thought of as a pregnancy‐ specific syndrome, data suggests that women with a history of preeclampsia have an eightfold increased risk of cardiovascular death when followed over their lifetime (Irgens et al., 2001). Risk factors for preeclampsia include nulliparity, preexisting hypertension, obesity, diabetes mellitus, thrombophilias, and a family history of preeclampsia (Dekker, 1999). Observations to date suggest that the earliest pathological change in preeclampsia occurs in the uteroplacental circulation resulting in placental insuYciency or ischemia, which may be considered stage 1 of the disease (Roberts, 2000). In stage 2, the diseased placental tissue (ischemic placenta) in turn secretes circulating factors that cause generalized endothelial cell injury in the mother, resulting in the clinical syndrome of preeclampsia (Roberts, 2000; Roberts et al., 1989). Current understanding of the pathogenesis of preeclampsia will be reviewed with an emphasis on evidence that an imbalance in circulating angiogenic factors (Bdolah et al., 2004) and their interaction with the maternal vasculature may be responsible for the clinical phenotype of preeclampsia.
II. Abnormal Placentation and Placental Ischemia (Stage 1) It is widely accepted that the placenta plays a central role in the pathogenesis of preeclampsia as it occurs only in the presence of the placenta and the clinical symptoms remit dramatically postpartum after the delivery of the placenta (Page, 1939). In a case of preeclampsia with extrauterine pregnancy, removal of the fetus alone was not suYcient; symptoms persisted until the placenta was delivered (Shembrey and Noble, 1995). One report suggests that in cases of preeclampsia with discordant twins, selective feticide reverses preeclampsia, with the attenuation of symptoms occurring in a timeframe consistent with placental involution (Heyborne and Porreco, 2004). Several lines of evidence suggest that placental insuYciency is central to the pathogenesis of preeclampsia (Karumanchi et al., 2004): (1) Pathological
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examination of placentas from preeclamptic pregnancies reveals numerous placental infarcts and sclerotic narrowing of arterioles (De Wolf et al., 1975; Khong et al., 1986); (2) placental bed biopsies in preeclamptics have been noted to have inadequate trophoblastic invasion of maternal decidual arterioles leading to tight and constricted vessels (Gerretsen et al., 1981; Robertson et al., 1967); (3) maternal risk factors for preeclampsia include medical conditions that predispose a patient to underlying vascular insuYciency such as chronic hypertension, diabetes, systemic lupus erythematosus, as well as acquired and inherited thrombophilias (Dekker, 1999); (4) obstetrical conditions such as multiple gestations or hydatiform moles that increase placental mass but with a relative decrease of placental blood flow increase the risk of preeclampsia (Dekker, 1999); and (5) animal models of preeclampsia involve creating placental insuYciency by disrupting uterine blood flow (Casper and Seufert, 1995; Kumar, 1962). During normal placental development, cytotrophoblasts invade the maternal spiral arterioles and completely remodel the maternal spiral arterioles into large capacitance vessels with low resistance (Gerretsen et al., 1981; Robertson et al., 1967). This endovascular cytotrophoblast invasion involves replacement of not only the endothelium but also the highly muscular tunica media. In preeclampsia, there is shallow placental cytotrophoblast invasion of uterine spiral arterioles, leading to reduced placental perfusion and consequently placental insuYciency (Brosens et al., 1972). Paradoxically, even in the normal placenta, there is little or no invasion of the uterine venules. The primary event that contributes to the failed trophoblast invasion/diVerentiation in preeclampsia is unknown, but genetic, immunological, and environmental factors (such as hypoxia and nutritional deficiencies) are thought to play a role. Extensive studies (by Fisher et al., 1981) suggest that diVerences in O2 tension may be the governing factor that regulates the invasive behavior of the cytotrophoblasts (Genbacev et al., 1996, 1997). The remodeling of the spiral arterioles is thought to begin in late first trimester and is complete by 18–20 weeks. Although the exact gestational age at which the trophoblast invasion of these arterioles ceases is unclear, histological studies show that fewer invasive trophoblasts are seen in the decidua with increasing gestational age. DiVerentiation of trophoblasts along the invasive pathway involves alteration in expression of a number of diVerent classes of molecules, including cytokines, adhesion molecules and extracellular matrix molecules, metalloproteinases, and class Ib major histocompatibility complex molecule, HLA‐G (Damsky et al., 1992, 1994; Fisher and Damsky, 1993). During normal diVerentiation, invasive trophoblasts alter their adhesion molecule expression from those that are characteristic of epithelial cells (integrin 6/ 4, v/ 5, and E‐cadherin) to those of endothelial cells (integrin 1/ 1, v/ 3, PECAM, and VE‐cadherin), a process referred to as pseudo‐vasculogenesis (Zhou
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et al., 1993, 1997b). Both in vitro and in vivo studies show that trophoblasts obtained from patients with preeclampsia fail to undergo these alterations of adhesion molecules and pseudo‐vasculogenesis (Lim et al., 1997; Zhou et al., 1997a), a finding that is disputed by some groups (Kaufmann et al., 2003). The molecular pathways that regulate pseudo‐vasculogenesis may involve a vast array of transcription factors, growth factors, and cytokines (Zhou et al., 2003). Considerable attention has been given to angiogenesis‐related gene products such as VEGF, angiopoietin/tie, and ephrin family proteins and their role in regulating pseudo‐vasculogenesis and invasiveness. Interestingly, invasive trophoblasts have been found to express VEGF, PlGF, VEGF‐ C, and their receptors. Furthermore, blocking their signaling pathways decreases the expression of integrin 1 (a marker of pseudo‐vasculogenesis) in vitro (Zhou et al., 2002). However, in vivo evidence directly linking abnormalities of VEGF signaling to impaired pseudo‐vasculogenesis is lacking at the present time. More recently, the invasive trophoblasts were also found to express L‐selectin, an adhesion molecule that mediates leukocyte migration from blood to tissues (Genbacev et al., 2003). It has been hypothesized that abnormalities of the selectin system at the fetal‐ maternal interface may account for implantation failures and preeclampsia. Finally, trophoblast expression of HLA‐G, a nonclassical class I molecule that has shown to be decreased in preeclampsia, has been hypothesized to protect trophoblasts from NK cell attack at the implantation site (MoVett‐King, 2002). Long‐standing and severe preeclampsia is associated with placental changes such as atherosis, fibrinoid necrosis, thrombosis, and placental infarction (De Wolf et al., 1975). Although not all these lesions are uniformly found in patients with preeclampsia, there appears to be a correlation between the severity of the disease and the extent of the lesions. Furthermore, in about one third of preeclamptic women (especially in those with term preeclampsia), these placental changes are not present. Abnormal remodeling of the spiral arterioles results in placental ischemia, which in turn is thought to lead to the secretion of soluble factors into the maternal bloodstream. However, evidence establishing a causal relationship between abnormal placentation and the maternal syndrome is lacking.
III. Systemic Endothelial Dysfunction (Stage 2) Generalized endothelial dysfunction can account for most of the clinical aspects of preeclampsia (Roberts, 1998): hypertension through disturbed endothelial control of vascular tone, proteinuria from increased glomerular vascular permeability, coagulopathy as a result of abnormal endothelial expression of procoagulants, and liver dysfunction from hepatic ischemia.
