Advances in Developmental Biochemistry, Volume 3

ADVANCES IN DEVELOPMENTAL. BIOCHEMISTRY

Vofume3

1994

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ADVANCES IN DEVELOPMENTAL B OCH EMISTRY

Editor: PAUL M. WASSI RMAN Department of Cell and Developmenta1 Biology Roche Institute of Molecular Biology Nutfey,0New jersey

VOLUME 3

1994

@ JAl PRESS INC. Greenwich, Connecticut

London, England

Copyright 0 1994 byJAl PRESS INC. 55 Old Post Road No. 2 Greenwich, Connecticut 06836 ]A/ PRESS LTD. The Courtyard 28 High Street Hampton Hill, Middlesex TWl2 1PD England All rights reserved. No part of this publication may be reproduced, stored on a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, filming, recording, or otherwise, without prior permission in writing from the publisher. ISBN: 1-55938-865-X Manufactured in the United States of America

CONTENTS

vii

LIST OF CONTRIBUTORS

PREFACE Paul M. Wassarman

ix

EXPRESSION AND FUNCTION OF PROTEIN KINASES DURING MAMMALIAN GAMETOGENESIS Deborah L. Chapman and Debra 1. Wolgemuth

1

REGULATION OF THE DOPA DECARBOXYLASE GENE DURlNG DROSOPHlLA DEVELOPMENT Martha J. Lundell and Jay Hirsh

55

TRANSCRIPTION FACTORS IN MAMMALIAN DEVELOPMENT MURINE HOMEOBOX GENES S. Steven Potter

87

EXPRESSION AND FUNCTION OF C-MOSIN MAMMALIAN GERM CELLS Geoffrey M . Cooper

127

REGULATION OF PIGMENTATION DURING MAMMALIAN DEVELOPMENT Friedrich Beermann, Ruth G a d , and Gunther Schutz

149

INDEX

179

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LIST OF CONTRIBUTORS

Friedrich Beermann

Swiss Institute for Experimental Cancer Research (ISREC) Epalinges, Switzerland

Deborah L. Chapman

Department of Genetics and Development The Center for Reproductive Sciences Columbia University College of Physicians and Surgeons New York, New York

Geoffrey M. Cooper

Dana-Farber Cancer Institute Boston, Massachusetts

Ruth GanB

Division of Molecular Biology of the Cell German Cancer Research Center Heidelberg, Germany

Jay Hirsh

Department of Biology University of Virginia Charlottesville, Virginia

Martha J. Lundell

Department of Biology University of Virginia Charlottesville, Virginia

S. Steven Potter

Department of Pediatrics Children’s Hospital Research Foundation and University of Cincinnati College of Medicine Cincinnati, Ohio

vi i

viii

LIST OF CONTRIBUTORS

Gunther Schutz

Division of Molecular Biology of the Cell German Cancer Research Center Heidelberg, Germany

Debra J. Wolgemuth

Departments of Genetics and Development and Obstetrics and Gynecology The Center for Reproductive Sciences Columbia University College of Physicians and Surgeons New York, New York

PREFACE Advances in Developmental Biochemistry was launched as a series by JAI Press in 1992 with the appearance of Volume 1. The series is inextricably linked to the companion series, Advances in Developmental Biology, that was launched at the same time. As stated in the Preface to Volume 1: “Together the two series will provide annual reviews of research topics in developmental biologyhiochemistry, written from the perspectives of leading investigators in these fields. It is intended that each review draw heavily from the author’s own research contributions and perspective. Thus, the presentationsare not necessarily encyclopedic in coverage, nor do they necessarily reflect all opposing views of the subject.” Volume 3 of the series follows these same guidelines. Volume 3 of Advances in Developmental Biochemistry consists of five chapters that review specific aspects of mammalian and fly development. In Chapter 1, D. Chapman and D. Wolgemuth discuss the role of protein kinases, especially tyrosine- and serine/threoninekinases, in regulating cell cycle events during mammalian gametogenesis. In Chapter 2, M. Lundell and J. Hirsh discuss the regulation of the DOPA decarboxylase gene during Dmsophila development. DOPA decarboxylase is a key enzyme in biogenic amine biosynthesis and its expression is subject to both transcriptional and post-transcriptionalregulation. In Chapter 3, S. ix

X

PREFACE

Potter discusses the role of homeobox genes as master switches determining the developmental destinies of groups of cells during murine development. In Chapter 4, G. Cooper discusses the expression and function of the c-mos proto-oncogene in mammalian germ cells where it plays a central role in regulating the meiotic cell cycle. In Chapter 5, F. Beermann, R. GanB, and G. Schutz discuss the regulation of pigmentation during mammalian development,with emphasis on the production of melanin in mouse melanocytes. Finally, I am grateful to the authors for their excellent contributions, as well as for their cooperation and patience during the preparation of this volume. Paul M. Wassarman Series Editor

EXPRESSION AND FUNCTION OF PROTEIN KINASES DURING MAMMALIAN GAMETOGENESlS

Deborah L. Chapman and Debra J. Wolgemuth

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Introduction.. . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . 11. Protein Kinase Activity and Regulation of Cell Cycle Events in the Yeast and Selected Vertebrate Model Systems . . . . . . . . . . . . . . A. Serinefbeonine Kinases: Cdc2 and Cdks . . . . . . . . . . . . . B. Regulators of CdcUCdk2 Protein Kinase Activity: Cyclins . . . . . C. Dual-Specificity Kinases . . . . . . . . . . . . . . . . . . . . . . . D. Phosphatases: Cdc25 . , . . . . . . . . . . . . . , . . . . , . . . . 111. Role of Protein Phosphorylation in GermCell Differentiation . . . . . A. Historical Perspective . . . . . . . . . . . . . . . . . . . . . . . . B. Use of Mouse as a Model System: Oogenesis and Spermatogenesis IV. Early Germ Cell Differentiation . . . . . . . . . . . . . . . . , . . . . A. Kitandsteel . . . . . . . . . . . . . . . . . . . . . . . . , . . . .

Advances in Developmental Biochemistry Volume 3, pages 1-53. Copyright Q 1994 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-865-X 1

... 2 ... 3 ... 4 ... 5 ... 8 . . . 10 . . . 12 . . . 12 . . . 12 . . . 13 . . . 14 . . . 14

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DEBORAH L. CHAPMAN and DEBRA J . WOLGEMUTH

V. Mitoticstages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Kitandsteel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B . CyclinB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C . Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D . MAPKinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E . Raf . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Meiotic Stages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . MPF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B . CyclinB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C . CSF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D . Mos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Cdk2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. M015 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . G . Cdc25 Class of Phosphatases . . . . . . . . . . . . . . . . . . . . . . . H . Novel Kinases: mak and Nekl . . . . . . . . . . . . . . . . . . . . . . VII . Post-Meiotic Events . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Murinec-abl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B . MurinecycBl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C . Murineraf . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Fertilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX . Summary and Future Directions . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

16 16 17 18 20 22 23 23 24 25 26 29 29 30 30 31 32 33 33 34 35 35 36

PREFACE The phosphorylation and dephosphorylation of proteins constitute major mechanisms for controlling the transduction of extracellular signals into intracellular cues. which in turn control cell proliferation. survival. and differentiation. Signaling events are thus translated into developmental decisions. The phosphorylation of proteins results in a cascade of reactions and hence. amplificationof the stimulus by the regulation of several diverse cellular processes. One signal can thus result in pleiotrophic effects. These phosphorylation events are involved not only in the transduction of extracellular signals into the cell. but also in the restructuring of the cell in preparation for cell division. including chromosome condensation. spindle formation. formation of microtubule organizing centers. etc. In this chapter. we review critical stages of mammalian germ cell differentiation during which protein kinases (and their corresponding phosphatases) may be functioning. This discussion is not intended to serve as a comprehensive survey of various classes of kinases. such as serine threonine kinases. cyclic nucleotide kinases. proto-oncogene ki-

Expression and Function of Protein Kinases

3

nases, receptor tyrosine kinases, etc., known to be expressed in germ cells. The readers are referred to reviews by Hunter (1987) and Hanks et al. (1988) for detailed discussions of protein kinases and a review by Winer and Wolgemuth (1993) for a more extensive discussion of protooncogenes with protein kinase activity expressed during male germ cell development. Rather, we will place a particular emphasis on those kinases involved with regulating the eukaryotic cell cycle, with the bias that gametogenesis represents a specialized and highly regulated series of cell cycle events. The main classes of protein kinases that will be considered in this review are the tyrosine kinases and the serine/threonine kinases. Although results from the mouse system will be emphasized, results from other organisms will also be included, especially in particular stages of gametogenesis in which the mammalian system is poorly understood.

1. INTRODUCTION It has long been established that growth factors and their respective receptor tyrosine kinases function in the mitogenic pathway. However, expression of the receptor tyrosine kinases has also been observed in differentiated tissues, indicating that they may also have a role in differentiation (see, for example, review on growth factor signaling, Chao, 1992).Receptor tyrosine kinases consist of three distinct domains: the extracellular ligand binding domain, a transmembraneregion and the intracellular cytoplasmic tyrosine kinase catalytic domain (reviewed in Pawson and Bernstein, 1990).Conformationalchanges brought about by binding of the ligand to the receptor ultimately results in the activation of the kinase, and hence, transmission of the signal from the extracellular to the intracellular domain. Binding of the ligand to the receptor can result in the dimerization of the receptor which, in turn, causes adjacent intracellular kinase domains to be brought together (reviewed in Ullrich and Schlessinger, 1990 and Schlessingerand Ullrich, 1992).This dimerization sets the stage for autophosphorylation of the kinase domain via intermolecular interactions between adjacent kinase domains. The tyrosineautophosphorylation domains within the receptor represent specific binding sites for cellular proteins with SH2 or SH3 domains (see below), which may be involved in proliferation and/or differentiation (reviewed in Schlessinger and Ullrich, 1992). Receptor tyrosine kinases not only autophosphorylate residues in the kinase domain; they also phosphory-

4

DEBORAH L. CHAPM AN and DEBRA 1. WOLGEMUTH

late exogenous substrates. In the cytoplasmic domain, serine/threonine and tyrosine residues serve as phosphorylation sites that regulate the activity of the receptor tyrosine kinase. Many non-receptor tyrosine kinases have been identified as products of retrovirally encoded oncogenes. Non-receptor tyrosine kinases can be divided into two groups: transmembrane and cytosolic families. The c-src tyrosine kinase is the prototype of the cytosolic tyrosine kinases. Regions within these non-receptor tyrosine kinases share homology with the Src kinase, known as Src homology 2 and 3 (SH2 and SH3) domains, and mediate protein-protein interactions between the receptor tyrosine kinases and the intracellular targets (reviewed in Cantley et al., 1991; Pawson and Gish, 1992; Mayer and Baltimore, 1993). Cytoplasmic serine/threonine protein kinases catalyze the transfer of phosphate groups to serine and threonine residues of target proteins. Serine/threoninekinases have been recognized as the products of protooncogenes (e.g., c-mos, c-raf) or as kinases intimately involved with the regulation of serinekhreonine kinase activity by CAMP.Some of these kinases specifically phosphorylate cellular structural proteins, such as histone, laminins, etc. Others phosphorylate still more kinases, resulting in either the activation or deactivation of downstream protein kinases. Specific examples in which serine/threonine kinases elicit specific cellular responses are discussed in this chapter.

II. PROTEIN KINASE ACTIVITY A N D REGULATION OF CELL CYCLE EVENTS IN THE YEAST A N D SELECTED VERTEBRATE MODEL SYSTEMS The eukaryotic cell cycle consists of distinct phases: DNA synthesis ( S ) and mitosis (M), which are separated by two gap phases (GI and G2). The identification of genes involved in the regulation of the eukaryotic cell cycle first came from studies of cell division control, cdc, mutants in both fission (Schizosaccharomyces pombe) and budding (Saccharomyces cerevisiae) yeasts (reviewed in Fosburg and Nurse, 1991). Progression through the cell cycle is regulated at two phases. Primary regulation of the cell cycle in S. cerevisiae is at START, which occurs during the GI phase. Passage through START signals commitment to the cell cycle. The chief point of cell cycle regulation in S. pombe is at the G2/M phase transition. Regulation at G2/M ensures that proper replication of the DNA has occurred prior to cell division (reviewed in Enoch and Nurse, 1991;

Expression and Function of Protein Kinases

5

Murray, 1992). This difference in regulation reflects the evolutionary divergence of the two species and underscores the diversity of control points even among lower eukaryotes. Genetic and molecular analysis of yeast cell cycle mutants has led to the identification of genes involved in regulating progression through the cell cycle. Many of these genes encode protein kinases. Phosphorylation of proteins is a particularly useful mechanism for controlling cell cycle progression because it is reversible. This means that following M phase, the cell can return to the GI phase of the next cell cycle. In the following paragraphs, we highlight the function of key cell cycle regulating genes in yeasts, S. pombe in particular,noting whether the functional homologues of these genes have been identified in higher eukaryotes, including mammals. A. Serinenhreonine Kinases: Cdc2 and Cdks A main regulator of the eukaryotic cell cycle is the 34 kDa serinelthreonine protein kinase encoded by the cdc2 gene in S. pombe (Simanis and Nurse, 1986). Homologues of cdc2 have been identified, by sequence homology and by functional complementation of cdc2 mutants, in all organisms thus far examined. Activation of the Cdc2 kinase results in the phosphorylation of various cellular substrates that ultimately drives the cell into M phase of mitotic and meiotic cell cycles (reviewed in Nigg, 1991). The Cdc2 protein kinase is required at two stages in the cell cycle of fission yeast, at START and at the GdM phase transition (reviewed in Nurse, 1990). Regulation of the Cdc;! Kinase

Although the level of Cdc2 remains relatively constant through the cell cycle, its kinase activity oscillates, peaking at the G2 to M phase transition of the cell cycle (Draetta and Beach, 1988; Moreno et al., 1989; Gautier et al., 1989). The kinase activity of Cdc2 is assayed by phosphorylation of an exogenous substrate, most commonly histone H1 (Arion et al., 1988). Activity of the Cdc2 kinase is dependent on its association with cyclin and its state of phosphorylation. Reviews by Nurse (1990), Clarke and Karsenti (1991), and Fleig and Gould (1991) provide extensive discussions on the regulation of Cdc2 kinase activity. During G2, Cdc2 is phosphorylated on tyr-15 in yeast (Gould and Nurse, 1989) and on both thr- 14 and tyr- 15 in animal cells (Krek and Nigg, 1991a; Norbury et al., 1991). Phosphorylation of Cdc2 at these sites is dependent on its

6

DEBORAH L. CHAPMAN and DEBRA I. WOLGEMUTH

association with a cyclin subunit (Cyc) (Meijer et al., 1991;Parker et al., 1991; Solomon et al., 1990; 1992). At the GdM phase transition, the tyr-15 residue in yeast (Moreno et al., 1989) and both thr-14 and tyr-15 residues in animal cells (Draetta and Beach, 1988; Morla et al., 1988, 1989) are dephosphorylated, concomitant with the activation of the kinase. In fission yeast, a mutation in cdc2 that changes tyr-15 into a nonphosphorylatable phe- 15, results in premature entry into mitosis, before S phase is complete (Gould and Nurse, 1989). Similarly, changing both thr-14 and tyr- 15 of Cdc2 into nonphosphorylatableresidues results in the deregulation of its kinase activity when the mutated protein is expressed in Xenopus oocytes or fibroblasts (Krek and Nigg, 1991b; Norbury et al., 1991;Pickham et al., 1992).Dephosphorylation of tyr-15 in yeast and both thr-14 and tyr-15 in animal cells is rate limiting in the activation of Cdc2. Coupling completion of DNA synthesis with the onset of mitosis is essential for the fidelity of transmission of genetic information. In yeast, failure to complete S phase prior to mitosis results in mitotic catastrophe, a lethal phenotype characterized by abnormal chromosome segregation. Cdc2 has been shown to be dephosphorylated on tyr-15 in cells treated with agents that uncouple S and M phases, such as caffeine and okadaic acid (Smythe and Newport, 1992). Tyrosine phosphorylation of Cdc2 thus couples S phase with the onset of mitosis. Activity of the Wee1 protein kinase, which negatively regulates Cdc2 (see Section II.C, Fig. 2), is responsible for mitotic delay due to DNA damage (Rowley et al., 1992). Inhibition of DNA synthesis negatively regulates the Cdc25 phosphatase, a positive regulator of Cdc2 (see Section II.D, Fig. 2) (Enoch and Nurse, 1990). Cdc2 is also phosphorylated on thr-167 in S. pombe and thr-161 in Xenopus (Gould and Nurse, 1989; Solomon et al., 1990; Krek and Nigg, 1991a). Phosphorylation at thr- 161/167 may be required for activation of the kinase by stabilization of the Cdc2/Cyc complex (Booher and Beach, 1986; Ducommun et al., 1991; Gautier et al., 1991; Gould et al., 1991; Solomon et al., 1990, 1992; Desai et al., 1992; Krek et al., 1992; Pickham et al., 1992). Changing thr-161/167 to nonphosphorylatable residues resulted in decreased cyclin binding to Cdc2 (Ducommunet al., 1990; Gould et al., 1991;Pickham et al., 1992).Inactivation of the Cdc2 kinase and exit from M phase requires dephosphorylation of Cdc2 at thr-1611167 residue (Gould et al., 1991; Lee et al., 1991; Lorca et al., 1991; Krek et al., 1992) and proteolysis of the cyclin regulatory subunit (Murray et al., 1989; Glotzer et al., 1991) (Fig. 1).

Expression and Function of Protein Kinases

7

Figure 7. The cell cycle as a Cdc2 cycle. Progression through the eukaryotic cell cycle is sensitive to the phosphorylation state of Cdc2. A block to DNA synthesis ( S ) prevents dephosphorylation, and hence activation, of Cdc2. Impaired spindle function will prevent deactivation of Cdc2 and thus blocks exit from M phase (Hoyt et al., 1991; Li and Murray, 1991; reviewed in Nurse, 1991). Exit from M phase requires destruction of the regulatory subunit, Cyc B. Dephosphorylation of Cdc2 at thr-161 may act to destabilize the Cdc2/Cyc B complex and thus allow the ubiquitination of Cyc B followed by its destruction.

Homologues of the cdc2 kinase, known as cyclin-dependent kinases (cdk), have been identified in a number of different species, including Drosophila (Lehner and O’Farrell, 1990a), goldfish (Hirai et al., 1992), Xenopus (Blow and Nurse, 1990; Paris et al., 1991; Gabrielli et al., 1992a), mouse (McConnell and Lee, 1989; Ben-David et al., 1991) and human (Elledge and Spottswood, 1991; Tsai et al., 1991; Meyerson et al., 1992). The Cdks may function at distinct stages of the cell cycle as compared to Cdc2 (Fang and Newport, 1991; Elledge et al., 1992; Rosenblatt et al., 1992; Yasuda et al., 1992; Stem et al., 1993). For

8

DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

example, immunodepletion of the cdk2 homologue of cdc2 from Xenopus eggs (Fang and Newport, 1991) or human fibroblasts (Pagan0 et al., 1993) blocks DNA replication (S) but has no effect on entry into mitosis (M), whereas immunodepletion of Cdc2 has no effect on S, but blocks entry into M. Amurine cell line, tsFT210, carrying a temperaturesensitive mutation in the cdc2 gene arrests at the G2 phase at the restrictive temperature (Th’ng et al., 1990). Since cells arrest at the G2 phase and not at the Gl/S phase, this implies that some kinase other than Cdc2 is functioningat the Gl/S phase transition, but that this kinase could not compensate for the absence of a functional cdc2. Furthermore, transfection of cdk2 into the mutant cell line at the restrictive temperature did not improve growth, while transfection of cdc2 did improve growth (Yasuda et al., 1992). These results indicate that cdc2 and cdk2 function at discrete phases of the cell cycle and that at least one more kinase in addition to cdc2 is necessary for progression through the GdS phase transition. Similar to Cdc2, the activity of the Cdk2 kinase in HeLa cells has been shown to be regulated by phosphorylation at thr-14, tyr-15, and thr-161 (Gu et al., 1992; Sebastian et al., 1993).