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Data from several studies support the theory that the maternal response in preeclampsia is secondary to generalized endothelial dysfunction. Studies have reported increased circulating fibronectin, factor VIII antigen, and thrombomodulin, all markers of endothelial cell injury in patients with preeclampsia (Friedman et al., 1995; Hsu et al., 1993; Taylor et al., 1991). Flow‐mediated vasodilation has also been found to be impaired in preeclamptic vessels, further suggesting abnormal endothelial function (Cockell and Poston, 1997; McCarthy et al., 1993). Decreased production of endothelial‐derived vasodilators such as prostacyclins, increased production of endothelins and enhanced vascular reactivity to angiotensin II also suggest abnormal endothelial function (Clark et al., 1992; Gant et al., 1973; Mills et al., 1999). Renal biopsies from patients with preeclampsia reveal diVuse glomerular endothelial swelling referred to as glomerular endotheliosis (Fisher et al., 1981). Finally, serum from preeclamptic women causes endothelial activation in human umbilical vein endothelial cells in vitro (Roberts et al., 1992). The identification of circulating factors mediating endothelial dysfunction has been the source of great research interest for decades. Several groups have reported alterations of cytokines/growth factors/chemicals such as TNF‐ , IL‐6, IL‐1 , IL‐1 , Fas ligand, neurokinin‐B, oxidized lipid products, and ADMA (asymmetric dimethyl arginine) that are released by the placenta and/or other maternal sources in preeclampsia (Benyo et al., 2001; Conrad et al., 1998; Page et al., 2000; Roberts and Cooper, 2001; Savvidou et al., 2003). However, there is no evidence that any of these molecules are etiological. Increased sensitivity to angiotensin II, a consistent feature of preeclampsia, has been found to be secondary to increased bradykinin (B2) receptor upregulation in preeclamptic patients (AbdAlla et al., 2001). This, in turn, was found to lead to heterodimerization of B2 receptors with angiotensin II type I receptors (AT1), and this AT1/B2 heterodimer was shown to increase responsiveness to angiotensin II in vitro. It is unclear, however, whether these alterations are pathophysiological or epiphenomena. Along similar lines, increased circulating concentrations of agonistic antibodies to the angiotensin‐1 (AT‐1) receptor have been reported in women with preeclampsia (Wallukat et al., 1999). Stimulation of the AT‐1 receptor by these autoantibodies might contribute to the vascular damage and the enhanced angiotensin II sensitivity noted in preeclampsia (Dechend et al., 2003; Xia et al., 2003). These antibodies have also been encountered in other examples of vascular injury such as vascular rejection (Dragun et al., 2005), suggesting that they may be secondary to the generalized microangiopathy of preeclampsia. Studies from several laboratories have demonstrated an increased placental expression and secretion of sFlt1 (soluble fms–like tyrosine kinase 1; see following), a naturally occurring circulating vascular endothelial growth
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factor (VEGF) antagonist in patients with preeclampsia (Koga et al., 2003; Maynard et al., 2003; Zhou et al., 2002). Importantly, when administered exogenously to rats, sFlt1 alone has been shown to be suYcient to induce a preeclampsia‐like phenotype (Maynard et al., 2003). A. Excess Circulating sFlt1, Impaired VEGF Signaling, and Antiangiogenic State VEGF is an endothelial‐specific mitogen that plays a key role in promoting angiogenesis. VEGF’s activities are mediated primarily by its interaction with two high‐aYnity receptor tyrosine kinases: kinase‐insert domain region (KDR) and fms‐like tyrosine kinase‐1 (Flt‐1) that are selectively expressed on vascular endothelial cell surface (Dvorak, 2002). Alternative splicing of Flt‐1 results in the production of an endogenously secreted protein referred to as sFlt1, which lacks the cytoplasmic and transmembrane domain but retains the ligand‐binding domain (He et al., 1999b; Kendall and Thomas, 1993). Thus, sFlt1 can antagonize circulating VEGF by binding to it and preventing VEGF’s interaction with its endogenous receptors located in the vasculature (Fig. 1). sFlt1 also binds and antagonizes placental growth factor (PlGF), another member of the VEGF family that is made in the placenta predominantly (Fig. 1). In vitro studies confirm that excess placental sFlt1 production induces an antiangiogenic state in the serum of preeclamptic women that can be rescued by exogenous VEGF and PlGF (Maynard et al., 2003). sFlt1 alone, when administered to pregnant rats, induced albuminuria, hypertension, and renal pathological changes of glomerular endotheliosis by antagonizing circulating VEGF and PlGF and inducing endothelial dysfunction. In addition, circulating levels of free VEGF and free PlGF were found to be decreased in conjunction with elevated sFlt1 in the bloodstream at the time of disease presentation (Chaiworapongsa et al., 2004; Koga et al., 2003; Maynard et al., 2003; Tsatsaris et al., 2003). Although sFlt1 is made in small amounts by other tissues (endothelial cells and monocytes), the placenta seems to be the major source of circulating sFlt1 during pregnancy as evidenced by a dramatic fall in circulating concentrations of sFlt1 after delivery of the placenta (Maynard et al., 2003). The increase in sFlt1 precedes the onset of clinical disease by at least 5 weeks (Hertig et al., 2004; Levine et al., 2004) and appears to be more pronounced in severe and early‐onset preeclampsia (Levine et al., 2004). When free PlGF and free VEGF were measured throughout pregnancy, it was found to be decreased in preeclamptics well before the onset of clinical disease (Levine et al., 2004; Polliotti et al., 2003; Taylor et al., 2003). Finally, decreased urinary PlGF has also been reported to precede clinical preeclampsia (Levine et al., 2005).
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Figure 1 Mechanism of action of sFlt‐1. sFlt‐1 protein, derived from alternative splicing of Flt‐1, lacks the transmembrane and cytoplasmic domains but still has the intact VEGF- and PlGF-binding extracellular domain. During normal pregnancy, VEGF and PlGF signal through the VEGF receptors (Flt‐1) and maintain endothelial health. In preeclampsia, excess sFlt‐1 binds to circulating VEGF and PlGF, thus impairing normal signaling of both VEGF and PlGF through their cell‐surface receptors. Thus, excess sFlt‐1 leads to maternal endothelial dysfunction. Reproduced with permission from Bdolah et al., 2004.
VEGF is known to stimulate angiogenesis and promote vasodilation by stimulating NO and prostacyclin formation (signaling molecules that are decreased in preeclampsia) (He et al., 1999a). Furthermore, a significant percentage of cancer patients receiving VEGF‐signaling antagonists develop hypertension and proteinuria (Kabbinavar et al., 2003; Yang et al., 2003). Even a 50% reduction of renal VEGF production in genetically modified mice resulted in glomerular endotheliosis and proteinuria (Eremina et al., 2003). These data suggest that excess sFlt1, by neutralizing VEGF and PlGF, may play a causal role in the pathogenesis of the maternal syndrome in preeclampsia. Data suggests that VEGF may be particularly important in maintaining the health of fenestrated endothelium (Risau, 1998), which is found in the renal glomerulus, choroid plexus, and the hepatic sinusoids— organs disproportionately aVected in preeclampsia. It has been shown that VEGF induces endothelial fenestrae in vitro (Esser et al., 1998) and even a 50% decrease in VEGF production in the glomerulus in mice leads to not only glomerular endotheliosis but also loss of glomerular endothelial fenestrae (Eremina et al., 2003).
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B. sFlt1 and Maternal–Fetal Conflict Substantial evidence suggests that sFlt1 is, at least partially, responsible for the endothelial dysfunction of preeclampsia. Why then does the placenta release a factor that damages the maternal endothelium? A conventional interpretation would be that sFlt1 performs some other function and that endothelial damage is an occasional maladaptive side eVect of its release into the maternal circulation, perhaps in women who are particularly ‘‘susceptible.’’ Another possibility should also be considered. The placenta may release sFlt1 into the maternal circulation to cause endothelial dysfunction because the associated vasoconstriction benefits the fetus by directing a greater share of maternal cardiac output to the intervillous space of the placenta. In this view, stage II of preeclampsia is an adaptive response of the conceptus to the placental insuYciency arising from stage I of preeclampsia (Haig, 1993, 1996). During pregnancy, the maternal systemic circulation can be conceptualized as consisting of two subcirculations, placental and nonplacental, arranged in parallel: the placental subcirculation consists of all the vessels that supply maternal blood to the intervillous space of the placenta; the nonplacental subcirculation consists of all vessels that supply other tissues of the maternal body (Fig. 2). Increases in the nonplacental resistance (Rn), as occurs in preeclampsia, would result in increased blood flow through the placental subcirculation, other things being equal (i.e., for unchanged cardiac output and placental resistance, Rp). More generally, any increase in the ratio of nonplacental to placental resistance (Rn/Rp) will result in a larger fractional share of maternal cardiac output flowing through the intervillous space (Haig, 1999).
Figure 2 A simple model of the maternal circulation during pregnancy. Maternal systemic cardiac output is shared between placental and nonplacental subcirculations. In stage I of preeclampsia, inadequate modification of maternal spiral arterioles results in increased placental resistance (Rp). In stage II of preeclampsia, placental release of sFlt1 causes a disproportionate increase in the nonplacental resistance (Rn) relative to the placental resistance (Rp). As a result, an increased share of maternal cardiac output is directed to the placental subcirculation. Modified from Haig, 1999.
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The hypothesis that preeclampsia is an adaptation of malnourished fetuses to increase their supply of nutrients posits that the endothelial dysfunction of preeclampsia disproportionately increases nonplacental resistance relative to placental resistance. This seems plausible—given the remodeling of spiral arterioles that occurs during the first half of pregnancy—but is yet to be experimentally demonstrated. The hypothesis is based on the evolutionary theory of parent–oVspring conflict (Trivers, 1974) that what is ‘‘best’’ for a parent is not always ‘‘best’’ for an oVspring, and vice versa. This conflict is illustrated in the clinical dilemmas of treating preterm preeclampsia: the longer that induction of delivery is delayed, the greater the risk to a mother’s health but the greater the benefits to the fetus. If the induction of preeclampsia is an adaptation to enhance fetal nutrition, then the adaptation need not be simple and could involve the release of multiple placental factors that target diVerent physiological systems of the mother. That is, sFlt1 may be just one component, albeit an important component, of a cocktail of substances that are released into maternal blood in nutritionally compromised pregnancies. C. Speculations About the Mechanisms of Preeclampsia If sFlt‐1 is an important cause of preeclampsia, there might be at least two kinds of predisposing factors. One might involve the overproduction of sFlt‐1. Conditions falling in this category might include multiple gestation, hydatiform mole, trisomy 13, and, possibly, first pregnancy. Another set of predisposing factors would include disorders that sensitize the maternal vascular endothelium to the antiangiogenic eVects of sFlt‐1. Such factors might include obesity, preexisting hypertension or renal disease, diabetes, and preexisting vasculitis. It is interesting that the alteration in the angiogenic factors in the serum of obese patients with preeclampsia was somewhat lower than that in lower‐weight preeclamptic patients (Levine et al., 2004; Thadhani et al., 2004a,b). It is not yet known whether diabetes, hypertension, and preexisting renal disease predispose to preeclampsia by increasing the production of sFlt‐1 or by sensitizing the vascular endothelium to its presence. D. Unanswered Questions There are limitations and several unanswered questions to the sFlt1 story. The precise mechanisms of excess sFlt1 production by the placenta are not known, and importantly, the role of sFlt1 in normal placental development and in placental pseudo‐vasculogenesis is not clear. No coagulation or liver
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function abnormalities or brain abnormalities (eclampsia) were reported in sFlt1‐treated animals. Moreover, genetic studies provide little support for a role for sFlt1. For example, both an Australian/New Zealand cohort (Moses et al., 2000) and an Icelandic cohort (Arngrimsson et al., 1999) have suggested a maternal susceptibility locus on chromosome 2, bearing no known relationship to sFlt1. One possible interpretation is that these studies detect a locus responsible for susceptibility to stage I of preeclampsia, whereas placental production of sFlt1 is responsible for stage II of the disease. Although, it is also possible that such loci are associated with transcription factors or splicing factors aVecting sFlt1 production, it seems more likely that there are other yet unidentified genetic factors that contribute to this multifactorial disease. On the other hand, the hypothesis that excessive production of sFlt1 may play a causal role in preeclampsia is supported by studies of the occurrence of this syndrome in mothers of infants with trisomy 13. The genes for sFlt1 and Flt‐1 are carried on chromosome 13. Fetuses with an extra copy of this chromosome should theoretically produce more of these gene products than their normal counterparts. The incidence of preeclampsia in mothers who carry fetuses with trisomy 13 is in fact greatly increased, as compared with all other trisomies or with control pregnant patients (Tuohy and James, 1992). It has been noted the women carrying trisomy 13 fetuses have a greater concentration of circulating sFlt1 as compared to normal karyotype controls, thus providing a molecular explanation for the increased risk of preeclampsia observed in these patients (Bdolah and Karumanchi, unpublished observations). Serum concentrations of sFlt1 have been found to be modestly elevated in patients with IUGR without preeclampsia (Tsatsaris et al., 2003), a finding that has not been confirmed by others (Shibita et al., 2004). Finally, although sFlt1 was elevated in most patients with preeclampsia, it was not elevated in some patients with mild preeclampsia (Levine et al., 2004). Thus, it is likely that additional synergistic factors that are elaborated by the placenta may yet be identified that play a role in the pathogenesis of the generalized endothelial dysfunction and vascular damage noted in preeclampsia.
IV. Conclusions In summary, preeclampsia is believed to be a two‐stage disease with an initial placental syndrome that is followed by the maternal syndrome (Fig. 3). The maternal syndrome in preeclampsia is a state of generalized endothelial dysfunction secondary to excessive amounts of circulating antiangiogenic factors (such as sFlt1) that are released by the diseased placenta (Fig. 3). The excess sFlt1 theory in the pathogenesis of preeclampsia fits very well with
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Summary of the pathogenesis of preeclampsia.
the maternal–fetal conflict that has been previously proposed as the basis of the development of preeclampsia. Understanding the mechanisms of placental dysfunction in preeclampsia should further clarify the etiology of preeclampsia. Future studies specifically characterizing the various circulating proteins elaborated by the preeclamptic placenta and understanding their relationship with already‐identified mediators of endothelial dysfunction such as AT1‐AA trophoblast debris and sFlt1 should help in clarifying the pathogenesis of the maternal syndrome. Although improvements in obstetrical and perinatal care have dramatically reduced morbidity and mortality from preeclampsia (especially in the developed world), there have been no significant breakthroughs in the treatment of preeclampsia over the last 40 years. The promising early results of agents such as aspirin and calcium supplementation have not been borne out in large randomized, controlled trials (Caritis et al., 1998; Levine et al., 1997). Therapeutic strategies aimed at rescuing the endothelial dysfunction with agents such as VEGF, PlGF, and prostacylins should be tested in patients with severe disease and hence might allow the delivery to be safely postponed. As understanding continues to advance based on molecular and genetic techniques, hopefully new interventions may improve the management of this important syndrome in the near future.
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Acknowledgments We would like to thank Vikas P. Sukhatme and Franklin Epstein for helpful suggestions and support. This work was supported by NIH grants (DK 065997 and HL079594) to S.A.K.