B. Regulators of Cdc2/Cdk2 Protein Kinase Activity: Cyclins A combination of genetic and biochemical analyses has indicated a direct interaction between Cdc2/Cdks and Cyc. Association of Cyc with Cdc2 or Cdk2 results in activation of the kinase. Cyclin was originally identified in sea urchin eggs as a protein whose level greatly increased upon fertilization and subsequently oscillated during the early cell divisions of the embryo (Evans et al., 1983). The level of Cyc increased during interphase of the cell cycle, reached a maximum just prior to M phase, and dropped at the anaphase/metaphase transition of each cell cycle. Since their original discovery in invertebrates, cyclins have been identified in organisms ranging from yeast to humans (reviewed in Pines, 1991). In vitro studies have demonstrated that the Cyc subunit is responsible for substrate specificity of the CdWCyc complex (Hinds et al., 1992; Thomas et al., 1992; Peeper et al., 1993). Cyc B associates with Cdc2 to form acomplex known as the maturation or M phase promoting factor (MPF) (Dunphy et al., 1988; Lohka et al., 1988; Gautier et al., 1988, 1990; Labbe et al., 1989a,b). MPF was originally identified by its role in meiosis, as an activity found in mature frog eggs that was capable of causing germinal vesicle breakdown (GVBD) when injected into immature oocytes (Masui and Markert,

Expression and Function of Protein Kinases

9

1971). MPF activity is evolutionarily conserved, as shown by the ability of cytoplasm from the eggs of various organisms, including mouse (Sorenson et al., 1985; Hashimoto and Kishimoto, 1988), to induce meiotic maturation of immature oocytes from other species. Cell fusion studies demonstrated that MPF activity is not restricted to meiotic maturation. When interphase nuclei are exposed to the cytoplasm of mitotic cells by cell fusion, they undergo premature chromosome condensation and nuclear envelope breakdown (Rao and Johnson, 1970; Matsui et al., 1972). Furthermore, cytoplasm from mitotically active embryonic cells (wasserman and Smith, 1978) and mammalian tissue culture cells at the G2 phase of the cell cycle (Sunkara et al., 1979) are also capable of inducing meiotic maturation of Xenopus oocytes. These results suggest that a common factor, MPF, is capable of driving cells into M phase of mitotic and meiotic cell cycles. Cyclins have been divided into three classes, GI, A, and B, based on their amino acid similarity and the timing of their appearance during the cell cycle (reviewed in Minshull et al., 1989b;Hunt, 1991; Pines, 1991). GI cyclins, originally described in the budding yeast S. cerevisiae, appear during GI and function at START, passage through which commits the cell to a round of cell division (reviewed in Nasmyth, 1990). Cyclins C, D, and E have been identified in humans and fall into the GI class of cyclins (Koff et al., 1991; Lew et al., 1991; Xiong et al., 1991). The Aand B-type cyclins were originally classified under the common heading of mitotic cyclins, since they can induce entry into M phase when complexed with Cdc2 (Swenson et al., 1986; Pines and Hunt, 1987; Draetta et al., 1989; Roy et al., 1991). However, Cyc A appears before and is destroyed earlier than Cyc B during the cell cycle (Minshull et al., 1989a;Whitfieldet al., 1990;Pines andHunter, 1991b).Cyc Aisbelieved to function during S phase, as evidenced by its ability to activate DNA synthesis in a GI extract, and by its nuclear localization from S phase onward (D’Urso et al., 1990; Pines and Hunter, 1991a). Moreover, antibodies against Cyc A and cyc A specific antisense plasmids inhibit entry into S phase when injected into GI phase cells (Girard et al., 1991; Pagano et al., 1992; Zindy et al., 1992). In general, the B-type cyclins appear during interphase and function at the Gdh4 phase transition to trigger entry of the cell into M phase. This may be an oversimplification, since several B-type cyclins from yeast appear to function during S phase (Bueno et al., 1991;Epstein and Cross, 1992; Fitch et al., 1992; Grandin and Reed, 1993). Destruction of Cyc B occurs via the ubiquitin-mediated

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DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

proteolysis pathway and has been shown to be required for exit of the cell from M phase (Murray et al., 1989; Glotzer et al., 1991). Both Cyc A and B can associate with either the Cdc2 or Cdk2 protein kinase in vitro, with the resulting complexes capable of phosphorylating histone H1 (Peeper et al., 1993, and references therein); however, Cyc B has not been shown to associate with Cdk2 in mammalian cells in vivo. Cdk2, in association with Cyc A, is activated at the G1/S transition and is localized in the nucleus through S phase, further supporting a role for Cyc A in S phase (Pines and Hunter, 1991a; Pagano et al., 1992; Rosenblatt et al., 1992). Assignment of function to the B-type cyclins has been complicated by the identification of multiple B-type cyclins in organisms as evolutionarily divergent as S. cerevisiae, Xenopus, mouse and human. Complementation analysis revealed that two S. pombe M phase cyclins, cdcl3 and cig.2, have distinct functions in mitosis (Bueno and Russell, 1993). At least nine B 1-related sequences have been genetically mapped in the mouse genome (Hanley-Hyde et al., 1992; Lock et al., 1992). The roles of these different B-type cyclins are not understood. Our results on cycBl and cycB2 expression during murine germ cell development suggest that they may have distinct functions in this process (see Sections V1.B and V1I.B).

C. Dual-Specificity Kinases S. Pombe strains that have mutations in the weel gene undergo cell division prematurely, at approximately half the normal size (Russell and Nurse, 1987a). In S. pombe, the weel gene encodes a dual-specificity kinase, having both serinehhreonine and tyrosine kinase activity (Featherstone and Russell, 1991; Parker et al., 1991, 1992). Wee1 negatively regulates the Cdc2 kinase by phosphorylating it on tyr- 15 (Featherstone and Russell, 1991; Parker et al., 1991, 1992). Although yeast Wee 1 has been designated a dual-specificity kinase, threonine phosphorylation of Cdc2 by Wee1 has not been detected. The human homologue of Wee1 is a tyrosine-specific protein kinase (Parker and Pinwica-Worms, 1992; McGowan and Russell, 1993). It can phosphorylate Cdc2 on tyr-15 in vitro, thereby inactivating the Cdc2/Cyc B complex. weel and mikl encode redundant protein kinases, which apparently cooperate to negatively regulate Cdc2 kinase activity (Lundgren et al., 1991).

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Expression and Function of Protein Kinases

Genetic studies indicated that Niml (“new inducer of mitosis”) negatively regulates the Weel protein kinase regulatory pathway in S. pombe (Russell and Nurse, 1987b; Feilotter et al., 1991). Overexpression of niml can drive cells into M phase at a reduced size and can rescue cdc25 mutant strains; however, niml is not required for cell viability (Russell and Nurse, 1987b). niml encodes a dual-specificity protein kinase, as evidenced by the ability of recombinant Niml protein to autophosphorylate on both tyrosine and serine residues (Coleman et al., 1993). In a cell free system, recombinant Niml phosphorylatesWeel specificallyin the C-terminal region which contains the catalytic domain. Therefore, Niml-specific phosphorylation results in a decrease in the ability of Weel to act as a negative regulator of Cdc2 kinase activity (Coleman et al., 1993) (Fig. 2). Since Weel kinase activity is down-regulated by phosphorylation,presumably there is a phosphatase that activates Weel. At this time, however, the Weel phosphatase is unknown. kinase X

niml (kinase)

0

= higher eukaryotes only

0

=yeast only

= negative regulator

+

= positive regulator

Figure 2. Regulatorsof the Cdc2 protein kinase. Schematic illustration of the regulators of Cdc2 kinase activity. Proteins that are circled indicate proteins that have been identified in higher eukaryotes only; the squared proteins have been identified exclusively in the yeast systems. The remaining proteins have been identified in both higher and lower eukaryotes. Protein phosphatases are indicated by p’tase.

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DEBORAH L. CHAPMAN and DEBRA J. WOLGEMUTH

D. Phosphatases: Cdc25 Cdc25 has been identified as a dose-dependent inducer of mitosis that functions in opposition to the Wee1 and Mikl protein kinases (Lundgren et al., 1991; reviewed in Millar and Russell, 1992). cdc25 mutant strains arrest in late G2, whereas strains overexpressing cdc25 divide at abnormally small sizes (Russell and Nurse, 1986; Gould and Nurse, 1989; Moreno et al., 1989). In cdc25 mutant strains, Cdc2 is phosphorylated and has low kinase activity (Booher et al., 1989). Because initially it did not reveal homology to any known phosphatase, cdc25 was not thought to encode the phosphatase responsible for activating Cdc2 (Russell and Nurse, 1986). However, evidence that cdc2.5 indeed encodes a tyrosine phosphatase was provided by the observation that a human protein tyrosine phosphatase complemented a cdc25 mutation in S. pombe by dephosphorylating the tyr-15 residue of Cdc2 and thus activating the kinase (Gould et al., 1990). cdc25 homologues have now been identified in a variety of organisms and are known to encode dual-specificity phosphatases capable of dephosphorylating proteins on tyrosine and serinehhreonine residues in vitro and in vivo (Dunphy and Kumagai, 1991; Gautier et al., 1991; Strausfeld et al., 1991; Lee et al., 1992). Dephosphorylation of Cdc2 at tyr-15 in fission yeast and at thr-14 and tyr-15 in animal cells results from Cdc25 activity (Russell and Nurse, 1987a; Dunphy and Kumagai, 1991; Gautier et al., 1991; Millar et al., 1991; Lee et al., 1991; Strausfeld et al., 1991; reviewed in Millar and Russell, 1992). In addition, murine (Sebastian et al., 1993) and Xenopus (Gabrielli et al., 1992b)Cdc25 can dephosphorylatethe thr-14 and tyr-15 residues of Cdk2, thus activating its kinase activity.

111. ROLE OF PROTEIN PHOSPHORYLATION IN GERM CELL DIFFERENTIATION A. Historical Perspective Observationsfrom various systems, including yeast, suggest that phosphorylation and dephosphorylation of proteins play important roles in the mitotic and meiotic cell cycles and the differentiation of germ cells. Extracts from mitotic HeLa cells contained phosphoproteins also present in other mitotic and meiotic cell types, but not in interphase cells (Davis et al., 1983). Exposure of Xenopus oocytes to progesterone results in a burst of protein phosphorylation shortly before GVBD (Maller et al.,

Expression and Function of Protein Kinases

13

1977). The qualitative changes of polypeptides found in resting Xenopus oocytes versus maturing oocytes have been demonstrated to involve phosphorylation (Maller and Smith, 1985; Karsenti et al., 1987). As in other systems, entry into M phase during mouse oocyte maturation is correlated with a burst of protein phosphorylation and activation of MPF as assayed by H1 kinase activity (Rime and Ozon, 1990). Treatment of mouse oocytes with an inhibitor of protein phosphorylation resulted in reversible inhibition of GVBD (Rime et al., 1989). Longer periods of incubation with the kinase inhibitor resulted in oocytes with condensed chromosomes but with an intact nuclear envelope, implying a role of protein phosphorylation in nuclear envelope breakdown (Rime et al., 1989; Motl'lk and Rimkevicovh, 1990). More recent studies have shown that oocytes allowed to undergo GVBD and then treated with the kinase inhibitor had decondensed chromosomes and had begun to reform the nuclear envelope, suggesting that protein phosphorylation is important in maintaining chromosomes in a condensed state (Motlik and Rimkevicov6, 1990). B. Use of Mouse as a Model System: Oogenesis and Spermatogenesis Germ cell development involves mitotic proliferation, meiotic recombination followed by reduction divisions, and subsequent differentiation to produce the highly specialized germ cells, the egg and the sperm. Although these events occur during both male and female germ cell development in mammals, the temporal pattern of their progression is quite different. In the mouse, primordial germ cells (PGCs), the germ cell precursors, enter the genital ridge by day 11.5 post coitum (P.c.). Once in the developing gonad, male germ cells continue to proliferate as they are surrounded by the developing seminiferous cords. During fetal development, male germ cells arrest in mitosis and continue to grow in size while in a quiescent phase. The male germ cells remain arrested until a few days after birth, at which time gonocytes resume mitosis and differentiate into spermatogonia. These spermatogonia undergo several mitoses and either remain as stem cells, renewing the population, or become committed to meiosis. Meiotically dividing primary spermatocytes undergo final replication of their DNA prior to entering into meiosis, including pre-leptotene, leptotene, zygotene, pachytene, and diplotene stages. In the mouse, day 8 of postnatal (p.n.) development marks the progression of the first wave of spermatogenetic cells into

14

DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

meiosis I (Nebel et al., 1961).After completing the first meiotic division, the secondary spermatocytesrapidly divide to form the haploid spermatids. By day 22 p.n., this first wave of germ cells has completed both meiotic divisions and has begun the complex morphological changes of spermiogenesis,which transform haploid spermatidsinto highly specialized spermatozoa (Nebel et al., 1961). Female mammalian germ cells, on the other hand, continue to undergo mitotic divisions after entering the embryonic gonad, and in most species, enter into meiosis, arresting at diplotene of meiosis I. At birth, these “resting” oocytes are found in primordial follicles, surrounded by a single layer of granulosa cells. Puberty is marked by the recruitment of a pool of oocytes into a growth period in response to cyclic variations in gonadotropins. During this growth period, the oocyte increases in size but remains arrested in prophase of meiosis I. Concomitantly, the number of granulosa cells surroundingthe oocyte increases. After completion of this growth and proliferative phase to form the mature follicle, the oocyte is stimulated to resume meiosis. Meiosis I is completed and the ovum is ovulated while arrested in metaphase of meiosis 11, awaiting fertilization to complete the second meiotic division (reviewed in Peters, 1969; Buccione et al., 1990).

IV. EARLY GERM CELL DIFFERENTIATION In the mouse, PGCs are derived from the epiblast (embryonic ectoderm). A group of approximately 100 PGCs can be detected in the mouse extra-embryonic mesoderm, just posterior to the definitive primitive streak in the day 7 p.c. animal (Ginsburg et al., 1990). By day 8 P.c., PGCs are found in the hindgut endoderm and at the base of the allantois. PGCs migrate from the allantois, along the hindgut, and reach the genital ridge by day 11.5 p.c. As they migrate, the number of PGCs increases to approximately 25,000 alkaline-phosphatase positive cells in the day 13.5 p.c. embryo, at which time they stop dividing (reviewed in McLaren, 1991). A. Kit and Steel Two existing mouse mutations have provided insight into the factors necessary for germ cell migration and proliferation. Mutations at either the dominant white spurring (W) or SteeE (SOlocus affect three migratory

Expression and Function of Protein Kinases

15

cell lineages: germ cells, hemopoietic stem cells, and melanocytes. Homozygotes for either mutation have a reduced number of germ cells in the genital ridge (Mintz and Russell, 1957; Russell, 1979; also see Besmer, 1991, and Morrison-Graham and Takahashi, 1993, for reviews). The W locus has been shown to encode the c-kit proto-oncogene, a receptor tyrosine kinase (Chabot et al., 1988; Geissler et al., 1988). SZ encodes a transmembrane growth factor that is the ligand of the c-kit receptor (Anderson et al., 1990a; Copeland et al., 1990; Flanagan and Leder, 1990;Huangetal., 1990;Martinetal., 1990;Williams etal., 1990; Zsebo et al., 1990a, b; reviewed in Witte, 1990). This ligand, also known as steel factor (SLF), stem cell factor (SCF), or mast cell growth factor (MGF), will be referred to as Steel (SI) in this review. Alternate splicing gives rise to two different mRNAs, both of which contain transmembrane domains but give rise preferentially to either the soluble or transmembrane protein in transfected cells (Anderson et al., 1990a; Flanagan et al., 1991). A soluble form of S1 is formed by proteolytic cleavage of the transmembrane domain. This soluble extracellular protein is the only form of S1 produced in the Steel dickie (Sld) mutants (Brannan et al., 1991; Flanagan et al., 1991). Sld/Sldmiceare sterile and severely anemic and lack pigmentation in the coat, indicating the importance of the transmembrane form of Sl in development. The soluble form of S1 can stimulate mast cell proliferation,whereas the transmembraneprotein can stimulate proliferation and mediate cell-cell adhesion of mast cells (Flanagan et al., 1991). Binding of the S1 to the Kit receptor results in autophosphorylation of the dimerized receptor and activation of the tyrosine kinase (Rottapel et al., 1991). Once activated, the kinase can phosphorylate cellular substrates, ultimately resulting in a cellular response. c-kit is expressed in embryonic hemopoietic tissues, melanoblasts, and PGCs, which is in good agreement with the observed mutant phenotype (Nocka et al., 1989; Om-Urtreger et al., 1990; Keshet et al., 1991; Manova and Bachvarova, 1991). In the early genital ridge, c-kit expression decreases when the PGCs stop dividing at approximately day 13.5 p.c. (Manova and Bachvarova, 1991). SZ is expressed during embryogenesis in the cells associated with the migratory pathways and homing sites of PGCs, melanocytes, and hematopoietic stem cells (Matsui et al., 1990; Keshet et al., 1991). Whereas the W mutation is cell autonomous, the S1 mutation affects the microenvironment of the migratory cell lineages. S1 has therefore been implicated in the homing mechanism for migratory cells expressing the c-kit receptor.

16

DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

Both the transmembrane and soluble forms of S1 can support the survival, but not the proliferation, of PGCs in vitro (Dolci et al., 1991; Godin et al., 1991). However, a combination of soluble S1 and leukemia inhibitory factor (LIF) has been shown to support the survival and in some cases the proliferation of PGCs in culture, but only for a short time (Matsui et al., 1991). The addition of basic fibroblast growth factor (bFGF) to this cocktail allows continued proliferation of PGCs and therefore long-term culture of PGCs in vitro (Matsui et al., 1992; Resnik et al., 1992). Although bFGF apparently can induce PGC proliferation in vim, it is not known whether this type of regulation of PGC proliferation occurs in vivo.

V. MITOTIC STAGES As previously discussed, in mammals the events of germ cell development exhibit sexual dimorphism with respect to their temporal progression. With few exceptions, all of the mitotic cell divisions of the female germ cells, oogonia, occur during embryonic development, while the male germ cell progenitors, Type Astern spermatogonia, continue to proliferate throughout most of the life of the adult animal. A. Kit and Steel

Both somatic and germ cell lineages express c-kit in the testis; c-kit mRNA (Manova et al., 1990; Sorrentino et al., 1991) and protein (Yoshinaga et al., 1991) have been detected in type A2 through type B spermatogonia up to the preleptotene stage of meiosis I, whereas Leydig cells at all stages express c-kit (Manova et al., 1990). In the female, c-kit mRNA and protein have been localized to oocytes in the embryo just before birth and to oocytes at all stages from primary oocytes through the fully grown oocyte (Manova et al., 1990; Horie et al., 1991). The presence of the Kit receptor on the surface of fully grown oocytes may indicate a function for the receptor during meiotic maturation, although injection of Kit monoclonal antibodies had no effect on oocyte maturation (Yoshinaga et al., 1991). S1 is expressed in the Sertoli cells of the adult testis (Motro et al., 1991; Rossi et al., 1991) and in the follicular cells of growing follicles (Keshet et al., 1991;Motro et al., 1991;Manova et al., 1993). In both the male and female, the S1-expressingcells (Sertoli

Expression and Function of Protein Kinases

17

and follicle cells) are in direct contact with those germ cells expressing c-kit (spermatogonia, oocytes). The differentially spliced SZ mRNAs, encoding either the soluble or transmembraneform of S1, are expressed in distinct patterns in the Sertoli cells during postnatal development of the mouse testis (Marziali et al., 1993). At day 6 postnatal the two &As were equally expressed, whereas in the adult, the smaller mRNA encoding the transmembrane S1 was the predominant transcript (Manova et al., 1993; Marziali et al., 1993). Several in vitro systems have been developed to assay S1function. In mast celVSertoli cell co-culture experiments, Sertoli cells derived from SZd/Sldmice could not support mast cell proliferation,whereas those derived from +/+ mice could (Tajima et al., 1991). Similarly, in primary Sertoli CelVspermatogoniaco-cultures, Sertoli cells derived from normal mice could bind spermatogonia, while the mutant Sertoli cells could not (Marziali et al., 1993). Microinjection of Sld/SZdSertoli cells with a plasmid expressing the transmembrane form of S1 rescued the binding defect (Marziali et al., 1993). Recent data from Rossi et al. (1993) further suggest arole for the KitlSl receptor/ligand in Sertoli cell regulation of spermatogenesis.Treatment of primary mouse Sertoli cell cultures with FSH, the physiological regulator of Sertoli cell function, results in an increase in both soluble and transmembrane Sl mRNAs in the Sertoli cells. This soluble S1 was found to stimulate DNA synthesis (S phase) in isolated cultures of type A but not type B spermatogonia.Yoshinaga et al. (199 1) had previously found that intravenousinjection of monoclonal antibodies against the Kit receptor depletes the seminiferous tubules of differentiating type A spermatogonia but has no effect on undifferentiated type A spermatogonia. Furthermore, intraperitoneal injection of Kit monoclonal antibodies completely blocks mitosis of differentiating type A spermatogonia but has no effect on the mitoses of gonocytes or primitive type A spermatogonia or the meiosis of spermatocytes.