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Index A A . See -amyloid peptide ABA. See Abscisic acid Abscisic acid (ABA), 89–90, 99 environmental stresses and, 90 PCD regulation by, 232–233 treatment with, 231 ACE. See Angiotensin converting enzyme ACT. See Activator of CREM in testes Activator of CREM in testes (ACT), 154–156 Active oxygen species (AOS), 226 PCD and, 230, 233 TEs and, 238 Acute hydrocephalus, 26 AD. See Alzheimer’s disease Adenosine, CP and, 18 ADHs. See Alcohol dehydrogenases Adult muscle stem cells, 117–121 Aerenchyma, 252 Caþþ signaling in, 240 development of, 240 ethylene signaling in, 239–240 formation trigger for, 239 PCD in, 239–241, 249 biochemical data for, 241 DNA laddering in, 240–241 PM invagination and, 240 Age, 85–86 CNS and, 27 ethylene and, 88 modeling of, 199–223 photosynthesis decline and, 85 Albumin, 6, 30 bovine serum, 151–152 Alcohol dehydrogenases (ADHs), 13, 34 Aldehyde dehydrogenase 1 family (ALDH), A2 member of (ALDH1A2), 14 ALDH. See Aldehyde dehydrogenase 1 family Aleurone cells, 228–229, 252 PCD and, 229–231, 248 ALS. See Amyotrophic lateral sclerosis
Alzheimer’s disease (AD), 2, 200, 204–205, 208–211, 215 characteristics of, 28 CP and, 28–30 CSF secretion rates and, 26 CSF TTR reduction in, 29 CSF turnover restoration in, 31 familial, 11 genes of, 209 hallmarks of, 208 Amino acid, 17 Amyloid Beta Protein Precursor (APP), 208 intracellular domain (AICD), 209 -amyloid peptide (A ), 29 Amyloid plaques, 208–209, 211 Amyotrophic lateral sclerosis (ALS), 205, 212–215 Caenorhabditis elegans model of, 213 Drosophila melanogaster model of, 212–215 glial cells and, 214 oxidative damage from, 212 Androgen, 134–137, 146, 151 action mechanisms of, 137–142, 150 liver and, 135 modes of action of, 133 nonclassical responses of, 139–141, 151 prostate cells and, 140 Sertoli cells and, 140 spermatogenesis and, 140 primary response genes of, 137–138 prostate and, 150 secondary response genes of, 138–139, 150 definition of, 150 steroid hormones and, 131 TF induction by, 133 Androgen receptor (AR), 134, 136, 140, 142–146, 167 dimerization of, 133 domains of, 142–143 exon removal, 145 gene location of, 142 modular structure of, 142
313
314 Androgen receptor (AR) (cont.) spermatogenesis and, 143–146 testosterone production and, 146 Androgen response elements (AREs), 133, 138, 157 lack of, 139, 150 Androgen-induced transcription factor, 133 Angioblasts, 63, 70–71 definition of, 55 formation of, 57 grl/hey2 and, 64 induction of, 57 restriction of, 59 migration and diVerentiation of, 55–57 origin and specification of, 55–59 specification of flk1 and, 58 signals for, 57 Angiogenesis, 54 preeclampsia and, 300 VEGF stimulation of, 303 Angiotensin converting enzyme (ACE), 6 Anther, 242 Anthocyanins, 84 Antiangiogenic state, 302–303 Antipodal cells, 244 Aortic bifurcation, 64 AOS. See Active oxygen species Apical meristem, 241 Apoptosis, 227, 235 activation of, 227 PCD and, 247, 248–250 AR. See Androgen receptor Arabidopsis, 87–88 BRs and, 238 cysteine proteases found in, 94 cytokinin levels in, 90 ethylene constitutive overproduction by, 88 gfa2 mutant of, 245 GS and, 96–98 hxk1 knockout mutant, 86–87 IPT and, 90 JA and, 244 leaf senescence in, 86 lipid degradation in, 96 MAPK and, 100 nutrient recycling in, 96 ore4 mutant, 85 PCD and, 228 ethylene and, 246 roots of, 241–242
Index SAG gene expression in, 98–99 senescence-specific gene SAG12, 86 transcription factor genes in, 101–102 Arachnoid membrane, 25 Arcuate nucleus (ARC), 16 ARE. See Androgen response elements Arginase gene, 138–139 Arginine vasopressin (AVP), 27, 33 CP perfusion inhibition by, 27 CSF secretion inhibition by, 27 Aromatase, 147 Arterial endothelial cells development of, 74 diVerentiation of, 53, 62, 65 VEGF promotion of, 66–67 fate of, 68 plasticity of, 68–69 Arterial endothelial diVerentiation, 62, 65 Arterial endothelium, 65 Arterial-venous diVerentiation Notch signaling role in, 61 regulation of, 60 Arteries, 60 Asparagine (AS), 97–98 Aspartate aminotransferase (ASPA), 97 Autophagy, 227, 235 definition of, 227–228 pathway of, 93–95 PCD and, 247 Avian embryos, 57 AVP. See Arginine vasopressin Axial vessels, 54 Axon-guidance families, 75 B Basement membrane. See Extracellular matrix Basic FGF, 12 Basic helix-loop-helix (bHLH), 54 grl/hey2 and, 63 spermatogenesis and, 167–168 Blood-brain barrier (BBB), 7, 30, 264, 266, 274 cytokines and, 272 disruption of, 22 gelatinase regulation of, 279 LEP transport across, 19 Blood-CSF barrier (BCSFB), 2, 4, 7, 9, 16, 20, 30 function of, 18
315
Index glucose transport across, 17 LEP transport across, 14, 19 PRL transport across, 20 Blood-testis barrier (BTB), 265 biology of, 263–296 cadmium disruption to, 278, 283 cadmium model of, 268–269 composition of, 285 cytokines and, 272–274, 282 damage to, 278 dynamic modeling for, 267–270 dynamics of, 271 regulation of, 270–285 ECM and, 282 integrity loss for, 279 2-MG and, 281 model potentials for, 269–270 molecular architecture of, 267 protease/protease inhibitors and, 282 regulation of, 263–296 model of, 282–285 spermatogenesis and, 282 restructuring of, 270–271 spermatogenesis and, 263–296 BMPs. See Bone morphogenic proteins Bone marrow, 115 Bone marrow derived cells (BMDC), 117–120, 125 Bone morphogenic proteins (BMPs), 11 Bovine serum albumin (BSA), 151–152 Brain development, 8–16 estrogen and, 149 fluid compartments of, 3–6 protection of, 7 TTR entrance into, 9 volume of, 2 Brain injury ischemic, 2 traumatic, 21 types of, 21 Brain interstitial fluid (ISF) bulk flow of, 3–5 CSF and, 3–5 direction of flow of, 4 sources of, 3–5 volume transmission and, 5 Brain parenchyma, 4, 6 penetration of, 5
Brassinosteroids (BRs), 89, 99 PCD and, 247 TEs and, 238 Brd transcription factors, 168–169 Breast cancer, 140–141 BRs. See Brassinosteroids BSA. See Bovine serum albumin C Caenorhabditis elegans, 200 ALS model of, 213 APP homologues in, 210 mutant polyQ proteins and, 206 PCD and, 227 PD model and, 208 CAG repeats, 203–206 Calcium signaling (Caþþ), 100, 231, 251 androgen and, 140 cAMP-response element (CRE), 137 Pdha2 silencing and, 172 cAMP-response element binding protein (CREB), 137, 153–156 actions of, 153–154 functional depletion of, 156 phosphorylation of, 141 Sertoli cell target genes of, 156 spermatogenesis and, 156 Cancer/testis antigens, 174, 176 spermatogenic role of, 169–170 Cardinal vein assembly of, 69–71 embryonic, 68 formation of, 56, 71 posterior, 53 Cardiotoxin, 126–127 Cardiovascular death, 298 Carotenoids, 84 Carotid artery, 74 Carrier proteins, 34 Caspases, 227 homologues of, 228 Catabolism, 84, 92 Cathepsins, 275–278 types of, 275 CD45, 118–119 expression of, 116 CD45þ, 117, 120–124 CD45þ/Sca1þ cells, 121–123, 127 muscle stem cell fate and, 124–125
316 Cell death. See also Programmed cell death; Senescence execution process, 91–98 chlorophyll degradation, 92 lipid degradation, 95–96 nutrient recycling, 96–98 protein degradation, 93–95 Cellular membrane, 92 degradation of, 95 Cellular retinol-binding proteins (CRBPs), 14 Central nervous system (CNS), 1, 3, 11, 14 adult neuronal cells in, 24 age related changes and, 27 chemical communication in, 5 development of, 2, 6–16 CP-CSF system and, 31 disorders of, 23 early development of, 10 failure of, 2 homeostasis of, 3 injury to, CP-CSF System role in, 21–23 LEP and, 15–16 malformation of, 7–8 normal development of, 10 normal growth of, 8 repair process of, 35 stem cells in, 23–24 Cereal crops, 85 Cerebellum LEP synthesis by, 16 normal development of, 14 Cerebral cortex, 8 LEP synthesis by, 16 Cerebral ventricles, 2, 4 formation of, 6 Cerebrospinal fluid (CSF), 1–2, 4, 11 AD and, 26, 29, 31 adenosine and, 18 AVP and, 27 brain protection and nourishment by, 7–8 brain volume percentage of, 2 cytokines interleukin-1 , 28 drainage of, 30 hydrocephalus and, 25 FGF2 elevation in, 23 formation rate of, 7 glucose secretion by, 17 ISF and, 3–5 limiting factor in, 7 net flux of, 6 neuronal organization and, 7
Index primary function of, 2 PRL and, 20 production of, 3 protein composition of, 5–6 protein origin of, 5–6 rising levels of, 27 secretion of, 25–28, 26 measurement of, 26 SVZ B cells and, 24 TGF and, 22–23 TTR secretion into, 9 turnover of, 30–31 FGF2 and, 30 volume increase of, 25 volume transmission and, 5 Chemorepellents. See CP-derived chemorepellents Chiari malformation, 23 Chlorophyll, 84, 92 Chloroplasts breakdown of, 91 protein degradation in, 93 vacuoles and, 93–95 Choroid plexus (CP), 1–2, 4, 6, 8, 31–35 AD and, 28–30 adenosine and, 18 aging and, 26 appearance of, 7 AVP and, 27 blood flow to, 7 composition of, 4 diVerentiation of, 6–7 embryonic, 8, 11–12 formation of, 6 growth factor secretion by, 31 growth factor synthesis by, 32, 33 growth factors identified in, 21 IGF2 expression in, 9–10 mitochondrial dysfunction and, 28 morphogenesis of, 10 morphological changes in, 28 neuronal survival promotion by, 31 PRLR concentration in, 20 progenitor cells and, 31 protein and peptide synthesis in, 28–30 role of, 3 senescence, protein and peptide synthesis and, 28–30 transport systems in, 16–21 Choroid plexus-cerebrospinal fluid system. See CP-CSF system
317
Index Choroidal epithelium, 3, 4, 7–8, 10, 14 factors produced by, 21 glucose net flux across, 17 growth of, IGF2 and, 29 IGF2 production by, 22 IGF2 secretion by, 11 position of, 1 synthesis by, 1–2 TGF 1 production by, 22 transport protein expression, 18 transporter and, 16–17, 17 polarized distribution of, 17 Choroidal LEP receptor, 15 Choroidal transplant, 35 Chromatin assembly factor-1 (CAF1), 163–164 Chromatin condensation, 243 Cib. See Naþ-nucleoside cotransport system Cloche mutation, 58 CNS. See Central nervous system CP. See Choroid plexus CP-CSF axis, 29 active role of, 2–3 function decline in, 2 senescence related dysfunction of, 31 CP-CSF system, 1–52 adulthood and, 16–24 development of, 6–8 post-injury role of, 21–23 role in maintaining CNS homeostasis of, 3 senescence of, 24–31 CP-derived chemorepellents, 12–13, 31, 33 axon guidance role of, 12–13 CP-derived growth factor, 2, 21 CpG, 171, 180–181 DNA methylation and, 171–172 methylation of, 179 mutation of, 178 CRBPs. See Cellular retinol-binding proteins CRE. See cAMP-response element CRE modulator protein (CREM), 142, 153–156 actions of, 153–154 activator of, 154–156 downstream targets of, 154 isoforms of, 154 male sterility and, 154 CREB. See cAMP-response element binding protein CSF. See Cerebrospinal fluid CSF-borne
bioactive substances, 8 lipophilic compounds, 18 oligopeptides clearance of, 20–21 processing/degradation of, 20 CSF/plasma concentration ratio, 6 Cystatin C, 275–278, 276 Cysteine aspartate specific proteases (Caspases), 227–228 Cytokines, 28, 33 BTB dynamics and, 272–274, 282 endothelial dysfunction and, 301 interleukin-1 , 28 Cytokinins, 90, 237 D Defense-related (DR) genes, 87–88 Delta-Notch mediated lateral inhibition, 63 Delta-Notch signaling, 60–63, 74–75 Dementia, 25 estrogen and, 29 nicotine and, 29 Dermomyotome, 114 DLAV. See Dorsal longitudinal anastomotic vessels Dmrt1. See Doublesex and Mab-3-related transcription factor-1 DNA laddering, 226–227, 231, 233, 235, 243, 248–250, 251 aerenchyma and, 240–241 leaf sculpting and, 236 leaf senescence and, 237 petal senescence and, 247 TE and, 238–239 DNA methylation, 135, 170–181 CpGs and, 171–172 direct regulation by, 177 factor binding inhibition by, 179 hypomethylation, 173–177 indirect regulation by, 177 Pgk2 and, 175–176 tissue-specific expression and, 171 transcription inhibition by, 176–177 Dnmt1, 171 Dopaminergic neurons, 207 Dorsal aorta assembly of, 69–71 bifurcation of, 64 development of, 69–70 embryonic, 68
318 Dorsal aorta (cont.) formation of, 56, 71 patterning of, 53 Dorsal longitudinal anastomotic vessels (DLAV), 71–72 Doublesex and Mab-3-related transcription factor-1 (Dmrt1), 162–163 Drosophila melanogaster, 62 AD model in, 209–211 age-related disease modeling in, 199–223 ALS model in, 212–215 anatomy and development of, 201–202 drug screening with, 202 fALS model in, 212–213 generation time of, 200–201 life span of, 201 modeling with, 216–217 mutations of, 201 neurodegenerative disease study in, 200–203, 215–216 neurofibrillary tangles and, 211 PD model in, 207–208 PS homologue in, 210 therapeutic drug testing on, 216 Duct of Cuvier, 54 E ECF. See Extracellular fluid ECM. See Extracellular matrix Ectoplasmic specialization (ES), 263–264, 270 Embryo cardinal vein of, 68 CP and, 8, 11–12 development, 228–235 elimination of, 234, 252 PCD in, 234–235, 248 vasculature of development of, 53 formation of, 53–81 vessel, 74 Embryogenesis, 6 Embryonic myogenesis, 114, 124 Endoderm, 57 Endoreduplication, 233 Endosperm, 228 cell death in, 229–234 PCD in, 232–233, 248 pattern of, 232
Index Ricinus communis, 233–234, 248, 252 degeneration in, 233–234 lipid mobilization in, 233 Endothelial cells arterial, 59–69 tip cells, 74 types of, 72 venous, 59–69 Endothelial dysfunction, 297 circulating factors and, 301 cytokines and, 301 preeclampsia and, 305 systemic, 300–306 TNF- and, 301 vascular, 298 Endothelial markers, 58 Endothelial progenitor cells, 53 Endothelial tubes, 53 Environmental stress, 87 senescence signals and, 100 Enzyme inhibitors, 34 Enzymes, 34 Ependyma, 4 EphB4, 59, 60, 62 ephrinB2, 59, 60, 68 expression of, 67 VEGF and, 66 Epididymal cells, 138, 143 Equilibrative nucleoside transporter 1 (rENT1), 18 ER. See Estrogen receptor ERK. See Extracellular-regulated kinase ES. See Ectoplasmic specialization Estradiol, 147, 149, 152 Estrogen, 131, 134 cell stimulation by nonclassical, 148–149 primary, 148–149 secondary, 148–149 cell type response to, 148 dementia and, 29 male fertility and, 152 spermatogenesis and, 136, 152 steroid hormones and, 131 testis function and, 147–149, 153 TTR and, 29 Estrogen receptor (ER), 136, 148 Ethylene aerenchyma and, 239–240 age and, 88 biosynthetic genes, 232
319
Index leaf senescence, 237 PCD and, 232, 246–247 production of, 88 receptor genes, 232 Extracellular fluid (ECF), 3, 5 Extracellular matrix (ECM), 264, 266, 269, 274, 280 BTB regulation and, 282 components of, 275 migration across, 283 Sertoli cells and, 270–271 Extracellular proteolysis, 274–275 Extracellular-regulated kinase (ERK), 140–141
Fluorescence-activated cell sorting (FACS), 115–116, 122 fms-like tyrosine kinase-1 (Flt-1), 302–303 sFlt1, 304, 307 action of, 303 excess, 302–303 maternal-fetal conflict, 304–305 mechanisms of, 305–306 production of, 306 trisomy 13, 306 Follicle-stimulating hormone (FSH), 143, 152 receptor of, 167 TNF- and, 273 G
F FACS. See Fluorescence-activated cell sorting fALS. See Familial Amyotrophic lateral sclerosis Familial Alzheimer’s disease (FAD), 209 PS and, 211 Familial Amyotrophic lateral sclerosis (fALS) Drosophila melanogaster model of, 212–213 mutations in, 212 Female gametophyte, 244–245 Ferredoxin-dependent GOGAT (FD-GOGAT), 96 Fetal Sertoli cells, 135 FGF2. See Basic FGF FGFR1. See Fibroblast growth factors, FGF2 Fibroblast growth factors (FGFs) autocrine regulation by, 12 FGF2, 23, 32 age related changes to, 27 CSF secretion reduction by, 27 CSF turnover and, 30 neuronal survival and, 27 receptor for (FGFR1), 23 retrograde neuronal transport of, 23 juxtacrine/paracrine regulation by, 12 flk1, 57–58 defects in, 73 inactivation of, 57 mutations of, 58 Flowering PCD and, 242–247 self-incompatibility and, 242 Flt-1. See fms-like tyrosine kinase-1