B. Cyclin B Evidence for the involvement of a Cdc2/Cyc B complex in the mitotic stages of germ cell development comes from studies in DrosophiZu. Cyc B transcripts are abundant and uniformly distributed in the early Drosophilu embryo (Whitfield et al., 1989, 1990; Lehner and O'Farrell, 1990b).Maternally derived Cyc B transcripts are also concentrated at the posterior pole of the oocyte. Later, during embryogenesis, Cyc B tran-

18

DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

scripts become incorporatedinto the pole cells, the precursors of the germ cells, where the level remains high through embryonic development (Whitfield et al., 1989, 1990; Lehner and O'Farrell, 1990b; Raff et al., 1990).Sequencesin the 3' region of the maternally derived Cyc BmRNA are responsible for localizing the transcript to the posterior pole and for repression of translation (Dalby and Glover, 1992; 1993). Translation of the Cyc B mRNAin the pole cells is repressed until pole cell proliferation begins in the newly formed gonads. Zygotic transcriptionof CycB occurs in the gonads of the first instar larva and is presumably necessary for subsequent gonial stem cell division (Dalby and Glover, 1993). C. Phosphatases

As previously discussed, cdc25 encodes the dual-specificity phosphatase that dephosphorylates Cdc2 and Cdk2 at thr- 14 and tyr- 15, thereby activating its kinase activity. Homologues of cdc25 have been identified in various species. In the following paragraphs we present data supporting a role for Cdc25 in the mitotic division in higher eukaryotes and discuss the regulation of Cdc25 activity. The DrosophiZu cdc25 homologue string was identified by its ability '~ (Edgar and O'Farrell, 1989; to complement an S. pombe c d ~ 2 5mutant Jimenez et al., 1990). Bacterially expressed Drosophila string can dephosphorylate Cdc2Kyc B (pre-MPF), thus activating the kinase in a cell free system (Kumagai and Dunphy, 1991). During early Drosophilu embryogenesis, string expression corresponds to the pattern of zygotic cell division (Edgar and O'Farrell, 1989). Ectopic expression of string mRNA suggested that its activity is rate limiting in controlling the timing of embryonic cell divisions (Edgar and O'Farrell, 1990). string minus embryos undergo cell cycle arrest in GZ of cell cycle 14, following depletion of maternal mRNAs (Edgar and O'Farrell, 1989). string transcripts were also detected later, in the mitotically proliferating follicle cells during the early stages of oogenesis of the adult fly (Courtot et al., 1992). Thus far, three human cdc25 homologues have been identified: cdc25A (Galaktionov and Beach, 1991), cdc25B (Galaktionov and Beach, 1991; as CDC25Hu2 in Nagata et al., 1991), and cdc25C (as CDC25 in Sadhu et al., 1990; Galaktionov and Beach, 1991). Each of these human homologues can complementa cdc25" mutant in S.pombe. Anti-Cdc25C antibodies microinjected into HeLa cells inhibit entry into mitosis (Millar et al., 1991), whereas anti-Cdc25A antibodies arrest the cell at

Expression and Function of Protein Kinases

19

mid-mitosis (Galaktionov and Beach, 1991). These results indicate that in higher eukaryotes,different cdc25 genes may be functioning at distinct stages of the cell cycle and may activate distinct Cdc2/Cyc complexes (Galaktionov and Beach, 1991; Millar et al., 1991; reviewed in Millar and Russell, 1992). A murine homologue of cdc25, cdc25M2, shows highest homology to the human cdc25B and can complement an S. pombe cdc25" mutant (Kakizuka et al., 1992).Bacterially expressed Cdc25M2possesses phosphatase activity (Kakizuka et al., 1992). cdc25M2 is expressed in mouse embryos in a wide variety of differentiatingtissues that contain dividing cells. In the developing liver there appeared to be a correlation between the level of cdc25M2 expression and the level of mitoses; expression of cdc25M2 is high in the liver of day 13.5 p.c. embryos and decreases by day 16.5 p.c. (Kakizuki et al., 1992). cdc25M2 transcripts have also been detected in total RNA isolated from adult testis and ovary, suggesting a role for the phosphatase in mammalian gonadal function ( S . Wu and D. J. Wolgemuth, unpublished observations). In S. pombe, Xenopus, and human cells, Cdc25 activity has been shown to be regulated during the cell cycle by phosphorylation (Ducommun et al., 1990; Moreno et al., 1990 Kumagai and Dunphy, 1992; Hoffman et al., 1993). Hyperphosphorylation of the N-terminal region of Cdc25 at the onset of mitosis resulted in a dramatic increase in phosphatase activity, while dephosphorylation of the protein resulted in a decrease. The unphosphorylated form of Cdc25C was unable to induce maturation of Xenopus oocytes, while a stable, thiophosphorylated Cdc25C could (Hoffman et al., 1993). Thus, while Cdc25 controls the activity of a kinase, it is itself controlled by a kinase. In vitro studies demonstrated that the Cdc2/Cyc B kinase is responsible for Cdc25 phosphorylation, which could explain the self-amplification of MPF activity observed when MPF is injected into Xenopus oocytes (discussed in Hoffman et al., 1993). Okadaic acid, a potent inhibitor of serinehhreonine protein phosphatases type-1 (PP-1) and type 2A (PP-2A), completely blocked dephosphorylation of Cdc25C in Xenopus egg extracts, while specific inhibition of PP-1 had no effect (Clarke et al., 1993). The Cdc2/Cyc B kinase complex in these extracts was active as assayed by its ability to phosphorylate histone HI. These results suggest that PP-2A negatively regulates Cdc2/Cyc B kinase activity by dephosphorylatingthe activating phosphatase Cdc25C. In addition, the fission yeast type 2A-like protein phosphatase pp2a has been shown genetically to interact with

20

DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

cdc25 and weel, inasmuch as a deletion of pp2a can partially suppress a conditionallylethal mutation of cdc25 but is lethal when combined with a wee1 mutation (Kinoshita et al., 1993). As previously mentioned, activation of the Cdc2/Cyc B kinase complex is inhibited when DNA synthesis is blocked. In Xenopus egg extracts that have been supplemented with nuclei to mimic the presence of unreplicated DNA, the block to CdcYCyc B activation can be overcome by the addition of stably phosphorylated Cdc25C or by treatment with okadaic acid (Clarke et al., 1993). These results further suggest that inhibition of Cdc2/Cyc B kinase activity, due to the presence of unreplicated DNA, can be accomplished by maintaining Cdc25C in a dephosphorylated and hence inactive state, probably by keeping the level of PP-2Aactivity toward Cdc25C elevated. Other phosphatases have also been identified and may be implicated in mitotic and meiotic germ cell functions. For example, INH was originally isolated from a Xenopus oocyte cell free system as an inhibitor of pre-MPF activity (Cyert and Kirschner, 1988). ZNHencodes a protein phosphatase 2A that negatively regulates MPF activity by dephosphorylating Cdc2 on thr-161 (Lee et al., 1991; Solomon et al., 1990).

D. M A P Kinase MAP kinase (MAPK; also known as microtubule-associatedprotein-2 kinase), mitogen-activated protein kinase, and extracellular signal regulated kinases (ERKs) constitute a group of serine/threonine protein kinases of 40-45 kDa that are activated in response to various extracellular stimuli in a wide variety of cell types (Rossomando et al., 1991; also see reviews in Cobb et al., 1991; Pelech and Sanghera, 1992; Thomas, 1992).MAPKs play a central role in various signal transduction pathways (Fig. 3). Extracellularligands (insulin, growth factors) bind to and activate receptor tyrosine kinases, which ultimately lead to mitosis via activation of MAPK. MAPK is believed to be involved in both the re-entry of quiescent cells into the cell cycle (GdG1phase transition) and in M phase of oocyte maturation. Targets of MAPK include structural proteins, microtubule-associated protein-2 (MAP2), and myelin basic protein (MBP); transcription factors c-myc, c-jun, and ~ 6 2 ~ and ' ~ ;two serine/threonine protein kinases, S6 kinase I1 (rsk) and MAP kinaseactivated protein kinase-2 (MAPKAPkinase-2) (Stokoe et al., 1992; see Howe et al., 1992, and references therein). The levels of MAPK mRNA do not appear to change during the cell cycle; instead, regulation of MAPK activity occurs post-translationally,

Expression and Function of Protein Kinases

21

via phosphorylation (Gotoh et al., 1991b). MAPK activity is dependent on phosphorylation on both tyrosine and threonine residues, since it is inactivated by treatment with serinehhreonineand tyrosine phosphatases (Ahn et al., 1990; Anderson et al., 1990b; Crews et al., 1991; Seger et al., 1991; Wu et al., 1991; Posada and Cooper, 1992). MAPK is not phosphorylated on tyrosine residues in immature Xenopus oocytes, becomes phosphorylated during oocyte maturation, and returns to the dephosphorylated and inactive state approximately 30 minutes after fertilization(Gotohetal., 1991a;Ferrelletal., 1991;Posadaetal., 1991). MPF can activate MAPK in both Xenopus oocytes and cell free extracts of interphase eggs (Gotoh et al., 1991b). This presumably involves an intermediate kinase, since treatment of oocytes with cyclohexamide prior to injection with MPF did not affect tyrosine phosphorylation of MAPK (Ferrell et al., 1991). Furthermore, MPF cannot directly phosphorylate and activate purified MAPK that has been dephosphorylated by treatment with phosphatase 2A, a serinekhreonine protein phosphatase (Gotoh et al., 1991b). Therefore, although protein synthesis is not required for MPF-induced activation of MAPK, an intermediate between MPF and MAPK must exist. This intermediate MAPK activator (MAPK kinase, MAPKK) is a 45 kDa phosphoprotein capable of phosphorylating MAPK on serine/threonine and tyrosine residues (Matsuda et al., 1992; Nakielny et al., 1992a; Kosako et al., 1993). Like MAPK, the activity of MAPKK is regulated by phosphorylation. During oocyte maturation MAPKK is phosphorylated on threonine residues (Kosako et al., 1992), and this phosphorylation is required for its activity (Ahn et al., 1991; Gomez and Cohen, 1991; Kosako et al., 1992; Matsuda et al., 1992). MPF can activate both MAPKK and MAPK in vitro,with the activation of MAPK lagging behind that of MAPKK; however, MPF cannot activate either purified MAPKK or MAPK that has been dephosphorylated by phosphatases (Matsuda et al., 1992). MAPKK and MAPK are therefore believed to function downstream of MPF (Fig. 3). Recent data from yeast have shed some light on the MAPK signal transduction pathway. In S. cerevisiae,the pheromone-dependent transmission of signals necessary for mating involves the redundant protein kinases FUS3 and KSSl, which are members of the MAPK family (Boulton et al., 1990; Elion et al., 1991; Gartner et al., 1992). Genetic analysis indicated that FUS3 activity requires STE7 and STEll (Gartner et al., 1992;Stevenson et al., 1992).FUS3 has been shown to be activated by STE7 in vitro (Errede et al., 1993). STE7 is related to MAPKK

22

DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

c-ras c-raf

4 / 4

MAPKK

MPF

MAPK

transcrktion factors c-myc c-jun

kinases

x

MAPKAP kinase

~ 6 2 ~ ' ~ ribosomal S6

MAP2 MRP

C-fOS

Figure 3. MAP kinase regulatory pathway. The MAP kinase signaling pathway begins with activation of the receptor tyrosine kinase (RTK) by exogenous signals, such as growth factors and insulin. The signal is then transmitted into the cell via activation of the Raf serine/threonine kinase either directly by the RTK or through the GTP-binding protein, Ras. The signal is then transmitted to the nucleus and to other cytoplasmic proteins via MAPKK and MAPK.

(Kosako et al., 1992, 1993; Nakielny et al., 1992b) and, like MAPKK, STE7 is regulated by phosphorylation, which may occur via STEll (Cairns et al., 1992; Zhou et al., 1993). Proteins in both fission and budding yeasts (byrl, wisl, PBS2, STE7) are related to MAPKK (Kosako et al., 1992; 1993). These results indicate a conservation of the kinase cascade in eukaryotes (reviewed in Sprague, 1992). E. Raf

The c-ruf-1 proto-oncogene encodes a 74 kDa (Raf- 1) cytoplasmic serinekhreonine protein kinase. The role of Raf- 1 in signal transduction has been reviewed in Rapp (1991) and Heidecker et al. (1992). Phosphorylation of Raf-1 and its subsequent activity are increased by treatment of cells with growth factors or mitogens (Morrison et al., 1988; Rapp, 1991). Raf-1 has been shown to associate with the PDGF receptor in a ligand-dependent manner, with autophosphorylation of the receptor being required for the association (Morrison et al., 1989; App et al., 1991) and subsequent tyrosine phosphorylation of Raf-1 (Momson et al., 1988,

Expression and Function of Protein Kinases

23

1989). These data indicate that Raf-1 acts downstream of growth factor receptors in the signal transduction cascade. Where Raf-1 acts in this cascade is not clear, but evidence is mounting that indicates it acts downstream of rusl and upstream of MAPKK (reviewed in Roberts, 1992). Although Raf-1 shows no homology to STEll, a S. cerevisiue activator of MAPKK, it can activate both MAPKK and MAPK (Dent et al., 1992; Howe et al., 1992; Kyriakis et al., 1992). With regard to the potential function of Raf-1 in mammalian germ cells, the testis is a site of abundant expression of c-ruf-1 mRNAs. Northern blot and in situ hybridization analysis has shown that although c-ruf-1 is expressed most abundantly in early pachytene spermatocytes, some transcripts were also detected in germ cells from type A and B spermatogoniathrough to the round spermatid stage (Wolfes et al., 1989; Wadewitz et al., 1993).

VI. MEIOTIC STAGES Meiosis by definition constitutes unique aspects of cell cycle regulation; for example, there is no intervening DNA synthesis (S phase) between the first and second metaphase stages. During the second reduction division of meiosis the requirement for S is bypassed, such that there is an uncoupling of S and G2/M. In Xenopus oocytes, activation of meiosis via dephosphorylationof Cdc2 at thr-14 and tyr-15 is linked to exposure of the oocytes to progesterone, an external signal. This is in opposition to the internal signal (i.e., completion of S phase) in mitotically dividing cells. A. MPF Mouse oocytes can undergo GVBD in the presence of protein synthesis inhibitors (Wassarman et al., 1976; Schultz and Wassarman, 1977; Hashimoto and Kishimoto, 1988), whereas GVBD in Xenopus oocytes requires new protein synthesis (reviewed in Maller, 1985). New protein synthesis is required for meiosis I1 in both organisms (Gerhart et al., 1984; Hashimoto and Kishimoto, 1988). Activation of Xenopus oocytes by progesterone or injection of MPF results in increased protein synthesis followed by GVBD; however, the induction of these events occurs more rapidly with MPF injection than with progesterone (Wasserman et al., 1982). Furthermore, the pattern of protein phosphorylation from

24

DEBORAH L. CHAPMAN and DEBRA J. WOLGEMUTH

progesterone- and MPF-induced maturation of Xenopus oocytes is identical (Wasserman et al., 1982). These results suggest that progesterone and MPF are part of the same signal transduction pathway that brings about oocyte maturation, and that MPF is downstream of progesterone in this pathway. Cyc B is a necessary regulatory component of MPF. Xenopus and sea urchin oocytes contain a stockpile of Cyc B associated with Cdc2 in a complex known as pre-MPF, which is kept inactive by phosphorylation of Cdc2 on thr-14 and tyr-15 residues (Edgecomb et al., 1991; Gautier and Maller, 1991; Kobayashi et al., 1991). Injection of antisense cycBI, cycB2, and cycA oligonucleotides into Xenopus oocytes did not prevent progesterone-induced maturation; therefore, synthesis of new cyclin is not required forxenopus oocyte maturation (Minshull et al., 1991). MPF activity during mouse and Xenopus oogenesis has been assessed by injecting cytoplasm from oocytes at different stages into immature oocytes and by histone H1 kinase assays of cellular lysates (Gerhart et al., 1984; Hashimoto and Kishimoto, 1988; Choi et al., 1991; Fulka et al., 1992). MPF activity was low in immature oocytes, peaked at metaphase I and 11, and was low again immediately following metaphase I and 11. Furthermore, MPF activity was highest in cells containing the hypophosphorylated forms of Cdc2, thus confirming previous observations that the hypophosphorylated Cdc2 is the active form (reviewed in Clarke and Karsenti, 1991, and Fleig and Gould, 1991). Murine Cdc2 is not phosphorylated in the late phases of the first and second meiotic cell cycles (Choi et al., 1992). Treatment of oocytes in the late phase of meiosis I or in meiosis I1 with sodium orthovanadate, an inhibitor of tyrosine phosphatase activity, did not inhibit histone H1 kinase activity. However, it was able to inhibit kinase activity when oocytes were treated at early stages of meiosis I. Choi et al. (1992) suggest that the oocyte escapes the normal requirement for S phase prior to M phase by not phosphorylating Cdc2 in late meiosis I or in meiosis 11.

B. Cyclin B The four different B-type cyclins identified in S.cerevisiae (Clbsl4) may have distinct functions during the mitotic and meiotic cell cycles. While both Clb3 and Clb4 appear to have a role in the S phase of the mitotic cell cycle, they are not involved in the premeiotic S phase (Grandin and Reed, 1993). Instead, cZb3 and cZb4, as well as cZbl (an M phase cyclin) were expressed and activated the CDC28 kinasejust before

Expression and Function of Protein Kinases

25

the first meiotic division. Furthermore, while both Clbl and Clb2 are involved in the M phase of mitotic cycles, only Clbl is involved in meiosis. In studies designed to identify cyclins involved in murine germ cell development, cDNAs encoding the murine cyclin B 1 (cycBl)and cyclin B2 (cycB2) were isolated from an adult mouse testis cDNA library (Chapman and Wolgemuth, 1992, 1993). Northern blot and in situ hybridization analyses revealed that both cycBl and cycB2 were expressed in the germ cell compartment of the testis. While cycB2 was expressed primarily in pachytene spermatocytes, cycBl was expressed predominantly in the early round spermatids. The pattern of cycB2 expression in the testis conforms to the predicted cyclin function in meiotic cell division. Section VI1.B provides a continued discussion of cycBl expression in the post-meiotic germ cells in the testis. Interestingly, in the ovary in situ analysis revealed cycB2 transcripts in both germ cells and somatic cells, specifically in the oocytes and granulosa cells of growing and mature follicles. cycBl transcripts were localized primarily to the oocytes of growing and mature follicles, with a much lower level detected over the granulosa cells of large follicles. The pattern of cycBl and cycB2 expression in ovulated and fertilized eggs was also examined by in situ hybridization analysis. While the steady-state level of cycBl and cycB2 signal remained constant in oocytes and ovulated eggs, the signal of both appeared to decrease following fertilization. In addition, both cycBl and cycB2 transcripts were detected in the cells of the inner cell mass and the trophectoderm of the blastocyst. The lineage- and developmental-specificexpression of cycBl and cycB2 in the ovary and testis contrasts with theirco-expression in the blastocysts.

C. CSF In the oocyte, active MPF kinase triggers GVBD, chromosome condensation, and spindle formation. Following extrusion of the first polar body and ovulation, vertebrate oocytes are arrested at the metaphase of meiosis 11, presumably because of high MPF activity, which prevents the cell from exiting M phase. Fertilization or artificial activation of these eggs releases them from meiotic arrest and results in completion of metaphase 11.The presence of a meiotic inhibitor in unfertilizedxenopus eggs was demonstrated by injecting the cytoplasm of an egg into a blastomere of a two-cell embryo (Masui and Markert, 1971). This

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DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

cytoplasm caused cleavage arrest of the blastomere at the metaphase stage. This cytostatic factor, CSF, is believed to function by stabilizing active MPF (reviewed in Masui, 1991). CSF has been characterized as a calcium-sensitive factor in Xenopus eggs, the activity of which is overcome by an influx of free calcium at fertilization (Watanabe et al., 1989). CSF activity first appears at GVBD of meiosis I and remains high until fertilization (Watanabe et al., 1989). A CSF-like activity is also present in mouse oocytes, as shown by the arrest at metaphase of blastomeres that have been fused with maturing mouse oocytes (Balkier and Czolowska, 1977). Thus CSF, like MPF, is a highly conserved activity; however, it is not clear what the components are. We discuss below the evidence that suggests that at least part of CSF may be encoded by c-mos.

D. Mos c-mos, the cellular homologue of the Moloney murine sarcoma virus transforming gene, encodes a cytoplasmic serine/threonine protein kinase. The site of expression of c-mos in normal tissues had eluded detection until the gonads of adult animals were examined. High levels of c-mos RNAs are restricted to the germ cells of the testis and ovary of adult mice (Propst et al., 1987). A series of studies employing Northern blot and in situ hybridization analysis (Goldman et al., 1987;Mutter and Wolgemuth, 1987; Propst et al., 1987; Iwaoki et al., 1993) established that expression of c-mos is further restricted to specific stages of germ cell differentiation. In the male, c-mos is most abundant in early round spermatids and drops off as the spermatids differentiate. In the female, c-mos RNA is not detectable in primary resting oocytes, but accumulates soon after the oocyte enters the growth phase, reaching a level in fully grown oocytes that has been estimated to be several orders of magnitude higher than that found in early spermatids (Goldman et al., 1987). c-mos RNA levels drop between metaphase I and I1 and drop to undetectable levels aEter fertilization and early embryogenesis (Mutter et al., 1988). These initial observations of this highly restricted pattern of expression of c-mos suggested a possible role during meiosis or post-meiotic events in both the male and female germ line. A critical advance in the studies of the function of c-mos in germ cells, especially oocytes, was the identification of the Xenopus homologue of the murine gene. Since fully grown Xenopus oocytes arrested in prophase of meiosis can be collected in large numbers and induced to undergo maturation in vitro in response to progesterone, biochemical analysis of Mos function was feasible.

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c-mos expression is required for progesterone-induced GVBD of Xenopus oocyte maturation (Sagata et al., 1988). Injection of in vitro synthesized Xenopus c-mos RNA into oocytes can induce maturation in the absence of progesterone (Freeman et al., 1989; Sagata et al., 1989a). Antisense experiments further demonstrated that translation of mos mRNA is required for progression from metaphase I to metaphase I1 in Xenopus oocytes (Daar et al., 1991; Kanki and Donoghue, 1991). Injection of c-mos RNA into a two-cell Xenopus embryo results in cleavage arrest at metaphase and high MPF activity (Sagata et al., 1989a). Furthermore, egg cytosol immunodepleted of Mos no longer has the capacity to arrest the cell cycle when injected into two cell embryos (Sagata et al., 1989b). Together, these results suggested that Mos is part of the CSF activity described by Masui (Masui and Markert, 1971). These results indicate that the Mos kinase has two roles in oocyte maturation: 1) induction of GVBD in oocytes and 2) arrest at metaphase I1 in the mature egg. Injection of antisense c-mos oligonucleotides into GV stage mouse oocytes revealed an inhibition of first polar body emission (Sagata et al., 1988; Paules et al., 1989). Using a similar experimental approach, O’Keefe et al. (1989) showed that oocytes completed metaphase I but there was no initiation of metaphase 11. In these oocytes, chromosomes began to decondense, some of the DNA underwent replication, the nuclear membrane reformed, and cleavage to the two-cell stage occurred. The discrepancies in these experiments may be due to slight differences in the timing of injection: the first two studies may have employed injection of the antisense oligonucleotides at a slightly earlier time in meiosis I. Mos antibodies injected into fertilized Xenopus eggs have revealed that antibodies that inhibit Mos kinase activity also arrested the zygote at the pronuclear stage, whereas antibodies that had no effect on the kinase activity allowed cleavage and entry into the two-cell stage (Zhao et al., 1991). Therefore, Mos protein kinase activity is required for the completion of pronuclear breakdown following fertilization. In Xenopus oocytes, synthesis of Mos occurs only during progesteroneinduced oocyte maturation (Sagata et al., 1988). Prior to GVBD, Mos is only partially phosphorylated and is highly unstable. During and after GVBD, Mos becomes progressively phosphorylated, coincident with its increased stability in metaphase I1 arrested eggs. Phosphorylation at ser-3 appears to be responsible for the stability of the Mos in metaphase I1 arrested eggs, as assayed by injecting various mutated c-mos RNAs

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DEBORAH L. CHAPMAN and DEBRA 1. WOLGEMUTH

into Xenopus oocytesand examining their stability following progesteroneinduced maturation (Nishizawa et al., 1992). Furthermore, whereas wild-type Mos was stable and phosphorylated at ser-3 in metaphase TI arrested eggs, kinase-inactiveMos was unstable and not phosphorylated on ser-3. These results suggest that Mos autophosphorylateson ser-3 to form the stable protein found in the mature egg. These results are consistent with the results of Freeman and colleagues (1992) wherein ser-3 was identified as a major phosphorylation site in vivo in Xenopus. Interestingly, when assayed for their ability to induce meiotic maturation, the non-phosphorylatable mutant Mos was able to elicit GVBD. Thus GVBD-inducing Mos and the metaphase 11-arresting Mos are different forms of the same protein and may have different substrate specificities at these two distinct times in oocyte maturation (Nishizawa et al., 1992). It has been suggested that Mos might function to stabilize MPF via phosphorylation of the Cyc B subunit (Sagata et al., 1989b; reviewed in Hunt, 1992). For example, introduction of a proteolysis-resistantCyc B mutant caused cell cycle arrest at metaphase in both Xenopus eggs and embryos (Murray et al., 1989). Furthermore, mouse eggs microinjected with antisense c-mos-specific oligonucleotides failed to reactivate MPF and accumulate Cyc B following the completion of meiosis I1 (O’Keefe et al., 1991). Immunoprecipitates of Mos from Moloney murine sarcoma virus-transformedNM 3T3 cells and Xenopus eggs have been shown to phosphorylate Cyc B in vitro (Roy et al., 1990). c-rnos-specific antisense oligonucleotides resulted in the reduction of Cyc B phosphorylation in Xenopus oocyte extracts (Roy et al., 1990). However, the induction of oocyte maturation does not occur viaphosphorylation of cyc B, as shown by the following experiments. Although coinjection of c-mos and cyc B into Xenopus oocytes accelerates meiotic maturation, there was no increase in Cyc B phosphorylation in vivo (Freeman et al., 1991). Cdc2 can also phosphorylate CycB 1 and CycB2 on serine residues in Xenopus eggs (Izumi and Maller, 1991). However, replacing these serine residues with non-phosphorylatable residues does not result in functional differences between mutant and wild-type cyclins (Izumi and Maller, 1991). No changes in the kinetics of M phase Cyc B degradation or the induction of mitosis or meiosis were observed, demonstrating that in Xenopus, phosphorylation of Cyc B by Cdc2 is not required for cyclin function or for its degradation following meiosis or mitosis.