GAL4/UAS system CAG repeats and, 203–206 function of, 201 PD and, 207 Galactolipids, 96 GAs. See Gibberellic acids GATA factors, 168 GDF-15. See Growth/diVerentiation factor 15 GDH. See Glutamate dehydrogenase Gelatinases, 279–280 Gene expression, 103 testis-specific, 170–181 tissue-specific, 175–177, 181 transcription regulation, 101–102 GFAP. See Glial fibrillary acidic protein GH. See Growth hormone Gibberellic acids (GAs), 230 PCD and, 229, 247 PSV and, 229 Glial fibrillary acidic protein (GFAP), 24 Glucose, 17 GLUT1. See Naþ-independent glucose transporter Glutamate dehydrogenase (GDH), 97 Glutamate synthase (GOGAT), 96–98 Glutamine synthetase (GS), 96–98 G-protein-mediated signaling, 100 Grey matter, 25 Gridlock (grl) mutation, 54 grl/hey2, 54, 65, 68 bHLH and, 63 expression of, 64 phenotypic characteristics of, 63–64 role of, 63–66
320 Growth hormone (GH), 19–20 IGF and, 29 Growth/diVerentiation factor 15 (GDF-15), 12, 32 GS. See Glutamine synthetase Gymnosperms, 234 H HD. See Huntington’s disease Heat-shock factors (Hsfs), 136, 158–159 Hemangioblast, 57 Hematopoietic cells, 116 Hematopoietic markers, 58 Hematopoietic stem cell (HSC), 57, 118–119, 121–123 Hexokinase (HXK), 86 Hippocampus, 8 Homeobox genes, 164–167. See also Reproductive homeobox genes Hox subfamily, 164 spermatogenesis and, 164–165 Hormone responses nonclassical, 136–137, 139–141, 151 prostate cells and, 140 Sertoli cells and, 140 spermatogenesis and, 140 primary, 136–138 secondary, 136, 138–139 HR. See Hypersensitive response HSC. See Hematopoietic stem cell Hsfs. See Heat-shock factors Human congenital heart disease, 73 embryonic stem-cell research, 117 life expectancy of, consequences of, 200 neurodegenerative disease in, 199 venous arterialization and, 68–69 Huntington’s disease (HD), 203–207, 216 key features of, 203 sequestration of acetylases, 206 HXK. See Hexokinase Hydrocephalus, 13, 23. See also Normal pressure hydrocephalus aging and, 25–28 CSF drainage and, 25 ventriculomegaly and, 25 Hypersensitive response (HR), 226 Hypertension, 297, 300, 307
Index Hypomethylation, 171, 173–174 tissue-specific, 175–177 Hypothalamus, 16 I ICL. See Isocitrate lyase ICP. See Intracranial pressure IGF1. See Insulin-like growth factor 1 IGF1R. See Insulin-like growth factor 1 receptor IGF2. See Insulin-like growth factor 2 IGF2R. See Insulin-like growth factor 2 receptor IGFBPs. See Insulin-like growth factor binding proteins Insulin-like growth factor 1 (IGF1), 10 Insulin-like growth factor 1 receptor (IGF1R), 10 Insulin-like growth factor 2 (IGF2), 9–11, 21–22 biallelic expression of, 10 distal eVects of, 11 expression of, 9–10 increase in, 21–22 mRNA presence of, 10 neuroprotection by, 29 Insulin-like growth factor 2 receptor (IGF2R), 10 Insulin-like growth factor binding proteins (IGFBPs), 11 IGFBP2, 29 Interleukin-1 (IL-1 ), 33 Intersegmental vessels, 72 excessive branching of, 74 formation of, 71 growth inhibition of, 73 patterning of, 53–54 Interstitial fluid. See Brain interstitial fluid Intracranial pressure (ICP) aging and, 26 elevation of, 25 formula for, 25–26 NPH and, 26 Intrathecal fraction (IF), 6 IPT. See Isopentenyl transferase Ischemic brain injury, 12, 21 Ischemic insult, 10 Ischemic stroke, 35 ISF. See Brain interstitial fluid
321
Index Isocitrate lyase (ICL), 97, 98 Isopentenyl transferase (IPT), 90 J jagged1 mutant, 61 Jasmonic acid (JA), 88–89, 99 Arabidopsis and, 244 exogenous application of, 89 K Kidney, 139 androgen and, 138 estrogen and, 149 L Lactate dehydrogenase C (LDH-C), 172 Leaf development, 236–242 Leaf sculpting, 236, 252 PCD and, 248 Leaf senescence, 83–112, 237, 252. See also Senescence Caþþ signaling and, 237 chlorophyll degradation and, 92 cysteine proteases associated with, 94 cytokinin production upregulation in, 237 diVerential gene expression in, 98–99 DNA laddering and, 237 ethylene signaling reduction in, 237 factors of, 83 gene expression during, 98–99 transcription regulation of, 101–102 induction of, 83 nuclear control of, 98 PCD and, 249 protein degradation and, 93–95 regulating signals of, 85–91 ABA, 89–90 age, 85–86 BRs, 89 cytokinins, 90 ethylene, 88 JA, 88–89 NO, 91 PAs, 91 RG, 86 SA, 89 stresses, 87–88 sugars, 86–87
signal perception for, 100 signal transduction for, 100 suppression of, cytokinins and, 90 transcription factors, 101–102 transcriptome, 87 visible signs of, 84, 92 LEP. See Leptin LEP receptor (LEPR), identification of, 15 Leptin (LEP), 14 BCSFB-mediated transport of, 14 CNS developmental role of, 15–16 energy regulation by, 15, 19 functional, CNS abnormalities and, 15 neuronal projection developmental role of, 16 transport of, 19 Leptomeninges, 9 IGF2 production by, 22 Lewy bodies, 207–208 Lewy neurites, 207 Leydig cells, 135, 139, 145, 150, 152 ablation of, 144 androgen and, 135 estrogen and, 148 fetal development of, 146 gene expression in, 172 steroidogenesis in, 146 LH. See Luteinizing hormone L-histidine, 21 Lipids catabolites of, 98 degradation of, 95–96 enzymes for, 95 Liver, 8 androgen and, 138 estrogen and, 149 Luteinizing hormone (LH), 143, 146 Lymphatics, 4 Lysosomes, 227 M Macrophage-inhibiting cytokine 1 (MIC-1), 12, 32 Malate synthase (MS), 97, 98 Male contraception, 285–286 Male fertility, estrogen and, 147, 152 Male germ cells, 80, 134, 135, 173–174, 178, 265, 284 Dmrt1 expression in, 162 migration of, 263–264
322 Male germ cells (cont.) specific gene expression in, 172–174 spermatogenesis and, 271 Mammalian spermatogenesis. See Spermatogenesis MAP. See Mitogen-activated protein MAPK. See Mitogen-activated protein kinase Marrow-derived myogenic progenitors, 118 Maternal circulation, 304 MDR. See Multidrug resistance MDR1 P-glycoprotein (Prp), 18–19 Meninges, 6 Mesangioblasts, 117 Mesoderm, 57, 72 Methyl jasmonate (MeJA), 88 2-MG, 280–282, 284 basal level of, 281 Sertoli cells and, 283 TGF- 3 and, 281 TNF- and, 281 MIC-1. See Macrophage-inhibiting cytokine 1 Microangiopathy, 298 Micropyle, 244 Mind bomb mutant (mib), 61 Mitochondria, 91–92, 226, 243 Mitogen-activated protein (MAP), 140–141, 151 Mitogen-activated protein kinase (MAPK), 100, 283 2-MG and, 281 TGF- 3 and, 273–274, 278 Monocarpic plants, 86 Moyamoya syndrome, 23 MP. See Stem cells, main population MS. See Malate synthase Multidrug resistance (MDR), 18 Multidrug resistance-associated protein 1 (MRP1), 18, 19 Multipotent cell, 113 Muscle contractile fiber of, 114 degeneration of, 115 developmental origin of, 113–114 exercise-induced damage, 126 extreme damage to, 125–127 regeneration of, 113, 122, 124 CD45þ and, 120 functionality of, 126 regulatory factors of, 115
Index repair of, 120 postnatal, 114–116 stem cells of, 122, 123, 127 definition of, 114 Myelomonocytic cells, 119 Myf5, 114–115, 122 MyoD, 115, 122–123 Myofibers, 114, 118–119 Myogenesis, 116, 118 embryonic, 114 regenerative, 113–130 Myogenic markers, 124 Myogenic progenitors, 113, 118, 121, 125 Myogenic regulatory factors, 124 Myotome, 114 N NADP-dependent glutamate synthase (NADP-GOGAT), 97 Naþ-independent glucose transporter (GLUT1), 17 Naþ-nucleoside cotransport system (cib), 18. See also rCNT3 NCCs. See Nonfluorescent chlorophyll catabolites Neurodegenerative diseases Drosophila melanogaster models of, 200–203, 215–216 future research on, 216–217 human, 199 therapies for, 35 Neurofibrillary tangles (NFTs), 208–209 Drosophila melanogaster and, 211 Neurogenesis adult mammalian brain and, 23–24 normal, 7 Neuromuscular junction (NMJ), 201 Neuronal diVerentiation, 13 Neuron-specific enolase (NSE), 6 Neuropeptides, 33 Neuropil, 9, 22 Neuropilin-1 (NRP-1), 68 Neuropilin-2 (NRP2), 13 Neurotrauma, 35 NFTs. See Neurofibrillary tangles Nitric oxide (NO), 90–91, 99 antioxidant properties of, 230–231 signaling by, 230–231 VEGF and, 303 NMJ. See Neuromuscular junction