Expression and Function of Protein Kinases

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E. Cdk2 Several observations suggest a role for Cdk2, possibly in cooperation with Mos, to arrest eggs at metaphase 11. In the meiotic cell cycle of Xenopus oocytes, Cdk2 kinase activity is low at GVBD in meiosis I and increases by metaphase I1 (Gabrielli et al., 1992a). Xenopus oocytes injected with antisense cdk2 oligonucleotides and then stimulated with progesterone underwent GVBD and completed meiosis I but failed to arrest at metaphaseI1and entered the first embryonic cell cycle (Gabrielli et al., 1993). These eggs failed to arrest at metaphase I1 even though active Mos was present. Injection of purified Cdk2 into these oocytes could restore the arrest at metaphase I1 with high Cdc2 kinase activity. Progesterone-treated oocytes that were allowed to progress through GVBD and then injected with cyclohexamide and purified Cdk2 arrested at metaphase I1 with high H1 kinase activity and stable Cyc B2, whereas oocytes injected with cyclohexamide alone lost H 1 kinase activity and Cyc B2 was degraded (Gabrielli et al., 1993). These results indicate that following completion of GVBD, Cdk2 is the only protein needed to be synthesized to arrest eggs at metaphase 11. It would be interesting to know whether Cdk2 kinase levels are also elevated in secondary spermatocytes, where the metaphase divisions do not have arrest control points.

F. M015 M015 was isolated in a screen to identify kinases that were involved in meiotic maturation in Xenopus oocytes (Shuttleworth et al., 1990). M015 encodes a 40 kDa protein that shares 40% amino acid identity with human Cdc2 kinase. M015 is most likely a Cdk2 kinase rather than a Xenopus cdc2 homologue. M 0 1 5 transcripts accumulate during oogenesis and are deadenylated during meiotic maturation and degraded during embryogenesis following the mid-blastula transition. M015 is largely restricted to immature oocytes as deadenylation results in the reduction of M015 translation. Injection of antisense-specific MOI5 oligonucleotides into Xenopus oocytes followed by treatment with progesterone resulted in acceleration in the rate of H1 kinase activation and maturation. These results suggest that M015 may act as a negative regulator of meiotic maturation in Xenopus.

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DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

G. Cdc25 Class of Phosphatases

twine encodes the second Drosophilu cdc25 homologueto be identified (Alphey et al., 1992; Courtot et al., 1992). Both string and twine are abundantly expressed in nurse cells, during oogenesis, with maternal transcripts persisting through to the syncitial stage of embryonic development (Alphey et al., 1992; Courtot et al., 1992). However, whereas string transcripts were detected in both somatic and germ cells, twine expression is restricted to the germline. mine transcripts were not detected in mitotically dividing germ cells or in the mitotically proliferating follicle cells. In the testis, twine transcripts were detected in the growing stage of the pre-meiotic cysts (primary and secondary spermatocytes), but not after meiosis I1 in the haploid germ cells. twine is essential for male and female fertility. In flies carrying a missense mutation in twine, meiosis does not occur in the male; the testis contains cysts of 16 rather than 64 spermatids with 4N DNA content. Signs of sperm differentiation appear; for instance, there is elongation of the sperm tail and compaction of the nuclear DNA into the sperm head, but the sperm heads are larger and the DNA is never as tightly bundled as in the wild type. Furthermore, no motile sperm were present in the seminal vesicles. In the female mutants, a variability in the meiotic defects was observed. DNA appeared compacted and nuclear envelope breakdown and formation of spindles occurred. Although entry into meiosis appeared to occur at the correct stage, meiosis was not complete. Jessus and Beach (1992) showed that the level of the Xenopus Cdc25 did not oscillate during meiotic maturation or during early embryonic cell divisions. Instead, the association of Cdc25 with the Cdc2Kyc B complex (MPF) did oscillate in a cell cycle dependent manner. Maximal association coincided with maximal Cdc2 kinase activity. Cyc B can directly activate the intrinsic protein tyrosine phosphatase activity of Cdc25 proteins in the absence of Cdc2, suggesting that interactions between the Cdc25 and the Cdc2/Cyc B complex probably occurs via Cyc B (Galaktionov and Beach, 1991). H. Novel Kinases: mak and Nekl

Amurine male germ cell-assocatedkinase (rnak)was isolated by virtue of its weak homology to the v-ros tyrosine kinase (Matsushime et al., 1990). Although it was isolated because of its homology to a tyrosine kinase, sequence analysis revealed that it belongs to the serinehhreonine

Expression and Function of Protein Kinases

31

kinase gene family. Mak is 40% identical to the yeast and human Cdc2. Northern analysis revealed mak expression in male meiotic and postmeiotic germ cells in both rats and mice. The expression pattern of mak and its homology to cdc2 implies a role for Mak in the meiotic and post-meiotic processes. Nekl was identified by screening mouse cDNA expression libraries with anti-phosphotyrosineantibodies (Letwin et al., 1992). It encodes a protein that is 42% identical to the serinehhreonine protein kinase domain of the Aspergillus nidulans protein kinase NIMA (NIM = “never in mitosis”). NIMA has been shown to control the initiation of mitosis in A. niduluns (Osmani et al., 1988). Three different nimA mutants arrest in the G2 phase and contain Cdc2 with an unphosphorylated tyr-15 residue (Osmani et al., 1991). Although this form of Cdc2 is an active kinase, as shown by H1 kinase assay, these mutants cannot initiate mitosis. These results suggest that the Cdc2 and NIMA protein kinases act in parallel on separate pathways to initiate mitosis (reviewed in Murray, 1991). Blocking either pathway can arrest mitosis, even in the presence of the other fully active kinase. Bacterially expressed Nekl can phosphorylate exogenous substrates, mainly on serine/threonine residues but also on tyrosine residues, and therefore encodes a dual-specificity kinase (Letwin et al., 1992). Nekl transcripts can be detected in the developing gonads of day 15.5 p.c. embryos, in the prospermatogonia and the Sertoli cells of the testis and in the oocytes of the ovary. The embryonicovary at day 15.5p.c. contains oocytes that are in zygotene and pachytene of meiosis I. In the adult testis, Nekl signal was detected in Sertoli cells, at a low level in the spermatogonia and at higher levels in spermatocytes and early round spermatids. In the adult ovary, Nekl transcripts were detected in growing oocytes up to ovulation and in the proliferatinggranulosacells, but were not detected in primary follicles. The early corpus luteum also expresses Nekl.

VII. POST-MEIOTIC EVENTS The striking difference between the temporal progression of meiosis in the male and female germ cells extends to post-meiotic stages as well. The egg is essentially frozen in metaphase 11, awaiting the signal of fertilization to continue with the reduction division, whereas the male gamete proceeds to complete metaphase I1 and begins to differentiate. The differentiation events that follow transform somatic-appearingcells

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DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

into the highly specialized, motile spermatozoa. This stage does not represent a defined stage of the cell cycle per se; however, especially in the male, considerable changes in the organization of the nuclear components occur, which to a certain extent resemble nuclear changes during metaphase in mitotic cells. For example, the chromatin becomes highly compacted and reorganized. Spermiogenesis in mammals and other vertebrates is characterized by the systematic replacement of the somatic-type histones with testis-specific histones and transition proteins and, ultimately, the complexing of DNA with the highly basic protamines (Goldberg et al., 1977). A model for chromatin packaging in mouse sperm suggests a function for protamine phosphorylation and dephosphorylation (Balhorn et al., 1982). A. Murine c-abl Although c-abl, which encodes a non-receptor tyrosine kinase, is expressed at a low level in all mouse tissues examined (Wang and Baltimore, 1983), including male and female germ cells (Iwaoki et al., 1993), a unique -4-kb transcript is expressed in haploid spermatids (Ponzetto and Wolgemuth, 1985). This testis-specific transcript apparently has a 5’ sequence identical to the larger somatic transcript (Meijer et al., 1987; Oppi et al., 1987), although the 3’ end lacks -1 kb of untranslated sequences and utilizes an alternative polyadenylation site that lacks an AU-rich destabilization sequence (Meijer et al., 1987; Duggal et al., 1987). This may account for the increased stability of the message, which is observed at the highest level in elongating spermatids (Meijer et al., 1987; Iwaoki et al., 1993). Only a portion of the testis-specific 4-kb c-abl transcript is associated with polysomes; the remainder is associated with ribonucleoprotein particles and monosomes, suggesting that it is stored for translation at a later stage of development (Zakeri et al., 1988). The predicted amino acid sequence from cDNAs (Meijer et al., 1987; Oppi et al., 1987), as well as studies on c-Abl proteins (Meijeret al., 1987; Ponzetto et al., 1989),indicate that the 4-kb transcript produces a protein similar in size (-150 kDa) to that resulting from the Type I somatic transcript. Abl is phosphorylated on serine and threonine residues in vivo but normally not on tyrosine residues (Pendergast et al., 1987). The phosphorylation status of Abl varies in a cell-cycle-specificmanner. In NIH 3T3 cells, Abl is phosphorylated on three sites during interphase and seven additional sites during mitosis (Kipreos and Wang, 1990). Two of the interphase sites and all of

Expression and Function of Protein Kinases

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the mitotic sites could be phosphorylated in vitro by the Cdc2 kinase. Since Cdc2 has been shown to play a possible role in the regulation of Abl during the cell cycle, it would be of interest to determine whether Cdc2 plays a similar role in non-proliferating spermatids. B. Murine cycB1 As previously mentioned, Northern blot and in situ hybridization analysis revealed that the highest level of cycBl expression in the testis was in the post-meiotic germ cells. Based on what is known about cyclin function in cell division, the high level of cycBl transcripts in the early round spermatids was initially surprising. These cells have completed the mitotic and meiotic divisions of gametogenesis and are about to undergo spermiogenesis, a process of morphogenetic differentiation, to yield the mature sperm. However, observations from other organisms may provide clues to a possible function for a Cdc2/Cyc B complex in germ cells at these stages. For example, in the sea urchin, sperm-specific histones HI and H2B are phosphorylated on serine residues found within a consensus sequence similar to that of Cdc2 (Hill et al., 1990). These histones are phosphorylated in spermatids but not in the mature sperm. The sequence of histone phosphorylation and dephosphorylationduring spermiogenesismay contribute to the fine tuning of chromatin packaging in the sea urchin sperm head. During trout spermiogenesis, phosphorylation of histones at serine residues within a Cdc2 consensus sequence is believed to alter their ability to bind DNA, thus facilitating the replacement of the histones by protamines (Sung and Dixon, 1970). Interestingly, mouse histone H1, transition protein 2 (TP2), and protamine 2 (mP2) contain consensus sequences for phosphorylation by Cdc2. An intriguing possibility is that the cyclin observed in the early spermatids is involved in phosphorylatingthese proteins by complexing with Cdc2 or Cdk2 to target kinase activity. C. Murine raf

B-ruA a member of the rufgene family of serine/threonine kinases, is expressed as two major transcripts of 4.0 kb and 2.6 kb in the mouse testis (Wadewitz et a]., 1993). B-rufexpression is limited to the germ cells and is particularly abundant in early spermatids. Northern hybridization analysis revealed that the two B-ruftranscripts are expressed in a stage-specific manner. Low levels of the 4.0-kb transcript are first

34

DEBORAH L. CHAPMAN and DEBRA J.WOLGEMUTH

detected in meiotic germ cells, whereas the 2.6-kb transcript is restricted to post-meiotic spermatids. c-ruf-1 and its related gene, A-ruJ are both expressed in the mouse epididymis. c-ruf- 1 is expressed as a 3.1-kb transcript at a uniform level throughout the length of the epididymis in all epithelial cell types (Winer et al., 1993). In contrast, the 2.6- and 4.3-kb A-raftranscripts are present in a segment-specificpattern in the epididymis.The highest level of A-ruf is in the principal epithelial cells of the proximal caput and decreases progressively in more distal regions. The high level of A-raftranscripts in the proximal caput suggeststhat A-rufmay play a key role in regulating the function of this particular region of the epididymis and, potentially, sperm function.

VIII. FERTILIZATION In the mouse, binding of the sperm to the zona pellucida (ZP) results in the acrosome reaction. The acrosome reaction involves fusion of the plasma and outer acrosomal membranes and subsequent release of the acrosomal contents. This process may be thought of as a signal transduction pathway: ligand (in the ZP) binds to a receptor (in the sperm) resulting in the transmission of the extracellular signal to the intracellular domain and ultimately causing an effect, in this case the acrosome reaction. A specific zona pellucida glycoprotein (ZP3), which binds to the plasma membrane overlying the acrosome, is believed to induce the acrosome reaction (Florman and Wassarman, 1985). '251-labeledZP3, when used to probe a Western blot of sperm proteins, recognized two major ZP binding proteins of 95 and 42 kDa (Leyton and Saling, 1989). Antibodies against phosphotyrosine also recognized a 95 kDa protein, in addition to 52 and 72 kDa proteins, in Western blots of sperm proteins (Leyton and Saling, 1989). By indirect immunofluorescense, the 52 and 72 kDa phosphoproteins were detected exclusively in capacitated sperm, while the 95 kDa protein was detected in both fresh and capacitated sperm. These results demonstrate that the 95 kDa protein can bind ZP3 and can serve as a substrate for a tyrosine kinase. Following exposure to the ZP, there is an increase in the number of cells with anti-tyrosine reactivity (Leyton and Saling, 1989). There is an increase in sperm membrane fluidity following capacitation; therefore, ZP3 may act to aggregate the 95 kDa proteins. Furthermore, if p95 encodes a receptor tyrosine kinase, then aggregation of the receptors via ZP3 could result

Expression and Function of Protein Kinases

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in autophosphorylationof the receptor. Recent data indicates that the 95 kDa protein, p95spERM, has tyrosine kinase activity that is stimulated by exposure to solubilized ZP (Leyton et al., 1992).Furthermore, inhibition of the ~ 9 5 kinase ” ~ activity ~ also inhibits ZP triggered exocytosis (acrosome reaction) in a dose-dependent manner (Leyton et al., 1992). These results indicate that ~ 9 5 is ~a regulator ’ ~ ~of gamete interaction. In vivo labeling of sea urchin eggs revealed that fertilization results in an eightfold increase in 32Porthophosphate incorporation into tyrosine residues of egg proteins because of an increase in the activity of plasma membrane-bound tyrosine protein kinases (Ribot et al., 1984). Tyrosine phosphorylation is therefore believed to be an important step in the events following fertilization.

IX. SUMMARY AND FUTURE DIRECTIONS Fertilization marks both the conclusion of the events of gametogenesis, with its highly specialized cell cycle control points, as well as the commencement of the cell cycles that will characterize the developing embryo. This switch from the germ cell to somatic cell program no doubt also involves changes in phosphorylation,but very little is known about the specific players in this process, especially in mammals. The identification of the responsible genes in more experimentally amenable vertebrates such as Xenopus can help in this regard; for example, the studies on Mos function in frog oocytes yielded much insight into the role this gene plays in gametogenesis and meiosis. However, given the subtle but distinct differences among species in the timing of progression of germ cell differentiation,the requirements for new protein synthesis, etc., as well as the major differences between spermatogenesis and oogenesis in all species, precise function of any specific gene will always remain to be tested in vivo in mammals, and in both the male and female. The power of the transgenic and gene targeting approaches, now almost routine in the mouse, suggest that it is this particular mammalian model system that will lead the way in our understanding of gametogenesisregulating genes, whether kinases or others.

ACKNOWLEDGMENTS This work was supported in part by NIH grant P50 HD05077 and NM training grant GM 07088-18.

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Thomas, L., Clarke, P.L., Pagano, M., and Gruenberg, J. (1992). Inhibition of membrane fusion in virro via cyclin B but not cyclin A. J. Biol. Chem. 267: 6183-6187. Tsai, L-H., Harlow, E., and Meyerson, M. (1991). Isolation of the human cdk2 gene that encodes the cyclin A- and adenovirus E1A-associated p33 kinase. Nature 353: 174-1 77. Ullrich, A., and Schlessinger, J. (1990). Signal transduction by receptors with tyrosine kinase activity. Cell 61: 203-212. Wadewitz, A. G., Winer, M. A., and Wolgemuth, D. J. (1993). Developmental and cell-lineage specificity of raffamily gene expression in mouse testis. Oncogene 3: 1055-1062. Wang, J., and Baltimore, D. (1983). Cellular RNA homologous to the Abelson murine leukemia virus transforming gene: expression and relationship to the viral sequence. Mol. Cell. Biol. 3: 773-779. Wassarman, P. M., Josefowicz, W. J., and Letourneau, G. E. (1976). Meiotic maturation of mouse oocytes in vitro: inhibition of maturation at specific stages of nuclear progression. J. Cell Sci. 22: 531-545. Wasserman, W. J., and Smith, L. D. (1978). The cyclic behavior of a cytoplasmic factor controlling nuclear membrane breakdown. J. Cell Biol. 78: R15-R22. Wasserman, W. J., Richter, J. D., and Smith, L. D. (1982). Protein synthesis during maturation promoting factor- and progesterone-induced maturation in Xenopus oocytes. Dev. Biol. 89: 152-158. Watanabe, N., Vande Woude, G. F., Ikawa, Y., and Sagata, N. (1989). Specific proteolysis of the c-mos proto-oncogene product by calpain on fertilization of Xenopus eggs. Nature 342: 505-5 11. Whitfield, W. G. F., Gonzalez, C., Sanchez-Herrero, E., and Glover, D. M. (1989). Transcripts of one of two Drosophila cyclin genes become localized in pole cells during embryogenesis. Nature 338 337-340. Whitfield, W. G. F., Gonzales, C., Maldonado-Codina, G., and Glover, D. M. (1990). The A- and B-type cyclins of Drosophila are accumulated and destroyed in temporally distinct events that define separable phases of the G2-M transission. EMBO J. 9: 2563-2572. Williams, D. E., Eisenman, J., Baird, A., Rauch, C., Van Ness, K., March, C. J., Park, L. S., Martin, U., Mochizuki, D. Y., Boswell, H. S., Burgess, G. S., Cosman, D., and Lyman, S. D. (1990). Identification of a ligand for the c-kit proto-oncogene. Cell 63: 167-174. Winer, M. A., and Wolgemuth,D. J. (1993). Patterns of expression and potential functions of proto-oncogenes during mammalian spermatogenesis. In: The Molecular Biology of the Male Reproductive System (De Kretser, D. M., ed.), pp. 143-179. Academic Press, San Diego. Winer, M. A., Wadewitz, A. G., and Wolgemuth, D. J. (1993). Members of the rafgene family exhibit segment-specific patterns of expression in mouse epididymis. Mol. Reprod. Dev. 35: 16-23. Witte, 0.N. (1990). Steel locus defines new multipotent growth factor. Cell 63: 5-6. Wolfes, H., Kogawa, K., Millette, C. F., and Cooper, G. M. (1989). Specific expression of nuclear proto-oncogenes before entry into meiotic prophase of spermatogenesis. Science 245: 740-743.

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REGULATION OF THE DOPA DECARBOXYLASE GENE DURING DROSOPHlLA DEVELOPMENT

Martha j. Lundell and jay Hirsh

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Distribution of DDC Protein and Monoamines in the Drosophilu CNS . . . 111. Transcriptional Regulation of the Drosophila Ddc Gene . . . . . . . . . . A. cis-Regulatory Elements in the Ddc Promoter . . . . . . . . . . . . . . B. trans-Acting FactorsThatMay Regulate DdcExpression . . . . . . . rv. Post-Transcriptional Regulation of the Drosophilu Ddc Gene . . . . . . . . V. Comparison of the Drosophilu Ddc Gene to the Vertebrate AADC Gene . . VI. Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Advances in Developmental Biochemistry Volume 3, pages 55-86. Copyright 0 1994 by JAI Press Inc. All rights of reproductionin any form reserved. ISBN: 1-55938-865-X

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PREFACE Dopamine and serotonin are biogenic amines found in multiple tissues of both invertebrates and vertebrates. In the central nervous system (CNS), they act as neurotransmitters, functioning in learning and memory pathways and having physiologic importance in several human diseases. The final step in the biosynthesis of dopamine and serotonin is mediated by a common enzyme, DOPAdecarboxylase (DDC). The gene encoding Drosophila DDC (Ddc) is one of the best characterized eukaryotic genes for tissue- and cell-specific regulation. Several cistranscriptional regulatory elements have been identified within the promoter of Ddc. Some of these elements regulate expression in nonneural tissue, some regulate expression in the entire CNS, and some are cell-specific elements that regulate expression in discrete neuronal subsets within the CNS. Protein factors that bind to some of these regulatory elements have been isolated. These factors are intriguing proteins, in that their structures and temporal patterns of expression implicate them in significant aspects of Drosophilu development. In addition to transcriptional regulation Ddc is also regulated post-transcriptionally by the alternative splicing of specific neural and non-neural mRNAs. The synthesis of distinct mRNAs for neural and non-neural tissue is a common theme for CNS genes expressed in additional tissues. It has recently been shown that the vertebrate Ddc homologue, AADC, also synthesizes two tissue-specific mRNA isoforms. Here we review the regulation of Drosophilu Ddc and compare it to the vertebrate AADC gene.

1. INTRODUCTION Biogenic amines are decarboxylated derivatives of tyrosine and tryptophan that are found in animals from simple invertebrates to mammals. These compounds are found in neural tissue, where they function as neurotransmitters, and in non-neural tissues, where they have a variety of functions. The enzymes involved in biogenic amine synthesis and many receptors for these compounds have been isolated from both invertebrate and vertebrate sources. In all cases, the individual proteins that effect biogenic amine metabolism and function show striking similarity between species, indicating that these are ancient and wellconserved pathways.

Regulation of the DOPA Decarboxylase Gene

0

NH2 -CH

-COOH

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I

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___)

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Figure 7. Biosynthetic pathways for biogenic amines. In Drosophila and vertebrates decarboxylation of DOPA and 5-hydroxy-tryptophan is catalyzed by the same enzyme, DDC. In vertebrates this enzyme is called amino acid decarboxylase (AADC). Only vertebrates further metabolize dopamine to norepinephrine and epinephrine. TH, tryosine hydroxylase; DDC, DOPA decarboxylase; DBH, dopamine b-hydroxylase; PNMT, phenylethanolamine N-methyltransferase.Tryp-OH; tryptophan hydroxylase.

A key enzyme in biogenic amine biosynthesis is DOPA decarboxylase (DDC), also known as amino acid decarboxylase(AADC) in vertebrates. As shown in Figure 1 this enzyme decarboxylates both DOPA and 5-hydroxytryptophan,to form dopamine and 5-hydroxytryptamine(serotonin), respectively. The regulation of the gene encoding DDC has been studied extensively in the fruit fly, Drosophila melanoguster. Functional cis-regulatory elements can be conserved between homologous genes of DrosophiZu and humans (Malicki et al., 1992). It is thus likely that much of the information regarding regulatory elements and factors gleaned from studies in Drosophila Ddc will be relevant to the regulation of vertebrate AADC. In this chapter, we will review recent results concerning the transcriptionaland post-transcriptionalregulation of Drosophilu Ddc. We will also discuss the structure and regulation of the vertebrate AADC gene relative to Drosophila Ddc.

MARTHA J. LUNDELL and JAY HIRSH

58 Embryonic Hatching

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Figure 2. Temporal profile of DDC enzyme activity during Drosophila development. Note the major peaks of DDC induction during lateembryogenesis, at pupariation, and at the time of eclosion of the adult from the pupal case. At these developmentaltimes when there is extensive synthesis and hardening of cuticle, the induced DDC in the hypoderm is involved in this process. Levels of DDC in the CNS show much less developmental variation (Hirsh, 1986). Figure adapted from Hirsh, 1986.

Drosophila Ddc is expressed primarily in the CNS and the hypoderm, the epithelial layer of the fly that secretes the cuticle. In the CNS, Ddc is expressed in a small subset of neurons that produce either dopamine or serotonin (Budnik and White, 1988; Valles and White, 1988). In the hypoderm, Ddc expression leads to synthesis of dopamine, which is further metabolized into quinones that have a vital function in the cross-linking, hardening, and pigmentation of the fly cuticle (Wright, 1987). The developmental profile of DDC activity in these two tissues is quite different (Hirsh, 1986).DDC is first detected during late embryo-

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genesis, between 16 and 20 hours after fertilization,both in the hypoderm and in the CNS (Fig. 2). DDC activity is induced to high levels in the hypoderm at two subsequent developmental stages, pupariation and eclosion. At both of these stages there is extensive cuticular synthesis and hardening. In contrast, DDC activity in the CNS remains relatively constant throughout post-embryonic development and accounts for only a small fraction of total DDC activity. However, the relative level of DDC activity per cell in the CNS is actually quite high because Ddc is expressed in only about 150 neurons within the CNS (Beall and Hirsh, 1987; Konrad and Marsh, 1987) but is expressed in most if not all hypodermal cells (Bray and Kafatos, 1991). Ddc is also expressed in a subset of glial cells (Beall and Hirsh, 1987) and in a portion of the gut, the proventriculus (Konrad and Marsh, 1987), but the metabolic functions in these sites of expression are unknown. Flies carrying Ddc null mutations die during late embryogenesis because of incomplete hardening of the mouth parts, resulting in an inability of the larvae to penetrate the eggshell. A small fraction of these embryos can be rescued by manually ripping the eggshell (Valles and White, 1986). A minority of these larvae will survive through larval development, but they invariably die when they attempt the pupal molt. These larvae lack all detectable DDC and serotonin in their CNS, demonstrating that serotonin is not absolutely essential for larval viability and development. Dopamine is presumably also absent from these larvae, although this has not been assayed directly. Valles and White (1986) have shown that serotonin neurons are present in these larvae, since incubation of a serotonin-deficient CNS in serotonin results in uptake into the normal serotonin neurons. This demonstrates that development of the serotonin neurons does not depend on serotonin, although serotonin could well affect the development of other neuronal subsets. These observations leave unanswered the physiological roles of dopamine and serotonin in the fly CNS. Vertebrates also show expression of AADC in both neural and nonneural tissues. AADC has been purified from kidney (Christenson et al., 1972), liver (Ando-Yamamoto et al., 1987), adrenal medulla (Albert et al., 1987), and pheochromocytoma (Coge et al., 1989; Ichinose et al., 1989). In the adrenal medulla dopamine is further processed into epinephrine and norepinephrine, which are released from the chromaffin cells during stress to increase heart rate and blood pressure. There are no detectable monoamines in the liver and kidney, and the function of AADC in these tissues is unknown. AADC activity has also been

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reported in a variety of other tissues (Lovenberg et al., 1962; Rahman et al., 1981). High levels of AADC and monoamines are found in two tumor cell types, pheochromocytoma and small cell carcinomas of the lung (Baylin et al., 1980; Nagatsu et al., 1985). As in Drosophilu, only a small number of cells in discrete regions of the vertebrate CNS synthesizemonoamines (Jaeger et al., 1984). In both Drosophilu and vertebrates this small population of neurons branches extensively to encompass large areas. The multiple roles of biogenic amines in the CNS are accomplished by releasing the neurotransmitters over large regions of the CNS. Specific physiological effects are established by receptors that are expressed in defined post-synaptic patterns. In Drosophila three serotonin receptors have been identified. They are expressed in specific cellular subsets, and all three are G protein-coupled trans-membrane receptors (Witz et al., 1990; Saudou et al., 1992). In vertebrates multiple receptors for serotonin, dopamine, and norepinephrine have been identified (reviewed in Kandel et al., 1991). All are G protein-coupled receptors, except for one that is linked to a ligand gated ion channel (Maricq et al., 1991). In the vertebrate CNS monoamines have been associated with a number of physiological functions (reviewed in Kandel et al., 1991). Serotonin has functions associated with mood, pain, sleep, learning, and memory. Dopamine has functions associated with schizophrenia, Parkinson’s disease, and cocaine addiction. In vertebrates, dopamine is further metabolized into two additional neurotransmitters, norepinephrine and epinephrine. Norepinephrine increases the excitability of cells in response to sudden sensory input such as fear. Epinephrine has been identified in specific neurons of the brain, but the function of these cells is unknown. In addition, AADC has also been found in a class of neurons that do not have any of the four neurotransmitters discussed above (Jaeger et al., 1983). These neurons may use one of the “trace amines,” tyramine, tryptamine, or phenylethylamine, as a neurotransmitter. There has been one report of a congenital deficiency for AADC in humans. A set of twins has been found with AADC activity levels that are 1% of control values (Hyland and Clayton, 1990). Aggressive treatment of these twins was initiated with multiple drugs aimed at stimulating dopamine receptors and slowing the degradation of any residual amines. As of the time of this clinical report, the twins were surviving at 17 months but suffering from multiple neurological defects, whereas a previous untreated sibling had died.

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II. DISTRIBUTIONOF DDC PROTEIN AND MONOAMINES IN THE DROSOPHILA CNS As mentioned above, Ddc is expressed in only about 150cells within the Drosophila larval CNS. Figure 3 shows a cartoon illustratingthese cells,

VL Figure 3. Sketch of DDC-expressing neurons in the Drosophila larval CNS. The CNS consists of brain lobes and a segmented ventral ganglion. Filled circles represent dopamine cells; open circles represent serotonin cells; grayed circles represent DDC cells that contain no detectable tyrosine hydroxylase or serotonin immunoreactivity, indicating that these cells may produce neither transmitter(Lundel1and Hirsh, 1994). M, medial dopamine neurons; VL, ventrolateral serotonin neurons; DL, dorsolateral dopamine neurons. The hatched rectangle shows the region of the ventral ganglion that i s shown in Figures 4 and 6.

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and Figure 4 shows a panel of projections illustrating DDC cells in the ventral ganglion, generated using a confocal laser scanning microscope. Three reagents are available that facilitate examination of the cellular pattern of Ddc expression. Antisera specific for Drosophila DDC allow detection of all DDC-expressing cells (Beall and Hirsh, 1987; Konrad

Figure4. DDC (A), serotonin (B), and tyrosine hydroxylase (C) immunoreactivity in the posterior region of a wild-type Drosophila ventral ganglion. Tyrosine hydroxylase (TH) encodes the rate-limiting step in dopamine biosynthesisand is a marker for dopamine cells. B and C are the same CNS assayed for both serotonin and TH. M, medial dopamine neurons; VL, ventrolateral serotonin neurons; DL, dorsolateral dopamine neurons. Short unmarked arrows in C show vacuolated cells that do not contain DDC immunoreactivity. The immunoreactivity in these cells may represent a nonspecific cross-reactivity of the rat TH antibody. The length bar in A is 50 pM. The images are confocal projections generated on a Molecular Dynamics-2000 confocal laser scanning microscope.

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and Marsh, 1987; Scholnick et al., 1991). Antisera specific for serotonin are in common use, and a commercially available antiserum made against the rat tyrosine hydroxylase (TH) is used to detect dopamine-producing cells. This enzyme performs the rate-limiting step in dopamine biosynthesis (Figure 1) and is well conserved between vertebrates and Drumphila (Neckameyer and Quinn, 1989). TH-expressing cells show good correspondence to those that react with catecholaminespecific reagents and dopamine-specific antiserum (Budnik and White, 1988; Lundell and Hirsh, 1994). DDC cells are either serotonin cells (open circles in Figure 3) or dopamine cells (closed circles). No cells have been detected that synthesize both transmitters. However, there is a set of cells in the subesophageal region of the ventral ganglion that express high levels of DDC but contain neither serotonin nor TH immunoreactivity (grayed cells in Figure 3) (Lundell and Hirsh, 1994). As mentioned previously, DDC cells that contain neither catecholamines nor serotonin are also found in the mammalian brain (Jaeger et al., 1983). The majority of DDC-expressing cells in the brain lobes are dopamine cells. Most of these dopamine cells have axons that project into a common axonal fiber extending anteriomedially within the brain lobe and then separating into finer fibers that cross between the lobes. The dopamine cells occur in small clusters of two to six cells, which suggests that these cells might share common lineages. The serotonin cells within the lobes are also found in pairs, and each pair projects axons into closely associated tracts. The pathwaysof the serotonintracts often parallel those of the dopamine cells but are distinct (Lundell and Hirsh, 1994). Figure 4 shows confocal images of the staining pattern for DDC (Fig. 4A), serotonin (Fig. 4B), and TH (Fig. 4C) in the segmental ventral ganglion of the CNS from third instar larvae. Panels B and C are the same CNS double stained with serotonin and TH. The DDC-expressing cells can be categorized into a set of paired ventral lateral serotonin cells (Fig. 4A,B; labeled VL in 4A), and two morphologically distinct types of dopamine cells, the medial dopamine cells (Fig. 4A,C; labeled M) and the dorsal-lateral dopamine cells (Fig. 4A,C; labeled DL). Figure 4 demonstrates clearly that individual DDC cells synthesize either serotonin or dopamine, but not both. One additional set of cells shows TH immunoreactivity in the ventral ganglion. These six large vacuolated cells are located more laterally than any other DDC cells (Fig. 4C, unlabeled short arrows). It is likely that the immunoreactivity in these cells results from a non-specific cross-reaction, since these cells are not

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detected with DDC (Fig. 4A) and have not been detected previously with catecholamine-specificreagents (Budnik and White, 1988). The axonal projections from the DDC-expressing cells in the ventral ganglion also show tendencies to follow common pathways. The projections from the ventral lateral serotonin cells extend medially to fuse with axons projecting from the contralateral serotonin cells. At the midline, this projection is met by an axonal projection from the medial dopamine cell.

111. TRANSCRIPTIONAL REGULATION OF THE DROSOPHILA Ddc GENE A. cis-Regulatory Elements in the Ddc Promoter The cell-specific expression of Ddc in the CNS is generated by a set of interacting transcriptional regulatory elements. Both tissue-specific elements, required for Ddc expression in the whole of the CNS, and cell-type-specific elements have been identified in the Ddc promoter. A summary of these elements is shown in Figure 5 . Study of the Ddc regulatory elements relies on the ability to integrate in v i m altered Ddc genes into the Drosophila germline using P-element transposable vectors. These vectors are integrated into a genetic background that contains a temperature-sensitiveDdc allele, DdP2. At 25 C, Ddcts2synthesizes a very low level of DDC protein, making it a suitable host for examining DDC expression from altered Ddc genes by indirect immunofluorescence (Beall and Hirsh, 1987; Konrad and Marsh, 1987). Furthermore, the low levels of DDC activity in the hypoderm of Ddcrs2result in altered cuticular pigmentation, making it easy to select for transformed flies that express Ddc at higher levels (Scholnick et al., 1983). Transformation of Ddc‘” with a Ddc gene that extends 1600 bp upstream from the transcriptional startpoint is sufficient for normal Ddc expression in both the CNS and hypoderm (Johnson et al., 1989; Lundell and Hirsh, 1992). A minigene that retains only 208 bp proximal to the transcriptional start point restores wild-type expression of DDC in the hypoderm (Hirsh et al., 1986), yet leads to inappropriate cell-specific expression within the CNS (Beall and Hirsh, 1987). Additional experiments have determined that Ddc regulatory elements are clustered in two regions, a proximal regulatory region located within 120 bp of the transcriptional start site, and a distal region located from -1000 to -1 600

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bp upstream of the start site (Fig. SB,C). The proximal region contains elements required for the tissue-specific expression of Ddc in the hypoderm and the CNS (Scholnick et al., 1986; Bray et al., 1988), and specific elements that regulate Ddc expression in glial cells (Mastick and Scholnick, 1992). The distal region contains most if not all of the elements regulating the neuronal-specific pattern of Ddc expression in the CNS as well as some tissue-specific CNS elements (Johnson et a]., 1989; Lundell and Hirsh, 1992). The distal region has enhancer properties in that it can function independently of position and orientation (Johnson et al., 1989). Normal Ddc expression in the CNS requires both the proximal regulatory region and the distal enhancer functioning in consort. More precise localization of regulatory elements within these regions has been accomplished using fine structure deletions and point mutant alterations (Bray et al., 1989; Johnson et al., 1989; Lundell and Hirsh, 1992). These studies have been guided by DNA sequence comparisons with the Ddc gene isolated from Drosophila virilis, which diverged from D. melanogasterS0-70 million years ago. This period has been sufficient to allow divergence of most nonessential DNA regions, whereas many regions with regulatory importance have been conserved (Bray and Hirsh, 1986; Johnson et al., 1989). Furthermore, there is sufficient conservation of regulatory elements that the D.virilis gene functions normally during development when integrated into the Ddc‘*-’mutation of D. melanogaster (Bray and Hirsh, 1986). Within the proximal promoter are found five sequence elements that are conserved between D. melanogaster and D. virilis (Scholnick et al., 1986) (Fig. 5). The 16-bp element I sequence is perfectly conserved between D. melanogaster and D. virilis (Bray and Hirsh, 1986). Inactivation of element I by deletion or point mutation leads to a selective loss of Ddc expression within the CNS (Scholnick et al., 1986; Bray et al., 1988) (Fig. 6A), without significantly affecting hypodermal Ddc expression. This demonstrates that element I is required specifically for expression of Ddc within the CNS. A repressing function for Ddc glial expression has been localized within another ofthe conserved elements in the proximal region (Mastick and Scholnick, 1992). Loss of this element, either by deletion or by point mutation, results in DDC overexpression in glial cells. Expression of DDC in glial cells of wild-type flies is almost undetectable, and these glial cells show no expression of dopamine or serotonin even in strains that show high levels of glial DDC expression (unpublished results).

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The distal enhancer also contains five sequence elements that are conserved between D. melunogaster and D. virilis (Johnson et al., 1989; Lundell and Hirsh, 1992) (Fig. 5). Both tissue-specific and cell-specific regulatory functions have been attributedto these elements. Here we will

A) Ddc

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figure 5. Transcriptional regulatory elements of Ddc. A. The Ddc exons and 2200 bp of 5' flanking sequences that are sufficient for normal Ddc expression. B. An enlargement of the proximal regulatory region. Striped boxes represent elements conserved between Drosophila melanogaster and Drosophila virilis. The largest element extending from -82 to -95 is actually two elements. Binding sites for potential regulatory factors are indicated. The X over Elf1 indicates that this factor is not likely to be the functional factor that binds to this site in vivo (see text). C.An enlargement of the distal enhancer.

Figure 6. In vivoexpression of DDC in wild-type and mutant Ddcstrains, showing the specificity of CNS transcriptional regulatory elements. WT. DDC staining in a wild-type ventral ganglion. A. contains a proximal regulatory region deletion that includes element 1 . It shows severely reduced DDC expression in all neurons that normally express DDC. B. D ~ c "contains ~ a clustered point mutation in element C of the distal enhancer and shows reduced DDC expression in the medial dopamine neurons, but normal levels of DDC in the serotonin neurons. C. Ddc'03 contains a deletion of the SER regulatory element in the distal enhancer. It shows reduced DDC expression in the serotonin neurons but normal levels in the medial dopamine neurons. These images are confocal projections that do not include the dorsolateral cells. Expression of DDC in the dorsolateral cells is reduced in D ~ c ~ but ~ . is" normal in D ~ cand ' ~ ~ Ddc7O3.The length bar in WT is 50 pM. 67

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limit our discussion to two regions that contain cell-type-specific elements, element C and SER. Loss of a functional element C, either by deletion or by point mutation (Johnson and Hirsh, 1990), results in a selective loss of DDC immunoreactivity in the midline dopamine cells of the ventral ganglion (Fig. 6B). This element is specific for this class of dopamine cells, since expression in the dorsal lateral dopamine neurons is unaffected. The second cell-specific regulatory element within the distal enhancer is named SER. Loss of this element, which is at the extreme distal edge of the enhancer, leads to a selective loss of DDC expression in the ventral lateral serotonin cells (Johnson et al., 1989; Lundell and Hirsh, 1992) (Fig. 6C). This element has been delimited to about 40 bp and shows unexpected complexity, consisting of two functionally redundant elements. These two subelements, SERLand SERR,are each sufficient to allow normal DDC expression in the ventral lateral serotonin neurons if the other is deleted. In spite of this functional similarity, no sequence similarity is apparent between the two regions. The region of conservation between D.melunoguster and D.virilis is limited to SERL. B. fransActing Factors That May Regulate Ddc Expression

Many of the cis-elements mentioned above bind proteins in nuclear extracts prepared from Drosophilu embryos. Several potential Ddc regulatory factors have been identified and cloned by screening phage expression libraries with radioactively labeled concatamers of these elements. Although DNA binding factors are straightforward to isolate, it is much more difficult to prove in vivo function. The complication is that many proteins can bind to multiple sequences, and many sequences can bind multiple proteins. The conclusive determination of in vivo function can only come from a combination of genetic and reverse genetic techniques (ie., examining the loss of function phenotype) and by examining gain of function phenotypes by inappropriately overexpressing the candidate regulatory product. These studies are feasible in Drosophilu, and the potential to perform these studies remains one of the strengths of Drosophilu as a model system. We will discuss four of these Ddc regulatory element binding factors, surveying the progress that has been made in determining their in vivo roles. Figure 5 shows the four factors, Elf 1 , Cf 1a, ZFH-2, and Adf 1 , and the regulatory sites to which they bind.

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figure 7. Homology between ZFH-2 and ATBFI. A. Homeodomains (rectangles)and zinc fingers (circles) of ZFH-2 and ATBFI. The nine most significant identities between the proteins are represented by the lines connecting the motifs, with the numbers on the lines indicating the percent identity. Other intergenic and intragenic identities between the zinc fingers and homeodomains were detected in this search, with identities between 30% and 40%. The shaded homeodomain represents the conserved ”pseudohomeodomain” I in each gene. 5. Sequence comparison of the conserved “pseudohomeodomain” I. The canonical homeodomain sequence is shown below the ATBFI and ZFH-2 homeodomain sequences. Identities are marked with I; conservative substitutions are marked with *. The four shaded amino acids represent significant changes from the canonical sequence, which are discussed in the text. Note especially that the ATBFI and ZFH-2 homeodomains share identical sequences in helix I l l and extending further to the right. Reproduced with permission from Lundell and Hirsh (1992). The factor Elfl was isolated as a factor that binds to the CNS-specific element I in the proximal regulatory region (Bray et al., 1988). Several lines of evidence pointed to Elfl as a strong candidate for a specific Ddc regulatory factor (Bray et al., 1989): First, mutations of element I that disrupt function also reduce the affinity of Elfl. Second, Elfl is the only element I binding activity detectable in an embryonic nuclear extract,

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and an antibody to Elfl can deplete the extract of element I binding activity. Third, in v i m studies on Elfl, also known as NTFl, show that it can activate transcription from the Ddc promoter (Dynlacht et al., 1989). Elfl immunoreactivity is localized in the nuclei of most epidermal cells and in specific neurons of the CNS. However, Elfl immunoreactivity is not found in Ddc-expressing cells in the CNS (Bray et al., 1989). Although the binding experiments are consistent with a role for elf1 in regulating Ddc, the failure to find Elfl co-localized in Ddc neurons is not consistent with simple regulatory models. A genetic analysis of Elfl has lead to the isolation of recessive lethal mutations within the gene encoding Elfl (Bray and Kafatos, 1991). This study shows that the gene encoding Elfl corresponds to a previously identified mutation, grainyhead. The grainyhead gene (now called EFI/grh) is a late embryonic lethal with defects in the larval head cuticle. A direct assay of the effects of a lethal Elf--l/grh allele on Ddc showsjust the opposite of what was expected: EZf--I/grhaffects Ddc expression in the hypoderm, but not in the CNS. Therefore, some factor other than Elf-1 must bind to element I to regulate CNS expression of Ddc. These results also indicate that Elf-1 may be interacting with hypodermal regulatory elements of Ddc; however, this interaction could be indirect, with Elf- 1 activating other transcription factors that could bind directly to Ddc. These experiments demonstratethe paramount value of a genetic analysis to assess the in vivo function of factors that have binding and transcriptional stimulatory activities in vitro. The factor Cfla was isolated as a factor that binds to the dopaminespecific regulatory element C in the distal enhancer (Johnson et al., 1989). As in the case with Elf-1, Cfla binds preferentially to a wild-type binding sequence versus a mutated binding sequence that lacks function in vivo. DNA sequence analysis shows Cfla to be a member of the POU-domain family of homeodomain transcription factors. These factors are characterized by a homeodomain as well as a second conserved region, the POU-specific domain. This is an interesting family of transcriptional regulatory proteins in that many other POU proteins have neural functions: The dwarfmutation in mouse results from a mutation in the POU protein Pit-1 (Li et al., 1990), required for development of three pituitary cell types, and the C. elegans POU protein Unc-86 is important for the proper differentiation of serotonin and dopamine neuronal lineages (Desai et al., 1988). In addition, the vertebrate POU proteins bml, brn2, bm3, and tstl, which are most similar to Cfla, are expressed in the developing vertebrate brain (He et al., 1989).