Index Nonfluorescent chlorophyll catabolites (NCCs), 92 Nongenomic responses. See Hormone responses, nonclassical Nonsteroid hormone nuclear receptors, 156–158 Normal pressure hydrocephalus (NPH), 2 CSF secretion rates and, 26 ICP and, 26 TNF- elevation in, 28 Notch signal pathway, 61–63 cell fate regulation by, 62 grl/hey2 and, 64 notch1 mutant, 61–62 notch1/notch4 mutant, 61 NRP2. See Neuropilin-2 Nuclear hormone receptors (NRs), 136, 151 Nucleoside, 17–18 Nutrient deficiency, 87 Nutrient recycling, 96–98 O Obesity (ob), 19 gene for, 14–15 Ontogenesis, 7 Organic ion transporter (OAT), 19 Orphan nuclear receptors, 156–158 Out of bounds mutation (obd), 72–73 Ovary, 149 Oxidative stress, 87 P PaO. See Pheide a oxygenase Parkinson’s disease (PD), 204, 207–208 CSF secretion rates and, 26 etiology of, 207 PAs. See Polyamines Pathogen infection, 87 Pathogenesis-related (PR) genes, 87 Pax3, 114, 121 Pax7, 121–123, 122, 123 PCD. See Programmed cell death PCI, 278–279 PD. See Parkinson’s disease Pdha2, 172, 174, 176–177 Pem/Rhox5, 137–139, 150, 159, 175, 181 cancer/testis antigens and, 169–170 fertility and, 165–166 GATA factors and, 168
323 gene targets of, 166 Sertoli cell expression of, 166 PEPT2, 20–21 Peptide/histidine transporter (PTH1), 21 Peptides role of, 8–16 transport of, 19–21 Peroxisome, 91 Petal senescence, 252 DNA laddering and, 247 future research directions for, 247–253 PCD and, 246–247, 250 Ricinus endosperm and, 247 Pheide a oxygenase (PaO), 92 Phosphatases, 100 Phosphoglycerate kinase-1 (Pgk1), 172 Phosphoglycerate kinase-2 (Pgk2), 172–177 Photosynthetic activity, 84, 92 Pial-glial lining, 4 Pituitary anterior, 19 AR-mediated events in, 137 Placenta, 298 abnormal, 298–300, 307 antiangiogenic factor release by, 301, 306 angiogenic factors released by, 297 insuYciency, 298–299 Placental ischemia, 298–300 Placental lactogen (PL), 19 Plant organ development, 225–261 Plant reproduction, 242–247 Plasma, 5 Plasma membrane (PM), 240 Pluripotent stem cells, 115 Plzf. See Promyelocytic leukemia zinc factor Pollen grains, 242 Polyamines (PAs), 90, 91, 99 Polyglutamine disorders, 203–207 PolyQ diseases, 203, 204 key features of, 203 mutant proteins and, 206 Postmitotic senescence, 83. See also Leaf senescence Postnatal muscle regeneration, 115 muscle repair, 114–116 PPDK. See Pyruvate orthophosphate dikinase Prealbumin. See Transthyretin Preeclampsia angiogenesis and, 300
324 Preeclampsia (cont.) characterization of, 297–298 complications of, 298 cytotrophoblasts and, 299 endothelial dysfunction of, 305 etiology of, 307 maternal syndrome, 306 maternal-fetal conflict, 304–307 mechanisms of, 305 pathogenesis of, 307 angiogenic factors in, 297–312 placental syndrome, 306 risk factors for, 298 severe, 300 stage 1, 298–300 stage 2, 300–306, 304 cytokines and, 301 stages of, 297 trisomy 13 and, 306 trophoblast diVerentiation, 299–300 VEGF and, 300–302 Presenilin (PS), 209 expression of, 210 FAD mutations and, 211 PS1, 209 PS2, 209 PRL. See Prolactin PRL/GH/PL family, 20 Progenitors, 113, 121, 125 Programmed cell death (PCD), 225–261 ABA regulation in, 232–233 aerenchyma and, 239–241, 249 biochemical data for, 241 DNA laddering in, 240–241 aleurone cells and, 229–231, 248 animal cell, 227 AOS and, 230, 233 apoptosis and, 247, 248–250 Arabidopsis and, 228 ethylene and, 246 autophagy and, 247 BRs and, 247 Caenorhabditis elegans and, 227 caspase-like activity and, 235 cell fate and, 251–252 definition of, 226 delay of, 232 DNA degradation and, 225–226 embryo in, 234–235, 248 endoreduplication and, 233 endosperm in, 232–233, 248
Index pattern of, 232 ethylene and, 232, 246–247 female gametophyte and, 244–245 flower male sexual organs and, 242–244 flowering and, 242–247 GAs and, 229, 247 leaf sculpting and, 236, 248 leaf senescence and, 249 petal senescence and, 246–247, 250 root cap and, 241–242, 249 root development and, 236–242 seed and embryo development and, 228–235 sexual organ abortion and, 242, 249 SI and, 245–246, 251 tapetum and, 243–244, 249 TEs and, 249, 251 DNA laddering in, 238–239 proteolysis and, 239 Prolactin (PRL) cognate receptor (PRLR), 20 transport of, 19–20 Promyelocytic leukemia zinc factor (Plzf), 161–162 Prostate androgen and, 138, 150 nonclassical responses and, 140 Protease inhibitors, 274–282, 276–277 BTB and, 282 2-MG and, 280 Proteases, 274–282, 276–277, 284 BTB and, 282 Protein, degradation of, 93–95 N release from, 96 ubiquitin pathway and, 95 Protein storage vacuoles (PSV), 229 Proteinuria, 297, 300, 307 Prp. See MDR1 P-glycoprotein PS1. See Presenilin, PS1 PS2. See Presenilin, PS2 PTH1. See Peptide/histidine transporter Pyruvate orthophosphate dikinase (PPDK), 97, 98 R RA. See Retinoic acid Radioiodinated human serum albumin (RIHSA), 30 RALDHs. See Retinaldehyde dehydrogenases RCC. See Red chlorophyll catabolite
Index rCNT3, 18 Reactive oxygen species (ROS) accumulation of, 85–86 increased levels of, 88 Receptor kinases, 100 Red chlorophyll catabolite (RCC), 92 Regeneration, muscle, 113 functionality of, 125 rENT1. See Equilibrative nucleoside transporter 1 Reproductive growth (RG), 86, 99 Reproductive homeobox genes (Rhox), 166–167 Retinaldehyde dehydrogenases (RALDHs), 13, 34 RALDH2, 14 Retinoic acid (RA), 2, 13–14 cerebellum and, 14 patterning of neuronal diVerentiation, 13 TTR and, 29 Reversible infertility, 285–286 Rhox8, 142 RIHSA. See Radioiodinated human serum albumin Root cap, 252 PCD in, 241–242, 249 Root development, PCD and, 236–242 ROS. See Reactive oxygen species Rubisco, 85 degradation of, 93 S SA. See Salicylic acid SAGs. See Senescence-associated genes SAH. See Subarachnoid hemorrhage Salicylic acid (SA), 89, 99 endogenous, 89 sALS. See Sporadic Amyotrophic lateral sclerosis SAS. See Subarachnoid space Satellite cells, 4, 114, 119–120, 125 definition of, 114–115 localization of, 115 ontogeny of, 122–123 origin of, 117 Pax7 expression in, 121 specification of, 121–122 Sca1, 116, 118–119 Sca1þ, 117, 121–123, 121 Schizophrenia, 25
325 Schwentine mutation, 58, 72–73 scl, 58–59 SDGs. See Senescence downregulated genes Secondary messengers, 137 Seed development, 228–235 Self-incompatibility (SI), 245–246, 252 evolution of, 245 PCD and, 245–246, 251 Semaphorin family (SEMA), 33, 73 Seminiferous epithelium, 263, 276–277, 278 cytokines and, 272 damage to, 280 extracellular proteolysis and, 275 2-MG and, 280–281 TJs and, 270–271 tubule of, 264 Senescence. See also Leaf senescence cellular level of, 91 cysteine proteases associated with, 94 mitotic, definition of, 84 PCD and, 225, 235 postmitotic, definition of, 84 protein degradation and, 93 receptor kinases and, 100 signaling of, mechanisms for, 100 syndrome, 84 transcription factors for, 101–102 Senescence downregulated genes (SDGs), 99 Senescence-associated genes (SAGs), 83, 99 analysis of, 84 Arabidopsis and, 98–99 senescence-specific gene SAG12, 86 cloning of, 98 identification of, 84 induction of, 87 regulation of, 102 upregulation of, 98 Sertoli cells, 134–135, 138–139, 145, 150, 159, 181, 265, 276–277, 279, 284 androgen action in, 151 androgen response and, 141 AR expression in, 143–144 AR function in, 143 AR loss in, 152 Cadmium targeting of, 268–269 cathepsins and, 275 CpG promoter expression in, 174 CREB phosphorylation and, 141 CREB target genes in, 156 Dmrt1 expression in, 162 E-box and, 167, 177