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Cfla binds DNA as a homodimer. In vitro binding experiments have shown that another DrosuphiZa POU protein, I-POU, can form heterodimers with Cfla (Treacy et al., 1991). I-POU can act as a negative regulator of Cfla DNA binding activity, analogous to the negatively acting proteins that can form heterodimers with bHLH transcription factors (Benezra et al., 1990; Ellis et al., 1990; Garrell and Modolell, 1990). Further evidence regarding the in vivo role of I-POU and the role of Cfl a as a potential regulator of Ddc awaits analyses of these genes. The factor ZFH-2 was identified by its binding to the serotonin cell specific element SERL in the Ddc distal enhancer (Lundell and Hirsh, 1992). The predicted ZFH-2 translation product encodes a large factor of M W >300 kDa, which contains 3 homeodomains and 16 zinc finger motifs (Figure 7). This protein was initially identified as a factor that binds to a conserved element in promoters of the Drosophila opsin genes (Fortini et al., 1991; Lai et al., 1991). Homeodomains and zinc fingers are DNA binding motifs found in numerous transcription factors, but the finding of multiple homeodomains and zinc fingers within one factor is unprecedented. At least two of the homeodomains are active in DNA binding and show differential binding specificity. Homeodomain I11 binds preferentially to the opsin element, whereas homeodomain I1binds the Ddc element preferentially (Lundell and Hirsh, 1992). Homeodomain I is unusual, having been termed a “pseudohomeodomain” by Fortini et al. (1991). It shows a very unusual amino acid composition in the portion of the homeodomain that normally is in direct contact with DNA, such that this motif may not be active in DNA binding. A human homologue of zfh2 has been identified that gives some clues to important motifs within ZFH-2. This gene, ATBF1, which binds to the promoter of the alpha-fetoprotein gene, encodes a protein with four homeodomains and 17 zinc fingers (Morinaga et al., 1991). The nine most similar regions between ZFH-2 and ATBFl are the three homeodomains and six of the zinc finger motifs. Figure 7 shows that these top nine identities are collinear between the two genes (Hashimoto et al., 1992; Lundell and Hirsh, 1992). This indicates that there has been evolutionary pressure to maintain all these motifs within a single protein and with the same linear order. This evolutionary pressure would occur if the motifs within each protein function coordinately rather than independentlyat their targets. It is interestingthat the most similar region between these two genes is homeodomain I, the “pseudohomeodomain,” which shows 78% identity. Thus, instead of being an evolutionary relic,

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this domain must have an important role within the proteins, even if its function is other than DNA binding. The expression profile of ZFH-2 gives clues to its in vivo role. The expression of ZFH-2 is limited almost exclusively to the CNS, starting during mid-embryogenesis and continuing into late larval development (Lai et al., 1991; Lundell and Hirsh, 1992). By late larval development ZFH-2 is expressed in about 20% of CNS neurons. If ZFH-2 has a role in regulating Ddc expression in the serotonin cells of the ventral ganglion, then ZFH-2 should be expressed within these neurons. This has been examined using indirectly labeled fluorescent antibodies and confocal microscopy to resolve colocalization of DDC and ZFH-2 in individual cells of the CNS. The answer is complex but interesting; ZFH-2 immunoreactivity is found in only the outside cell of each pair of serotonin neurons in the ventral ganglion, and in addition, ZFH-2 immunoreactivity is found in the medial dopamine cells, cells whose Ddc expression is not affected by inactivation of the ZFH-2 binding site (Lundell and Hirsh, 1992) (Fig. 8). The paired serotonin cells in the ventral ganglion have been shown to be mitotic sisters in grasshopper (Taghert and Goodman, 1984). Given the similarities between grasshopper and Drosophilu in this portion of the nervous system (Doe, 1992), we suspect that the same will hold for Drosophila. The finding of ZFH-2 in only one cell of each serotonin cell pair shows that these cells have differentiatedproperties not evident from morphological examination. The expression of ZFH-2 in only one serotonin cell does not correlate directly with the reduction of DDC expression in both cells when the ZFH-2 binding site is altered. If ZFH-2 is regulating Ddc in these cells, then there must be some subtleties in the regulation. One possibility is that ZFH-2 could act early in development in the progenitor cell of the paired serotonin cells. There are other possibilities as well, but a definitive answer will come from the genetic analysis of ~$22. zfh2 is located on the fourth chromosome (Fortini et al., 1991). Several marked P elements that are closely linked to zjh2 are facilitating this genetic analysis (Lundell and Hirsh, manuscript in preparation). The protein Adfl was initially isolated as a factor binding to the promoter of the alcohol dehydrogenase gene (England et al., 1990). It was also shown to bind to a site within the Ddc proximal regulatory region, which is important for repressing Ddc expression in glial cells (Mastick and Scholnick, 1992). Adfl contains a helix-turn-helix DNA binding domain that is related to the Myb family of proteins (England et al., 1992). There is currently no additional evidence to support an in vivo role for Adfl in Ddc regulation.

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Figure 8. Colocalization of DDC and ZFH-2 in larval CNS. This figure shows abdominal segments 4-7 of a third instar larval CNS. DDC immunoreactivity is cytoplasmic and is shown in red, whereas ZFH-2 immunoreactivity is nuclear and is shown in green. Only the outside cell of each pair of serotonin neurons expresses ZFH-2. One medial dopamine cell shown at the top also shows ZFH-2 expression. This projection does not include the dorsolateral cells, which also express ZFH-2. A similar projection has been published in Lundell and Hirsh (1 992).

IV. POST-TRANSCRIPTIONALREGULATION OF THE DROSOPHILA Ddc GENE The cis-regulatory elements described above and the factors that bind to these elements determine the spatial and temporal expression of the Ddc primary transcript. Like a number of other genes that are expressed in

MARTHA J. LUNDELL and JAY HlRSH

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Altered Ddc Genes

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Figure 9. Summary of Ddc mRNA splicing and protein products in the CNS and hypoderm of flies transformed with altered Ddc genes. Altered Ddc genes are shown on the left. The spliced mRNA products generated in the CNS (upper line) and hypoderm (lower line) are shown on the right. Arrows on each mRNA indicate sites of translational initiation. 1) Wild-type Ddc gene. All four exons are included in the CNS mRNA, whereas exon B is excluded from the hypodermal mRNA. The open reading frame of the CNS mRNA starts in exon 6, then fuses into the same reading frame as the hypodermal mRNA. This results in a 35 amino acid N-terminal extension on the CNS protein relative to the hypodermal protein. 2) Hsp/Ddc fuses the Hsp70 promoter to exon A. 3) DdCBAdeletes the DNA between exon A and C, fusing these exons. 4) DdPb' deletes the intron between exons B and C. 5) Dd?ORFcontains a frameshift mutation in exon B. The hatched line indicates a frameshift that results in a missense protein produced exclusively in the CNS.

both neural and non-neuronal tissues, lldc is also regulated at the level of RNA processing to produce different mRNAs in each tissue type (Morgan et al., 1986). The unique 4.0-kb Ddc primary transcript is alternatively spliced into two mRNA isoforms, a 2.3-kb transcript containing all four exons A, B, C, and D, and a 2.1-kb transcript containing exons A, C, and D (Fig. 9, line 1). The 2.3-kb mFWA is unique to the

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CNS, whereas the 2.1-kb mRNAis found in the hypoderm. The two RNA isoforms lead to distinct DDC protein products. In the hypodermal mRNA, translation begins at an AUG in exon C to produce a 54-kDa protein. In the CNS mRNA translation begins at an AUG in the CNSspecific exon B and continues in the same reading frame as the hypodermal mRNA. This results in a 57-kDa CNS DDC protein with a 35-amino acid amino-terminal extension that is chromatographicallydistinguishable from the hypodermal DDC protein. Ddc expression is limited to 150 neurons of the CNS, whereas most if not all hypodermal cells express Ddc (Bray and Kafatos, 1991). Is the alternative splicing of Ddc in the CNS a characteristic of all the cells in the CNS, or is this capability limited to those cells that normally express Ddc? To address this question a Ddc gene was constructed that expresses the Ddc primary transcript constitutively under the control of an hsp70 heatshock promoter (Hsp/Ddc, Fig. 9, line 2) (Shen et al., 1993). The hsp70 promoter functions at a basal level under non-heatshock conditions to produce Ddc transcripts in all cells of the CNS and of the entire animal. Examination of the HspDdc mRNA products from these animals, by reverse transcription-linked PCR or Northern blot analyses, indicates that the CNS produces exclusively the CNS-specific isoform, whereas the hypoderm produces only the hypoderm-specific isoform. This shows that there is a distinction between neural and non-neural splicing that is characteristic of the entire tissue. These results imply that neural and non-neural tissues contain different splicing factors that may be important determinants of tissue identity. Additional altered Ddc genes can be used to investigatethe physiological roles of the two DDC protein isoforms. A Ddc gene containing a fusion of exon A to exon C can produce only the hypodermal mRNA and the hypodermal protein isoform (Fig. 9, line 3) (Morgan et al., 1986). Ddc null larvae containing this gene, DdPA,synthesize wild-type levels of serotonin in the CNS and mature to fully viable adult flies with wild-type levels of DDC enzyme activity in both the hypoderm and in the CNS (Morgan and Hirsh, unpublished observations).These observations indicate that the DDC hypodermal isoform is fully functional when expressed in the CNS. Acomplementary construct in which exons B and C are fused, Dd&bc (Fig. 9, line 4), produces only the CNS mRNA and the CNS protein isoform in both CNS and hypoderm (Morgan et al., unpublished observations). Ddc null larvae containing this gene show near-wild-typeexpression of DDC enzyme activity in both the hypoderm and CNS and develop into viable adult flies. These observations indicate

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that the two DDC isoforms have indistinguishable functions and can be interchanged between the two tissues. Why Drosophila expresses these two isoforms via alternative splicing remains a mystery. The splicing specificity and translational starts for the two Ddc mRNAs can be used to generate flies that specifically lack DDC expression in the CNS. This is done by transforming flies with an altered Ddc gene that contains a frameshift mutation in exon B (Morgan et al., 1986) (Fig. 9, line 5). This gene, named Dd$ORF, expresses normally in the hypoderm, since exon B is not included in the hypodermal Ddc &A. In the CNS, however, translation initiates at the AUG in exon B and extends into the frameshift mutation, producing a nonfunctional protein. DdP2 flies transformed with Dd2.ORFlack detectable DDC enzyme activity in the CNS. TheDd2.0RFgene provides an initial opportunity to test the physiological role of Ddc in the CNS, since it is specifically deficient in expression within the CNS. To do this the Dd2.0RFgene was transferred from the D d P 2 genetic background, which has low levels of Ddc function but wild-type levels of serotonin, into a Ddc null background totally deficient for Ddc function and serotonin (Morgan and Hirsh, unpublished observations). Larvae from these flies still contain very low levels of serotonin in the CNS, at levels about 5% of wild type. The source of this synthesis is unclear; it may come from uptake of serotonin in the periphery or from low levels of DDC synthesized from translational reinitiation at the downstream ATG in exon C. A small fraction of these larvae survive to adulthood, but these flies are so weak that assays of behavior have not been possible. These flies indicate that Ddc expression in the CNS is a near-vital function. However, they also indicate that more subtle alterations of Ddc expression within the CNS will be required to generate flies that are healthy enough for assessment of behavioral function.

V. COMPARISON OF THE DROSOPHILA Ddc GENE TO THE VERTEBRATE AADC GENE Drosophila DDC belongs to a family of pyridoxal-dependentdecarboxylases that extends from prokaryotes to eukaryotic plants and animals. The members of this family show significant sequence similarity over much of their length, even though the individual proteins have quite different substrate specificities, including the amino acids tyrosine, tryptophan, phenylalanine, histidine, and glutamate, and the amino acid derivatives

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omithine, 5-hydroxytrptophan,and DOPA (deLuca et al., 1989;Jackson, 1990; Kang and Joh, 1990). Figure 10 shows a protein sequence comparison of the two DDC protein isoforms with human AADC and with a plant tryptophan decarboxylase (TDC) protein. The sequence identity between Drusuphilu DDC and the periwinkle TDC protein is -40%, whereas the sequence identity with human AADC is -60%. The regions of greatest similarity are in a region surroundingthe pyridoxal phosphate binding site. The regions of similarity start at the beginning of the hypodermal isoform of DDC. The 35-amino acid amino-terminal extension found on the CNS DDC isoform is not similar to sequences in any of the related decarboxylases. As shown in Figure 10, the TDC protein also contains an amino-terminal extension of about 20 amino acids that is unrelated to the fly CNS isoform. As discussed previously, the two fly DDC isoforms have proved functionally interchangeable in vivu, such that the biological rationale for these two protein isoforms remains a mystery. In spite of the similarity of the gene products, structures of the genes encoding vertebrate AADC and Ddc are quite different: Drusuphila Ddc contains four exons encompassing 4 kb of genomic DNA, whereas the human AADC gene contains 15 exons spread over 85 kb of genomic DNA (Sumi-Ichinose et al., 1992). Similar to Drusuphila Ddc, there are distinct neural and non-neural AADC -As in vertebrates (Krieger et al., 1991). One isoform is found in liver and kidney, and a different isoform is present in brain, adrenal medulla, and pheochromocytoma cells (Ichinose et al., 1992). However, unlike Drusuphila, the two vertebrate AADC mRNAs produce an identical protein product in both tissue types (Albert et al., 1987). In Drusuphilu the two tissue-specific mRNAs are generated by alternative splicing of a single primary transcript (Fig. 9). In vertebrates the two tissue specific AADC transcripts are generated from two alternative promoters (Fig. 11) (Albert et al., 1992; Ichinose et al., 1992; Thai et al., 1993). In neural tissue transcription initiates from exon N1, whereas in non-neural tissue transcription initiates from exon L1. This produces two distinct primary transcripts that are then spliced from the first exon (L1 or N1) to exon 2 to generate two tissue-specific mRNAs. Translation initiates within exon 2, such that the same AADC protein product is synthesized from both AADC mRNAs. Although there is a different pattern of exons spliced in the two AADC mRNAs, this apparent alternative splicing is a consequence of differences in transcription. In the non-neural mRNA that initiates at exon L1,

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Figure 10. Amino acid sequence comparison of the CNS and hypodermal forms of Drosophila DDC with human amino acid decarboxylase (HUMAADC) and periwinkle tryptophan decarboxylase (TDC). Identical amino acids are indicated by dashes, and periods indicate gaps inserted to maximize the alignment. The boxed region shows the pyridoxal phosphate binding site. Asterisks indicate residues that are perfectly conserved among eight pyridoxal decarboxylases (Jackson, 1990). The brackets indicate a region that shows greatest similarity among all pyridoxal decarboxylases (Jackson, 1990; Kang and Joh, 1990). Alignment performed per Altschul et al. (1990). 78

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figure 77. Alternate promoters are used for production of the vertebrate neural and non-neural AADC mRNAs. Non-neural AADC transcription initiates at exon L1, whereas neural transcription initiates at exon N1. The non-neural mRNA splices from exon L1 to 2, since the 5’ edge of exon N1 is a site of transcriptional initiation instead of a splice acceptor site. Translation initiates from the same AUG in exon 2 in both mRNAs, producing the same protein product in both tissue types. This scheme holds for both human and rat AADC, although the nomenclature of the exons differs. In rat AADC the exon N1 to 2 splice uses a splice acceptor site 5 bp downstream of the splice acceptor used for the exon L1 to 2 splice (Albert et at., 1992).

exon N1 is excluded because its 5’ end is a transcriptional start site instead of a splice acceptor site. This is in contrast to Drosophila Ddc, where there is a true choice between exons: in non-neural tissue the donor site of exon A is preferentially spliced to the acceptor site of site of exon C instead of exon B, and the opposite choice is made in the CNS (Fig. 9). The cloning of rat and human AADC is recent enough that no studies have been published reporting the minimal promoter region sufficient for normal tissue- or cell-specificregulation in aliving animal. However, the 560 bp flanking the human AADC neural exon, exon N1, has promoter activity in neuroblastoma cells (Thai et al., 1993).This region does not show promoter activity when introduced into several other cell types, indicating that it may correspond to a neural specific promoter region. This study also found several regions showing weak similarity

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to Drosophilu Ddc cis-regulatory elements in the intronic region between exons L1 and N1, but the significance of these regions remains to be established.

VI. PROSPECTS It is clear that the regulation of the Drosophilu melunoguster Ddc gene is complex. This is not surprising, considering the diverse functions of monoamines in multiple tissues. It is highly unlikely that a complete understanding of the Ddc regulatory apparatus will come from a study of this gene alone. Comparison of D. melunoguster Ddc with D. virilis Ddc has been useful in identifying important regulatory regions that are conserved, whereas unimportant regions show rapid divergence. Further insights may come from looking more closely at the vertebrate AADC gene and performing similar comparisons on the AADC regulatory regions from various eukaryotes. In addition, comparison of Ddc to other genes in the monoamine biosynthetic pathway or genes that have a pattern of expression similar to that of Ddc may detect common regulatory mechanisms that have evolved to achieve overlapping patterns of gene expression. This regulatory information derived from Drosophilu Ddc has a high likelihood of being relevant to the vertebrate AADC gene. It is now clear that genetic regulatory mechanisms and molecules can be conserved to a remarkable degree between vertebrates and flies. Many examples exist of important developmental regulatory molecules that are conserved from flies to humans, where they may perform analogous developmental roles (Wright, 1991;McGinnis and Krumlauf, 1992). Furthermore, specific vertebrate regulatory molecules can functionally substitute for their homologues in flies (McGinnis eta]., 1990), and regulatory regions from vertebrate genes can drive a pattern of expression in flies that resembles the vertebrate pattern of expression (Awgulewitsch and Jacobs, 1992; Malicki et al., 1992). How can this regulatory information be put to use? It should be possible by manipulation of regulatory elements in Ddc and related genes to design flies that have aberrant patterns of cell-specific expression that will show selective loss or gain of monoamines in particular cellular subsets. These flies can be generated by P-element transformation of genes that have altered regulatory regions, or by constructing hybrid genes, taking advantage of regulatory regions with inherently different

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patterns of expression. These flies could be used to address questions regarding the development of monoamine-containing neurons, the biological function of monoamines in the CNS, and what mechanisms exist for regulating specific function. It seems possible that understanding Ddc regulation may be of use in considering therapies for amelioration of at least one human disease, Parkinson’s disease. This disease is characterized by the destruction of cells of the substantia nigra, cells that normally produce dopamine. The standard treatment is to give patients L-DOPA, which is converted to dopamine within the brain by residual AADC (Kang et al., 1992). The degenerative nature of the disease makes the success of this treatment limited. Experiments with fetal transplants have had some success, but this is not a routine approach due to practical and ethical considerations. Recently, attempts have been made to treat this disease by injecting the brain of diseased rodents with immortalized cells that are transfected with the tyrosine hydroxylase (TH) gene. In initial attempts, TH-expressing fibroblasts produce DOPA but not dopamine (Fisher et al., 1991). In this situation, the animal must convert the DOPA to dopamine using residual AADC within the brain. However, recent strategies have used TH-transfected muscle cells that apparently express low levels of AADC (Jiao et al., 1993). These cells synthesize large quantities of dopamine and achieve long-term improvement of Parkinson’s symptoms. Although a portion of the increased efficacy of the muscle cells may be due to lengthened survival within the brain relative to fibroblasts, it is also possible that the direct production of dopamine is also a factor. Clearly, further studies will be needed to study the basis of the effectiveness of TH-transfected muscle cells and the potential role of endogenousAADC. Knowledge of the mechanisms by which the TH and AADC genes are regulated in a cell-specific pattern within the vertebrate brain would contribute to this effort.

ACKNOWLEDGMENTS This work was supported by an NM grant to JH, and by an NIH postdoctoral fellowship to MIL.

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NOTE ADDED IN PROOF The following references present work relevant to this manuscript that have been published since the time of submission. For a description on the embryonic developmental expression of DDC, TH, serotonin and dopamine see Lundell and Hirsh, 1994. For a more detailed examination of elements essential in the alternative splicing of Ddc see Shen et al., 1993 and Shen and Hirsh, 1994. For additional work on the use of gene transfection in the treatment of Parkinson’s disease see Kang et al., 1993. Kang, U. J., Fisher, L. J., Joh, T. H., O’Malley, K. L., and Gage, F. H. ( 1993). Regulation of dopamine production by genetically-modified primary fibroblasts. J. Neurosci. 13: 5203-521 1. Lundell, M. J., and Hirsh, J. (1994). Temporal and spatial development of serotonin and dopamine neurons in the Drosophilu CNS. Dev. Biol. 165,385-396. Shen, J., Beall, C. J., and Hirsh, J. (1993). Tissue-specific alternative splicing of the DrosophiZu dopa decarboxylase gene is affected by heat shock. Mol. Cell. Biol. 13,45494555. Shen, J., and Hirsh, J. (1994). Cis-regulatory sequences responsible for alternative splicing of the DrosophiEu dopa decarboxylase gene. Mol. Cell. Biol. 14, 7385-7393.