326 Sertoli cells (cont.) ECM and, 270–271 gelatinase secretion by, 279 gene expression in, 172 loss of AR expression, germ cells aVected by, 144–145 2-MG production in, 283 2-MG secretion by, 280 nonclassical responses and, 140 Pem/Rhox5 expression in, 166 Plzf-null, 162 proliferation of, 159–160 Dmrt1 and, 162–163 Rhox expression during, 166–167 Rhox8 expression in, 142–143 seminiferous tubule and, 264 Sox3 and, 159–160 steroidogenesis regulation by, 146 TGF- 3 and, 273 TNF- receptors in, 272 tPA and, 278 uPA and, 278 vitamin A deficiency and, 270 SI. See Self-incompatibility sICAM1. See Soluble intercellular cell adhesion molecule 1 Single gene mutations, 199 Skeletal muscle, 119 adult, 115 adult stem cells in, 116–125 AR-mediated events in, 137 regeneration of, 116, 118 Slit protein family of chemorepellents (SLIT), 33 SLIT2, 12–13 SLIT3, 13 SNAP-25. See Synaptosome-associated protein of 25 kDa SOD1. See Superoxide dismutase 1 Soluble intercellular cell adhesion molecule 1 (sICAM1), 6 Somatic cell, 180 testis genes in, 174–175, 178–179 Somatopleural mesoderm, 57 Somite, 113–114 Somitic paraxial mesoderm, 57 Sox proteins, 159–161 Sox3, 159–160 Sox9, 160–161 Sry, 159 transcription regulation by, 159
Index SP. See Stem cells, side population Sperm1, 165 Spermatids, 143, 276, 277 Brd3 expression in, 169 CAF1-deficiency and, 163 diVerentiation of, 145 estrogen signaling and, 147 Sertoli cells and, 264 Spermatogenesis, 135, 139, 150, 157 AR role in, 143–146 blocking of, 143–144 BTB and, 263–296 BTB regulation during, 282–285 CAF1 and, 163–164 estrogen and, 136, 152 extracellular proteolysis and, 275 gene regulation in, 131–197 germ cell movement during, 271 homeobox genes and, 164–165 nonclassical responses and, 140 Rhox genes and, 166 steps of, 145–146 suppression of, 269 testosterone regulation of, 136 transcription factor roles in, 170 Spermatogonia, 134, 276 Brd4 expression in, 169 Plzf expression in, 161–162 stem cells, 132 Spermatozoa, 132, 143 Spina bifida, 7–8 Spinal cord, 24 Splanchnopleural mesoderm, 57 Sporadic Amyotrophic lateral sclerosis (sALS), 212 Sry, 159, 174, 176 Stem cells adult, 117, 120 undiVerentiated, 124 adult skeletal muscle and, 116–125 main population (MP), 115, 116 muscle, 113–130, 122, 123, 127 Pax7 regulation of, 121–123 pluripotent, 23–24, 115 adult skeletal muscle, 115 bone marrow, 115 possible sources of, 23–24 side population (SP), 115, 118, 121, 122 diVerentiation of, 116 marrow (maSP), 115 muscle (mSP), 115, 122
Index spermatogonial, 132 Stem development, 236–242 Sterility, 163 Steroid hormones, 131–132, 151 androgen and, 131 estrogen, 131 routes used by, 133 synthesis of, 135 Stresses, 87–88 STX1. See Syntaxin 1 Subarachnoid hemorrhage (SAH), 21 Subarachnoid space (SAS), 3–4 volume increase in, 25 Subventricular zone (SVZ) aging and, 31 diVerentiation in, 24 neurogenesis in, 23, 30 pluripotent stem cells in, 24 types of cells in, 24 Sugars, 86–87 leaf senescence and, 86 senescence signals and, 100 signaling molecule activity of, 86 Superoxide dismutase 1 (SOD1), 212 Suspensor, 229, 252 limiting growth of, 235 PCD in, 234–235, 248 Synaptobrevin, 15 Synaptosome-associated protein of 25 kDa (SNAP-25), 15 Syncytium, 114 Synergids, 244, 252 PCD and, 250 Syntaxin 1 (STX1), 15 T T. See Testosterone T3. See Triiodothyronine T4. See Thyroxine Tapetum, 252 mutants of, 243 PCD and, 243–244, 249 TBC. See Tubulobulbar complex Telencephalon, 11 TEs. See Tracheary elements Testicular receptor 2 (TR2), 158 Testicular receptor 4 (TR4), 157–158 AR and, 157 network patterns of, 157–158 role of, 157
327 Testis, 143 androgen and, 138, 139–141 CpG and, 172 diVerentiation of, Dmrt1 and, 162–163 ER action in, 153 estrogen and, 147–149, 153 estrogen responsive genes in, 148–149 extracellular proteolysis and, 275 GATA factor expression in, 168 gene expression in, 170–181 gene identification in, 140 2-MG and, 281 primary functions of, 135 promoters of, 172 somatic cell gene expression in, 174–175, 178–179 somatic cells of, Rhox gene expression in, 166–167 sox family protein expression in, 159 spermatogenesis and, 135 steroid hormone synthesis and, 135 TGF- 3 and, 273 TR4 and, 157 transcription in, hormonal regulation of, 135–153 zinc-finger proteins and, 161 Testis-specific histone (TH2B), 173 Testosterone (T), 133, 136, 138 BSA and, 151–152 decrease in, 146 production of, AR role in, 146 spermatogenesis regulation by, 136 TF. See Transcription factor TGF- . See Transforming growth factor- Thylakoid membrane, 96 Thyroid gland, 8 Thyroid hormone, 157 Thyroxine (T4), 8, 9 TTR and, 29 tie2, 68 Tight junction (TJ), 263, 264 cadmium and, 268–269 Time-course studies, 175–176 TIMPs, 279–280 Tissue-type PA (tPA), 278–279 TJ. See Tight junction TNF- . See Tumor necrosis factor- Tox. See Rhox8 tPA. See Tissue-type PA TR2. See Testicular receptor 2 TR4. See Testicular receptor 4
328 Tracheary elements (TEs), 252 AOS and, 238 diVerentiation of, 237–239 vacuolar collapse in, 238 formation of, 238 PCD in, 249, 251 DNA laddering in, 238–239 proteolysis and, 239 Transcription factor (TF), 179 androgen-induced, 133 bHLH, 54 Brd, 168–169 GATA factors and, 168 germ cell-specific (CREM), 142 orphan NRs, 156–158 positive-acting, 170–171 repression of, 171 spermatogenic role of, 134, 167–170 testicular function of, 153–170 zinc-finger family, 161 Transcriptome, 87, 92, 93 Transforming growth factor- (TGF- ), 32 Transforming growth factor- (TGF- ), 2, 11–12, 22–23, 33 cell function regulation by, 11 cytokines and, 272 elevation of, 28 member of, 12 2-MG and, 280 neuronal organization during development by, 11 neuronal survival promotion by, 22 production of, 22 TGF- 3, 263–264, 278, 284 cadmium-induced, 283 cytokines and, 273–274 MAPK and, 273–274, 278 2-MG and, 281 release of, 283 Transthyretin (TTR), 6, 8–9, 34 aging and, 28–29 behavioral eVects of, 9 brain entry by, 9 CSF secretion of, 9 estrogen and, 29 nicotine and, 29 RA and, 29 sources of, 8 T4 and, 29 Triiodothyronine (T3) conversion to, 9
Index levels of, 8 Trisomy 13, excess sFlt1 and, 306 Tubulobulbar complex (TBC), 263, 266 Tumor necrosis factor- (TNF- ), 33, 263–264, 272–273, 283, 284 cytokines and, 272 elevation of, 28 FSH and, 273 2-MG and, 281 receptors for, 272 Sertoli cell activation by, 283 Tyrosine kinase, 10, 118 U Ubiquitin pathway, 95, 206, 208, 214 uPA. See Urokinase-type PA Upstream activating sequence (UAS), 201. See also GAL4/UAS system Urokinase-type PA (uPA), 278–279 Uterus, 149 V Vascular branching, 74 Vascular development, 54, 60 Notch pathway and, 61 Vascular endothelial growth factor (VEGF), 13, 33, 57, 62, 303, 307 angiogenesis and, 303 arterial endothelial cells and, diVerentiation of, 66–67 cytokines and, 272 defects in, 73 definition of, 302 dorsal aorta development control by, 69–70 ephrinB2 and, 66 expression of, 58 gain-of-function and loss-of-function studies using, 67 impaired signaling of, 302–303 inactivation of, 58 intracellular signaling of, 72 misexpression of, 67–68 NO and, 303 preeclampsia and, 300–302 role of, 66–68 Vascular endothelial growth factor receptor-2 (VEGFR-2/flk1), 55, 57 Vascular tubulogenesis, 71
329
Index Vasculogenesis, 54, 58, 63 Vasoconstriction, 304 Vasopressin, 27 Vegetable storage proteins (VSPs), 98 VEGF. See Vascular endothelial growth factor Veins, 60 Venous arterialization, 68–69 Venous endothelial cells, 53 fate of, 68 plasticity of, 68–69 Venous sinuses, 4 Vertebrate embryo, 53 Virchow-Robin spaces, 5 Volume transmission, 5 VSPs. See Vegetable storage proteins W Water stress, 87 WRKY transcriptor factor gene family, 101–103 X X chromosome, 142, 169 Xenobiotics, 18–19
Xenopus, 57, 69–70 VEGF expression in, 70 Xylem, 237
Y y10 mutation, 72–73 Yeast invertase, 86–87
Z Zebrafish blood vessel formation in, 75 embryonic vascular development of, 54 embryonic vasculature, anatomy of, 55 embryos, model organism, 72 genetics of, 53–81, 69 grl hypomorphic mutation, 64 mutant types, 61 angiogenic growth, 72 use as model organism, 54 vascular tubulogenesis in, 71 VEGF expression in, 70