REFERENCES A1bert.V. R., Allen, J. M.,andJoh, T. H. (1987). Asinglegenecodes foraromaticL-amino acid decarboxylase in both neuronal and non-neuronal tissues. J. Biol. Chem. 262: 9404-94 11. Albert, V. R., Lee, M. R., Bolden, A. H., Wurzburger, R. J., and Aguanno, A. (1992). Distinct promoters direct neuronal and nonneuronal expression of rat aromatic L-amino-acid decarboxylase. Roc. Natl. Acad. Sci. USA 89 12053-12057. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990). Basic local alignment search tool. J. Mol. Biol. 215: 4031110. Ando-Yamamoto, M., Hayashi, H., Sugiyama, T., Fukui, H., Watanabe, T., and Wada, H. (1987). Purification of L-dopa decarboxylase from rat liver and production of polyclonal and monoclonal antibodies against it. J. Biochem. 101: 405-414.

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Awgulewitsch, A., and Jacobs, D. (1992). Deformedautoregulatory element from Drosophila functions in a conserved manner in transgenic mice. Nature 358: 341-343. Baylin, S. B., Abeloff, M. D., Goodwin, G., Carney, D. M., and Gazdar, A. F. (1980). Activities of L-dopa decarboxylase and diamine oxidase (histaminase) in human lung cancers and decarboxylase as a marker for small (oat) cell cancer in cell culture. Cancer Res. 40: 1990-1994. Beall, C., and Hirsh, J. (1987). Regulation of the Drosophila dopa decarboxylase gene in neuronal and glial cells. Genes Dev. I : 510-520. Benezra, R., Davis, R. L., Lockshon, D., Turner, D. L., and Weintraub, H. (1990). The protein Id: a negative regulator of helix-loop-helix DNA binding proteins. Cell 61: 49-59. Bray, S. J., and Hirsh, J. (1986). The Drosophila virilis dopa decarboxylase gene is developmentally regulated when integrated into Drosophila melanogaster. EMBO J. 5: 2305-2311. Bray, S. J., and Kafatos, F. C. (1991). Developmental function of elf-I: an essential transcription factor during embryogenesis in Drosophila. Genes Dev. 9: 1672-1683. Bray, S. J., Johnson, W. A., Hirsh, J., Heberlein, U., and Tjian, R. (1988). A cis-acting element and associated binding factor required for CNS expression of the D. melanogaster dopa decarbxylase gene. EMBO J. 7: 177-188. Bray, S. J., Burke, B., Brown, N. H., and Hirsh, J. (1989). Embryonic expression pattern of a family of Drosophila proteins which interact with a CNS regulatory element. Genes Dev. 3: 1130-1145. Budnik, V., and White, K. (1988). Catecholamine containing neurons in Drosophila melanogaster: distribution and development. J. Comp. Neurol. 268 400-413. Christenson, J. G., Dairman, W., and Udenfriend, S. (1972). On the identity of DOPA decarboxylase and 5-hydroxytryptophan decarboxylase. Proc. Natl. Acad. Sci. USA 69: 343-347. Coge, F., Krieger-Poullet, M., Guillemot, J. C., Ferrara, P., Gros, F., and Thibault, J. C. (1989). Purification and partial sequencing of L-dopa decarboxylase from pheochromocytoma in rats. C. R. Acad. Sci. Ser. 3 309 587-592. deLuca, V., Marineau, C., and Brisson, N. (1989). Molecular cloning and analysis of cDNA encoding a plant tryptophan decarboxylase: comparison with animal dopa decarboxylases. Proc. Natl. Acad. Sci. USA 8 6 2582-2586. Desai, C., Garriga, G., McIntire, S. L., and Horvitz, H. R. (1988). Agenetic pathway for the development of the Caenorhabditis elegans HSN motor neurons. Nature 336 638-646. Doe, C. Q. (1992). Molecular markers for identified neuroblasts and ganglion mother cells in the Drosophila central nervous system. Development 116 855-863. Dynlacht, B., Attardi, L., Admon, A., Freeman, M., and Tjian, R. (1989). Functional analysis of NTF- 1, a developmentally regulated Drosophila transcription factor that binds neuronal cis-elements. Genes Dev. 3: 1677-1688. Ellis, H. M., Spann, D. R., and Posakony, J. W. (1990). extramacrochaerae, a negative regulator of sensory organ development in Drosophila, defines a new class of helix-loop-helix proteins. Cell 61: 27-38. England, B. P., Heberlein, U., and Tjian, R. (1990). Purified Drosophila transcription factor, Adh distal factor-I (Adf-I), binds to sites in several Drosophila promoters and activates transcription. J. Biol. Chem. 265: 5086-5093.

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England, B. P., Admon, A., and Tjian, R. (1992). Cloning of Drosophila transcription factor Adf-1 reveals homology to myb oncoproteins. Roc. Natl. Acad. Sci. USA 89 683-687. Fisher, L. J., Jinnah, H. A., Kale, L. C., Higgins, G. A., and Gage, F. H. (1991). Survival and function of intrastriatally grafted primary fibroblasts genetically modified to produce L-dopa. Neuron 6 371-380. Fortini, M. E., Lai, Z., and Rubin, G. M. (1991). The Drosophila f i - 1 and zfh-2 genes encode novel proteins containing both zinc-fingerand homeodomain motifs. Mech. Dev. 34: 113-122. Garrell, J., and Modolell, J. (1990). The Drosophilu extrurnucrochaefue locus, an antagonist of proneural genes that, like these genes encodes a helix-loop-helix protein. Cell 61: 39-48. Hashimoto, T., Nakano, Y., Morinaga, T., and Tamaoki, T. (1992). A new family of homeobox genes encoding multiple homeodomain and zinc finger motifs. Mech. Dev. 39: 125-1 26. He, X., Treacy, M. N., Simmons,D. M., Ingraham,H. A., Swanson,L. W., and Rosenfeld, M. G. (1989). Expression of a large family of POU-domain regulatory genes in mammalian brain development.Nature 340: 3542. Hirsh, J. (1986). The DrosophiZa melunoguster dopa decarboxylase gene: progress in understanding the in vivo regulation of a higher eukaryotic gene. In: Molecular Developmental Biology, pp. 103-1 16. Alan R. Liss, New York. Hirsh, J., Morgan, €3. A., and Scholnick, S. (1986). Delimiting regulatory sequences of the Drosophila mehnoguster Ddc gene. Mol. Cell. Biol. 6 454234557. Hyland, K., and Clayton, P. T. (1990). Aromatic amino acid deficiency in twins. J. Inherited Metab. Dis. 13: 301-304. Ichinose, H., Kurosawa, Y., Titani, K., Fujita, K., and Nagatsu, T. (1989). Isolation and characterization of a cDNA clone encoding human aromatic L-amino acid decarboxylase. Biochem. Biophys. Res. Commun. 164: 1024-1030. Ichinose, H., Sumi-Ichinose,C., Ohye, T., Hagino, Y, Fujita, K., and Nagatsu, T. (1992). Tissue-specificalternative splicing of the 1st exon generates 2 types of messengerRNAs in human aromatic L-amino acid decarboxylase. Biochemistry 31: 1154611550. Jackson, F. R. ( 1990). Prokaryotic and eukaryotic pyridoxal-dependentdecarboxylases are homologous. J. Mol. Evol. 31: 325-329. Jaeger, C. B., Ruggiero,D. A., Albert,V. R., Park, D. H., Joh, T. H., and Reis, D. J. (1984). Aromatic L-amino acid decarboxylase in the rat brain: immunocytochemicallocalization in neurons of the brain stem. Neuroscience 11: 691-713. Jaeger, C. B., Teitelman, G., Joh, T. H., Albert, V. R., Park, D. H., and Reis, D. J. (1983). Some neurons of the rat central nervous system contain aromatic L-amino acid decarboxylase but not monoamines. Science 219: 1233-1 235. Jiao, S., Gurevich, V., and Wolff, J. A. (1993). Long-term correction of rat model of Parkinson’s disease by gene therapy. Nature 362: 450-453. Johnson, W. A., and Hush, J. (1990). A Drosophilu “POU, protein” binds to a sequence element regulating gene expression in specific dopaminergicneurons. Nature 343: 467470. Johnson, W. A., McCormick, C. A., Bray, S . J., and Hush, J. (1989). A neuron-specific enhancer of the Drosophilu dopa decarboxylase gene. Genes Dev. 3: 676-686.

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Kandel, E. R., Schwartz, J. H., and Jessell, T. M. (1991). Principles of Neural Science. Elsevier Science Publishing Co., New York. Kang, U. J., and Joh, T. H. (1990). Deduced amino acid sequence of bovine L-amino acid decarboxylase: homology to other decarboxylases. Mol. Brain Res. 8: 83-87. Kang, U. J., Park, D. H., Wessel, T., Baker, H., and Joh, T. H. (1992). DOPA-decarboxylation in the striata of rats with unilateral substantia nigra lesions. Neurosci. Lett. 147 53-57. Konrad, K. D., and Marsh, J. L. (1987). Developmental expression and spatial distribution of dopa decarboxylase in Dmsophilu. Dev. Biol. 122: 172-185. Krieger, M., Coge, F., Gros, F., and Thibault, J. (1991). Different mRNAs code for dopa decarboxylase in tissues of neuronal and nonneuronal origin. Proc. Natl. Acad. Sci. USA 88: 2 161-21 65. Lai, Z., Fortini, M. E., and Rubin, G. M. (1991). The embryonic expression patterns if zfh-1 and zfh-2, two Drosophila genes encoding novel zinc-finger homeodomain proteins. Mech. Dev. 34: 123-134. Li, S., Crenshaw, E. B., Rawson, E. J., Simmons, D. M., Swanson, L. W., and Rosenfeld, M. G. (1990). Dwarf locus mutants lacking three pituitary cell types result from mutations in the POU-domain genepit-I. Nature 347: 528-533. Lovenberg, W., Weissbach, W., and Udenfriend, S. (1962). Aromatic L-amino acid decarboxylase. J. Biol. Chem. 237: 89-93. Lundell, M. J., and Hirsh, J. (1992). The zfh-2 gene product is a potential regulator of neuron-specific DOPA decarboxylase gene expression in Drosophilu Dev. Biol. 154: 84-94. Malicki, J., Cianetti, L. C., Peschle, C., and McGinnis, W. (1992). A human HOX4B regulatory element provides head-specific expression in Dmsophilu embryos. Nature 3 5 8 345-347. Maricq, A. V., Peterson, A. S., Brake, A. J., Myers, R. M., and Julius, D. (1991). Primary structure and function expression of the 5HT3 receptor, a serotonin-gated ion channel. Science 254: 432-437. Mastick, G. S., and Scholnick, S. B. (1992). Repression and activation of the Drosophilu dopa decarboxylase gene in glia. Mol. Cell. Biol. 12: 5659-5666. McGinnis, W., and Krumlauf, R. (1992). Homeobox genes and axial patterning. Cell 68: 283-302. McGinnis, N., Kuziora, M. A., and McGinnis, W. (1990). Human Hox-4.2 and Drosophila Deformed encode similar regulatory specificities in Dmsophilu embryos and larvae. Cell 63: 969-976. Morgan, B., Johnson, W. A., and Hirsh, J. (1986). Regulated splicing produces different forms of dopa decarboxylase in the central nervous system and hypoderm of Drosophilu rnelunoguster. EMBO J. 5: 3335-3342. Morinaga, T.,Yasuda, H.,Hashimoto,T., Higashio, K., andTamaoki, T. (1991). Ahuman alpha-fetoprotein enhancer-binding protein, ATBFl , contains four homeodomains and seventeen zinc fingers. Mol. Cell. Biol. 11: 6041-6049. Nagatsu, T., Ichinose, H., Kojima, K., Kameya, T., Shimase, J., Kodama, T., and Shimosato, U. (1985). Aromatic L-amino acid decarboxylase activities in human lung tissues: comparison between normal lung and lung carcinomas. Biochem. Med. 3 4 52-59.

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Neckameyer, W. S., and Quinn, W. G. (1989). Isolation and characterization of the gene for Drosophila tyrosine hydroxylase. Neuron 2: 1167-1175. Rahman, M. K., Nagatsu, T., and Kato, T.(1981). Aromatic L-amino acid decarboxylase activity in central and peripheral tissues and serum of rats with L-DOPA and L-5-hydroxytryptophan as substrates. Biochem. Pharmacol. 30: 645-649. Saudou, F., Boschert, U., Amlaiky, N., Plassat, J. L., and Hen, R. (1992). A family of Drosophila serotonin receptors with distinct intracellular signalling properties and expression patterns. EMBO J. ZZ: 7-17. Scholnick, S., Morgan, B. A., and Hirsh, J. (1983). The cloned dopa decarboxylase gene is developmentally regulated when reintegrated into the Drosophila genome. Cell 34: 37-45. Scholnick, S., Bray, S. J., Morgan, B. A., McCormick,C. A., andHirsh, J. (1986). Distinct central nervous system and hypoderm regulatory elements of the D. rnelanogaster dopa decarboxylase gene. Science 234: 998-1 002. Scholnick, S. B., Caruso, P. A., Klemencic, J., Mastick, G. S., Mauro, C., and Rotenberg, M. (1991). Mutations within the Ddc promoter alter its neuron-specific pattern of expression. Dev. Biol. 146 423-437. Sumi-Ichinose,C., Ichinose, H., Takahashi,E., Hori, T., and Nagatsu, T. (1992). Molecularcloning of genomic DNA and chromosomal assignment of the gene for human aromatic L-amino acid decarboxylase, the enzyme for catecholamine and serotonin biosynthesis. Biochemistry 31: 2229-2238. Taghert, P., and Goodman, C. S. (1984). Cell determination and differentiation of identified serotonin-immunoreactive neurons in the grasshopper embryo. J. Neurosci. 4: 989-1000. Thai, A. L. V., Coste, E., Allen, J. M., Palmiter, R. D., and Weber, M. J. (1993). Identification of a neuron-specific promoter of human aromatic L-amino acid decarboxylase gene. Mol. Brain. Res 17: 227-238. Treacy, M. N., He, X., and Rosenfeld, M. G. (1991). I-POU: a POU-domain protein that inhibits neuron-specific gene activation. Nature 3 5 0 577-584. Valles, A. M., and White, K. (1986). Development of serotonin-neurons in Drosophila mutants unable to synthesize serotonin. J. Neurosci. 6 1482-1491. Valles, A. M., and White, K. (1988). Serotonin-containing neurons in Drosophila melanogaster; development and distribution. J. Comp. Neurol. 268 400-413. Witz, P., Amlaiky, N., Plassat, J. L., Maroteaux, L., Borrelli, E., and Hen, R. (1990). Cloning and characterization of a Drosophila serotonin receptor that activates adenylate cyclase. Proc. Natl. Acad. Sci. USA 8 7 894043944. Wright, C. V. E. (1991). Vertebrate homeobox genes. Curr. Opin. Cell Biol. 3: 976-982. Wright, T. (1987). The genetics of biogenic amine metabolism, sclerotization and melanization in Drosophila melanogaster. Genetics 24: 127-222.

TRANSCRIPTION FACTORS IN MAMMALIAN DEVELOPMENT: MURINE HOMEOBOX GENES

S . Steven Potter

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. TheHomeobox . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

III. Homeo&x Gene Ciusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . IY Dispersed Homeobox Genes . . . . . . . . . . . . . . . . . . . . . . . . . . V. VI . VII . VIII .

Ix. X. XI .

The Paired Box . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Misexpression Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Targeted Homeobox Genes . . . . . . . . . . . . . . . . . . . . . . . . . . Mammalian-Dmsophila Homeobox Gene Exchanges . . . . . . . . . . . . HomeoboxGeneCircuitry: UpstreamandDownstreamGenes . . . . . . . A . Downstream . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B . Upstream Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . cis-Regulatory Elements . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Advances in Developmental Biochemistry Volume 3. pages 87.125 Copyright 0 1994 by JAI Press Inc All rights of reproductionin any form reserved ISBN: 1-55938-865-X

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1. INTRODUCTION The development of the single cell of the zygote into a mature mammal is a remarkable process. It starts with the microscopic fertilized egg and ends with a large conglomerate of trillions of cells, of thousands of different types, all interacting to form a functional unit. Encoded in the DNA of the zygote is a genetic program that directs this transformation. The essence of the developmental program is achieving correct temporal and spatial patterns of gene expression. Cell identities and behaviors are determined by the batteries of genes they express. If each cell of the developing mammal exhibits appropriate gene expression patterns then development can be expected to proceed normally. It follows that a full understanding of mammalian developmental genetics will require the characterization of each and every gene, to determine how it is activated at the right times and places during development. This is a rather depressing prospect, since the mammalian genome is believed to carry 50,000 to 100,000distinct genes, and analysis of the cis-regulatory elements of a single gene can consume the efforts of a laboratory for years. Fortunately there is a short-cut approach that should yield a solid understanding of the basic control mechanisms of development. While it is reasonable to assume that all of the genes of the mammalian genome have some significant function, it is nevertheless clear that some genes are more developmentally important than others. Most genes are of the “housekeeping” variety, or structural component genes, or genes that encode terminal differentiation products. These genes, while often essential for life or health, are driven by the process of development, and do not themselves control it. The genes that encode transcription factors, however, are responsible for controlling patterns of target gene expression. Each transcription factor can have an impact on the expression levels of a large battery of target genes, so relatively few transcription factor genes can govern the entire genome. As shown in Figure 1, transcription factor genes in turn must be regulated, resulting in a hierarchy that can be represented by a pyramid. Genes near the top of this pyramid are master switch genetic elements that initiate genetic cascades and control the developmental destinies of groups of cells. These genes activate other genes, some of which also encode transcription factors and in turn influence expression of still more genes. Such upper-level genes are sometimes responsible for initiating genetic programs that drive the formation of a structure or organ.

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Figure 7. Hierarchy of control of gene expression. A total of about 50,000 to 100,000 genes are necessary to encode a mammal, most of which encode housekeeping, structural component, or terminal differentiation gene products. Transcription factor genes regulate expression of the lowerlevel genes and are in turn controlled by other upper-level transcription factors.

I I . THE HOMEOBOX It is clear that the homeobox gene family includes members from the top of the pyramid. These genes were first identified by Drosophilu geneticists as particularly interesting; mutations in these genes often result in remarkable homeotic transformations of body parts. The dominant mutation, Antennupedia, for example, caused antenna1 imaginal disc cells to form leg structures protruding from the head, instead of antennae. When three of these genes (An@,ftz, Ubx) were cloned and sequenced they were found to carry, near their 3’ coding ends, similar sequence blocks of 180 bp (McGinnis et al., 1984a, 1984b). This area of common sequence was named the homeobox. This name has persisted although it is somewhat inappropriate, since in Drosophilu many maternal effect and segmentation genes, in addition to homeotic genes (which determine segment identities), carry the homeobox. The amino acid sequence encoded by the homeobox includes a helixturn-helix motif that has now been shown by X-ray crystallography to be structurally virtually identical to that found in various prokaryotic

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repressors (Kissinger et al., 1990). The helix nearest the carboxy terminus binds specific DNA sequences and is termed the recognition helix. Bacterial repressors, although structurally similar, share very little sequence similarity with the homeobox (Langhon and Scott, 1984). Interestingly, however, a computer search for sequence homology to the homeobox did reveal some small but significant similarity with the yeast Mat alpha 2 gene (Shepherd et al., 1984), which is involved in determining the yeast mating type and is believed to regulate the expression patterns of other genes. It is intriguing that the most developmentally interesting thing that yeast can do, switching of mating type, is in part controlled by a gene with amino acid coding sequence homology to the Drosophila homeobox genes. Considerable evidence indicates that the homeobox gene proteins are also involved in regulating the expression patterns of other genes. About one-third of the amino acids encoded by the homeobox are basic, and the homeobox proteins are found localized in the nucleus (White and Wilcox, 1985; Carroll and Scott, 1985). Furthermore, a number of reports have shown that homeobox proteins can bind to specific DNA sequences (Treisman et al., 1989;Hoey and Levine, 1988;Beachy et al., 1988). And more persuasive, perhaps, are the experiments that have used expression vectors carrying homeobox genes in Drosophila tissue culture cells. Ubx protein, for example, was found to autostimulate Ubx gene expression (a possible mechanism for maintaining the determined state) while repressing Antp transcription (Krasnow et al., 1989). Embryos mutant for particular homeobox genes have also been analyzed extensively for effects on expression patterns of other homeobox genes (Howard and Ingham, 1986). Ubx deletion mutants, for example, express Antp more posteriorly than normal, in segments where Ubx would normally be expressed (Hafen et al., 1984; Harding et al., 1985).Likewise, it appears that Ubx expression is normally repressed by iab-2, the next most posteriorly expressed homeotic gene (Struhl and White, 1985). In this manner it has been possible to begin to work out the network of interactions of the Drosophila homeobox genes.

111. HOMEOBOX GENE CLUSTERS Many mouse homeobox genes are found in four clusters (a-d) on chromosomes 2,6,11, and 15 (Fig. 2). Each Hox cluster carries about 10 genes and displays a number of interesting features. First, genes within

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Figure 2. The mammalian homeobox gene complexes. Sequence comparisons allow the genes of the four clusters to be vertically aligned into 13 paralogous groups, which are numbered at the top. The enclosed designations represent the recently agreed nomenclature, and the names beneath are the old murine and human nomenclatures. The genes of the Drosophila HOM-Ccomplex are listed at the bottom beneath their mammalian cognate groups. The clusters are illustrated with their 3' ends at the left with transcription occurring right to left as drawn, except for the Evx genes, and the Drosophila Dfd gene, which are reversed.

a cluster are transcribed in the same direction (Graham et al., 1989). Second, genes more 3' on the clusters are expressed at more anterior positions and at earlier times during development (Holland and Hogan, 1988; Schughart et al., 1988). Third, sequence comparisons indicate that each murine cluster is likely derived from a single primordial homeobox gene cluster, which is split in Drosophilu into the Bithorux and Antennupediu groups (Graham et al., 1989; Schughart et al., 1989). The homeobox genes of these clusters can be divided into 13 paralogous

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groups, with homeobox genes at equivalent positions within clusters showing the greatest homology. H o X - ~ ~for , example, is most like the Hox-b4, HOC-c4and Hox-d4 genes, which are found at comparable positions in other clusters. Furthermore,as discussed in more detail later, it has been shown that genes at the more 3’ ends of the clusters are induced by lower levels of retinoic acid, a putative morphogen. And finally, it is interesting to note that the human and murine Hox clusters are remarkably alike in sequenceand organization, with encoded equivalenthomeodomains that are often identical.

IV. DISPERSED HOMEOBOX GENES In addition to the four main complexes there are a number of scattered, unique chromosomal positions where homeobox genes are also found. Although they are individually dispersed, comparisons show that these homeobox genes can be grouped into subfamilies based on similarities in encoded homeodomains. This list of dispersed homeobox genes includes the K-2 (MHox) (Kern et al., 1992; Cserjesi et al., 1992); S-8 (Opstelten et al., 1991); Pax 3, Pax 6, and Pax 7 (Walther et al., 1991) genes with paired type homeodomains; the En-1 and En-2 engrailed genes (Joyner and Martin, 1987); the msh-1, msh-2, and msh-3 genes (Hill et al., 1989; Robert et al., 1989, 1991; Davidson et al., 1991; Holland, 1991;Monaghan et al., 1991;Mackenzieet al., 1991);the DZx-1 and Dlx-2 genes (Price et al., 1991; Robinson et al., 1991; Porteus et al., 1991, 1992; Dolle et al., 1992); the Mox-1 and Mox-2 genes (Candia et al., 1992); the Gsh-1 and Gsh-2 genes (Singh et a]., 1991); the cdx-1 and cdx-2 genes (Duprey et al., 1988);Dbx-1 (Lu et al., 1992);and goosecoid (Blum et al., 1992). Most of these genes show extreme homology to a Drosophila gene that is also dispersed. The mouse cdx-1 and cdx-2 genes are very similar to the caudal gene of Drosophila, the En-1 and En-2 genes are similar to engrailed, the Dlx-1 and Dlx-2 genes are similar to distal-less and so on. It is interesting that single homeobox genes in DrosophiZa are generally represented by at least two close homologues in the mouse. Presumably the increased number of genes is needed to encode the greater phenotypic complexity of the mammal. These dispersed homeobox genes often show rather unique expression patterns. Both the Dlx and Dbx genes, for example, are expressed in the developing embryonic forebrain, where no expression is detected for the Hox complex genes. And the S-8 and K-2 (MHox) genes are expressed

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uniquely in mesodermalcells, which again is a pattern distinctly different from those of the clustered Hox genes. The msh-1 and msh-2 genes are expressed in a range of developing tissues including the eyes, limbs, and heart. There is a growing body of evidence indicating that these msh genes indeed play an important role in limb development (discussed later).

V. THE PAIRED BOX The paired box is another sequence motif that is very well conserved during evolution and is found in a number of genes known to be of developmental significance in “lower” organisms. The paired box encodes the 128-aminoacid paired domain, which binds DNA (Chalapakis et al., 1991), and shows similarity, with a helix-turn-helix motif, to the homeodomain. Eight members of this gene family have been isolated by Peter Gruss and his colleagues and named Pax I-Pax 8 (Walther et al., 1991). These genes are generally expressed in a manner different from that of the clustered Hox genes. The Pax-3, Pax-7, and Pax-8 genes, for example, are expressed along the entire anteroposterior axis in the developing neural tube, whereas Hox genes have sharp anterior boundaries of expression in the neural tube. Transverse sections, however, show that the Pax genes are often expressed in limited sectors of the neural tube with Pax-6, for example, expressed only in the ventral part of the neural tube in day nine embryos (Walther and Gruss, 1991). While Pax gene expression in the CNS, in particular the neural tube and hindbrain, is common, there are exceptions. Pax-I is expressed in the sclerotome of differentiating somites and at later stages in the intervertebral discs of the developing vertebral column (Deutsch et al., 1988). The genes with paired boxes often, but not always, have homeoboxes as well. The homeoboxes that are associated with paired boxes have distinctive sequence characteristics that set them apart as a separate homeobox subfamily. Interestingly, these paired-type homeoboxes in mouse can sometimes reside on genes such as the K-2 (MHox)and S-8 paired-type homeobox genes, which have not been found to carry paired boxes. Mutations in paired box genes of mouse and humans have now been shown to cause interesting developmental abnormalities. The classical mutation undulated in mouse was found to be the result of an alteration

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in the Pax-I gene (Chalapakis et al., 1991), giving abnormal vertebral column development. Mutation of P a x 3 was shown to be responsible for the splotch mutation (Epstein et al., 1991), which in heterozygotes gives white spotting of the abdomen, which is apparently due to aberrant melanocyte migration. Homozygotes fc - the splotch mutation display variable phenotypes depending on the allele and the genetic background, but they generally suffer neural tube defects that can include exencephaly, meningocele, and spina bifida. Interestingly, mutation of the human homolog of Pax-3, HuP2, is responsible for Waardenburg’s syndrome, which manifests as a combination of deafness and pigmentary disturbances (Tassebehji et al., 1992; Baldwin et al., 1992). Moreover, mutation of Pax-6 and its human homologue have been shown to be responsible for the small eye (sey) mutation in mouse and congenital aniridia(1ack of iris) in humans (Hill et al., 1991). The small eye mutation is again semidominant, with heterozygotes showing distinctly smaller than normal eyes. The homozygotes have no visible eyes and incomplete nasal cavities, resulting in an inability to breathe during suckling and early postnatal death.

VI. MISEXPRESSION STUDIES The Antennupediu mutation in Drosophila results from a chromosomal rearrangement that brings the Antennupediu coding sequences under the control of a heterologouspromoter, causing misexpression during development (Frischer et al., 1986). It is quite informative to observe that expression of Antennupediu in antennal imaginal discs can drive these groups of cells down the wrong developmental pathway, causing them to form legs instead of antennae. The Antennupediu gene is apparently capable of initiating a genetic program that results in leg formation. Interestingly, this program can only be imposed on certain susceptible groups of cells. When the heatshock promoter is connected to Antennupediu in transgenic flies, heatshock gives transcription of Antennupediu in almost all cells of the embryo. Nevertheless, only the antennal imaginal discs are misdirected to generate legs (Gibson and Gehring, 1988). One presumes that the transcription factor milieu present in other cell types is not appropriate to allow the initiation of the genetic cascade that forms leg structures. It seems reasonable to hope that misexpression analysis of murine homeobox genes might also provide clues to their function.The f i s t such

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study was reported by Wolgemuth et al. (1989). These experiments had the dual purpose of searching for Hox-1.4 cis-regulatory elements and examining the biological effects of Hox- 1.4 overexpression. The normal Hox-1.4 promoter was used, with sequences extending 10 kb upstream of the homeodomain, and abutting the next gene in the cluster, Hox- 1.3. The transgene was tagged at the 3’ end with SV40 DNA (including some coding sequences of the large T antigen), so that transcripts from the transgene could be readily distinguished from transcripts of the endogenous Hox-1.4 gene. Northern blots and in situ hybridization showed that the transgene expression pattern resembled that of the endogenous gene, with some quantitative variation from tissue to tissue. In particular, two transgenic lines showed considerably higher levels of transgene expression in the gut. This correlated with a very interesting phenotype. The mice developed a profound enlargement of the colon, termed megucolon. These mice moved feces through the colon with great difiiculty, suffering what eventually became terminal constipation. These experiments were of particular note, since they were the first to associate an abnormal phenotype with a mammalian homeobox gene. Balling et al. (1989) next reported the effects of ectopic expression of the Hox- 1.1 gene in transgenic mice. They used an approach similar to that which had been successful in Drosophilu. Instead of a heatshock promoter they used the p-actin promoter, which also proved to give near-ubiquitousexpression.This promoter was connected to the Hox-1.1 coding sequences, and transgenic mice were made. A number of interesting malformationswere observed. The founders expressing the transgene typically died within a few days of birth, but one mosaic founder survived and provided a source of a number of additional transgenic progeny for analysis. These transgenics showed open eyes at birth, small weight, detached pinnae, and cleft secondary palate. Kessel et al. (1990) later reported that these animals also manifested some remarkable malformations of bones in the region of the cranio-cervical transition. Most interesting perhaps was a missing supraoccipital bone and an apparent splitting of the exoccipital bone, with a proatlas, not normally present, now located beneath it. These experiments suggested a role for mammalian homeobox genes in determining positional information and establishing segment identity. This further supported the conclusion that homeobox genes play similar roles in the development of mammals and flies. Two subsequent studies by Kaur et al. (1992) and McLain et al. (1992) used the same p-actin promoter to drive misexpression of the Hox-2.2

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and Hox-2.3 genes. This promoter appears to be particularly well suited for ectopic expression experiments in transgenic mice, since the earlier work in Drosophilu indicated that only a small subset of all tissues are sensitive to the morphogenetic effects of misexpression of a particular homeobox gene. The use of a well-characterized promoter that drives near-ubiquitousexpression, such as hsp70 for Drosophilu and p-actin for mice, allows the efficient screening of many tissues for possible developmental perturbation. Also, the use of one promoter for multiple studies facilitates functional comparisons of Hox genes. The phenotypes observed in the Hox-2.2 and Hox-2.3 mutants were similar in many respects to that seen when Hox-1.1 was misexpressed. In the craniocervical transition region, for example, all three groups of misexpression mutants showed reduced or absent supraoccipital bones, malformed basioccipitals, and split exoccipital bones. The most extreme effect in this region, however, the formation of the additional proatlas, was only observed for the Hox-1.1 mutants. Interestingly, the Hox-2.2 and Hox-2.3 mutants showed other skeletal abnormalitiesnot seen in the p-actin Hox-1.1 mice. For p-actin Hox-2.2 an apparently mosaic transgenic mouse had numerous vertebral segmentsthat were “confused”with respect to anterior-posteriorpositional identity. One vertebra, for example, had the 13th thoracic rib protruding from one side, and no rib on the other side. It apparently was half a 13 thoracic and half a first lumbar segment. Furthermore, the pelvis was attached one segment more posterior to the vertebral column on one side relative to the other. Careful analysis showed numerous thoracic vertebrae were of mixed positional identity, with a number 12 rib on one side and a number 11 on the other, or articulated with two ribs on one side and one on the other. Hox-2.3 misexpression also gave some dramatic skeletal abnormalities unique to this transgene. For example, the tuberculum anterior was shifted from the sixth to the seventh cervical vertebrae, again suggesting a shift in positional identity. Furthermore, an extra pair of ribs was found extending from what would normally represent the seventh cervical vertebrae. These additional ribs fused ventrally with those from the first thoracic vertebrae prior to attachment to the sternum. Moreover, the Hox-2.3 mutants showed ventricular septa1 defects in the heart that were not observed with the other constructs. In each of these studies dozens of transgenic animals were made and characterized. This is necessary because there is considerable individual variability in phenotype, presumably because of different sized transgene concatemers and different chromosomal integration sites. In some foun-

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der mice the transgenes do not express and the mice are normal. Other mice might show particularly extreme phenotypes due to enhancer elements flanking the transgene insertion site. The net result is that a number of mice must be examined and an average effect emerges, with some mice milder and some more extreme in phenotype. For the p-actin Hox-1.1, -2.2, and -2.3 mice expression results in early postnatal death. This means that no permanent mutant lines can be established and makes the analysis more difficult. Occasionally, however, a mosaic founder will survive and can be used to produce significant numbers of transgenic mutant progeny. Such mosaic founders were employed in both the p-actin Hox-1.1 and p-actin Hox-2.3 studies. Although such animals are extremely useful, it is important to note that all the progeny represent only one transgene insertion site and concatemer size. Therefore, although many animals can be readily made from such a founder, they do not contribute significantly to the description of the range of phenotypes possible with that particular transgene construct. A key question in the study of mammalian homeobox genes concerns the degree of functional overlap existing between different members of this large family. Because four different clusters have been duplicated from one original, paralogous genes at similar positions in different clusters have near-identical homeobox sequences and often exhibit similar expression patterns during development (Schughart et al., 1989; Graham et al., 1989; Kessel and Gruss, 1991). Anumber of studies have shown that many homeodomain proteins bind to similar or identical DNA sequences, suggesting common targets (Desplan et al., 1988; Treisman et al., 1989). In addition, gene targeting studies have shown that “knock out” of homeobox genes results in developmentalabnormalities less severe than would be predicted based on expression patterns. For example, targeted disruption of En-2 caused only foliation defects in the cerebellum (Joyner et al., 1991). Also, defects were not detected in Hox-1.5 and Hox-1.6 homozygous mutants in many tissues where these genes are normally expressed (Chisaka and Capecchi, 1991;Lufkin et al., 1991; Chisaka et al., 1992). Phenotypic analysis of dominant mutants generated by homeobox gene ectopic expression provides another means of determining functional overlap. Rather than directly monitoring normal gene function, this approach measures the ability of gene misexpression to perturb normal development. Functional overlap between genes is assessed by similar developmental defects caused by misexpression of different homeobox genes.

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As measured in this manner the Hox- 1.1, Hox-2.2, and Hox-2.3 genes exhibit significant functional overlap. This perhaps is not surprising since these genes have been grouped together based on sequence comparisons between homeoboxes (Graham et al., 1989). The homeobox of Hox-2.3 is identical for 59 of 61 amino acids to that of Hox-1.1 and for 60 of 61 amino acids to the Hox-2.2 homeobox (Graham et al., 1989). The in vivo functional ectopic expression assay demonstrates that this structural similarity is reflected in biological activity. Several other studies have used homeobox gene promoter swaps or altered versions of the normal promoter to generate misexpression mutants. Interestingly, a number of resulting homeotic transformationshave been observed, again involving the thoracic-lumbar boundary and the craniocervical transition regions. Jegalian and DeRobertis (1992) found that a segment of the normal Hox-3.3 promoter could drive ectopic posterior Hox-3.3expressionthat converted the Fmt lumbar segment into an additional thoracic segment, with attached ribs. One mosaic founder produced a number of transgenic progeny, 50% of which showed this transformation. Aside from rib cage defects, these animals were apparently fertile and normal. An analysis of a number of transient transgenics found addition-independenttransgenics with extra rib phenotypes, confirming that the effect was not dependent on an insertional mutation or a unique transgene integration site. Pollock et al. (1992) used a transgene construct with the Hox-1.4 promoter connected to the Hox-3.1 coding sequences to generate misexpression mutants. The Hox-1.4 promoter was shown to extend the anterior limits of Hox-3.1 expression, as predicted, but the phenotypic effects were observed in more posterior regions, where the transgene resulted in increased levels of Hox-3.1 transcription. The most remarkable effects were changes in vertebral patterning. Again, the first lumbar vertebral segment was converted to a thoracic segment, with attached ribs. Likewise, the thoracic segments above the lumbar vertebrae were anteriorized, as determined by rib morphologies. Moreover, dysmorphologies of the gut were observed and may have been related to the early postnatal deaths generally observed in mice expressing the transgene. Lufkin et al. (1992) also used a homeobox gene promoter exchange to drive misexpression of the Hox-4.2 gene. The Hox-1.6 promoter was observed to extend the anterior transcription boundary, and this region of ectopic expression gave a number of abnormalities,including absence

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or reduction of supraoccipital and exoccipital bones, and a striking addition of one to four neural arch bony structures anterior to the atlas. It is interesting to note that many of the homeobox genes used in these misexpression studies are closely related, and many of the observed effects are similar. The Hox-1.1 and Hox-2.3 genes are paralogous, as are the Hox-2.2 and Hox-3.3 genes, and all four of these genes are extremely closely related as measured by homeodomain amino acid sequence comparisons.Hox-1.1, -2.2, and -2.3 misexpression gave related malformations in the craniocervical transition, and Hox-2.2, -2.3, and -3.3 all gave transformations of vertebral segments with ribs. Misexpression of Hox-3.1, which is also very closely related to these other four, again gave homeotic transformations of vertebral segments contributing to the rib cage. In addition, misexpression of Hox-4.2, which is somewhatdifferent from the others studied, also gave some similar craniocervical transition abnormalities, such as a missing supraoccipital bone. Although these studies employed a battery of distinct promoters, the observed similarities in effects again strongly suggest that many of the proteins encoded by Hox genes are closely related functionally. These experiments further confirm a role for Hox genes in specifying segment identity, since misexpression alters segmentation patterns. Although useful, the murine transgenic misexpression studies conducted to date do not readily reveal the specific normal function of a Hox gene. Misexpression may generate an extra rib or delete a supraoccipital bone, but these are clearly not normal functions. Morgan et al. (1992) have used a retroviral system to misexpress mammalian Hox genes in the limb of the developing chick. These experiments more directly addressed the question of normal function. Five genes of the Hox-4 cluster have been shown to be expressed in a nested set of overlapping domains in the developing limb bud (Dolle et al., 1989). In the most distal, posterior portion of the limb bud all five of these genes, Hox-4.4, -4.5, -4.6, -4.7, and -4.8, are expressed, but as one moves anterior and proximal, fewer are active. In the most anterior region only Hox-4.4 is expressed, and just posterior to this -4.4 and -4.5 are expressed, and so on. It has been hypothesized that this expression pattern plays a role in specifying positional identity along the anterior/posterior axis of the limb. Morgan et al. (1992) directly perturbed Hox expression to determine effects on limb development. They inserted the Hox-4.6 coding sequences into the RCAS replication competent retroviral vector. Early limb buds (stages 10-12) were locally infected, to achieve complete limb infection by stage 21, when cells can still alter their developmental fate.

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The Hox-4.6 gene is normally expressed in the developing leg only in regions that generate digits 11, III, and IV. The retroviral infection therefore produced ectopic expression only in the region that normally produces digit I. Interestingly, this ectopic expression often resulted in conversion of this digit into an additional digit 11. That is, this alteration in the pattern of Hox gene expression in the most anterior part of the limb produced a homeotic transformation of digit I to digit 11. These results strongly support a role for the Hox-4 genes in specifying cell identity during limb development. The work of Song et al. (1992) suggests a role for the Hox-7.1 gene in maintaining cells in the dedifferentiated state. This gene is normally expressed in the distal mesenchyme of the developing limb. This region, termed the progress zone, lies beneath the apical ectodermal ridge and represents an area where undifferentiated cells proliferate. As cells exit the progress zone they assume positional identity and eventually form the components of the limb. Song et al. (1992) stably transfected F3 myoblasts with a Hox-7.1 expression construct and found evidence that the cells were now more dedifferentiated. First, Hox-7.1 expression blocked or delayed myotube formation when the cells were grown in low serum. Second, cells with high levels of Hox-7.1 expression showed much reduced levels of MyoD1 expression. Third, the transfected cells showed morphological changes suggestive of cellular transformation, and they were now capable of colony formation in soft agar. Furthermore, one of the stably transfected cell lines was able to produce tumors following subcutaneous injection into nude mice. It is interesting to note that transfection of the very closely related Hox-8.1 gene did not give these effects, indicating that even though Hox-7.1 and Hox-8.1 have nearly identical homeodomains, they are functionally distinct.

VII. TARGETED HOMEOBOX GENES Gene targeting, by homologous recombination in embryonic stem cells, promises to be a very powerful tool in the functional analysis of murine homeobox genes. This technique allows the generation of null mutants as well as mice with more subtle modifications of homeobox genes. Resulting homozygous mice will display developmental defects that

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directly reveal which developmental processes required the altered homeobox gene. The results to date are similar to what one might have predicted based on previous mutational analysis of this family of genes in Drosophila. Mice with modified homeobox genes have displayed malformations, deleted structures, and homeotic transformations. In general, however, the phenotypes observed in mice with homeobox gene mutations tend to be less dramatic than those seen in flies. The prevailing view is that this is the result of overlapping function of duplicated homeobox genes in mice. Indeed, almost every fruit fly homeobox gene appears to have at least two counterparts in the mammalian genome. There is one homeotic gene complex in Drosophila, the combined Antennapedia and Bithorax clusters, and there are four equivalent homeobox gene clusters in mice and humans. This duplication extends to the dispersed homeobox genes as well, with one Engrailed gene in Drosophila, and two in mice, one msh gene in Drosophila and two in mice, one distalles gene in Drosophila and three in mice, and so on. And the duplicated murine homeobox genes are still remarkably similar, with near-identicalamino acid sequencesfor encoded homeodomain. It is rare for homeodomains of paralogous genes in the cluster to differ at more than two or three positions, and the same is true for members of the groups of dispersed homeobox genes. The homeodomains of Gsh-1 and Gsh-2, for example, have two amino acid differences, En-1 and En-2 have three differences, and msh-1 and msh-2 have two differences. This striking sequence similarity is quite compatible with the concept of overlapping function. The engrailed gene En-2 was the first homeobox gene to be successfully targeted (Joyner et al., 1991). In this case the resulting developmental defects were remarkably innocuous. The homozygous mutants were viable and fertile and appeared externally to be normal. They did, however, have striking foliation abnormalities of the cerebellum. This is particularly interesting because the developing cerebellum is one place where the En-2 gene is expressed and En-1 is not, supporting the notion of functional overlap. Other homeobox gene targeting experiments have resulted in more dramatic developmental defects. Homozygous Hox- 1.5 null mutants showed a number of alterations (Chisaka and Capecchi, 1991). The mice died at or shortly after birth, apparently from circulation and/or pulmonary problems. These animals showed hypertrophy of the atria and

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enlargement of major veins, indicating cardiovasculardysfunction.Mice were found with persistent truncus arteriosis, and patent ductus arteriosis, as well as missing right carotid artery and valve and septa1defects. In addition, the mice were athymic, most lacked parathyroid tissue, and the thyroids were greatly reduced in size. Skeletal defects in the head and neck region were also observed. And the muscles surrounding the esophagus, and those connecting to the epiglottis, were disordered and may have contributed to breathing difficulties that resulted in the stomach and intestines of the homozygotes being filled with air. This severe developmental modification resulting from targeting of HOX1.5 shows that the paralogous Hox-2.7, Hox-3.6, and Hox-4.1 genes of this subfamily do not fully overlap with Hox-1.5 functionally. Nevertheless, development appeared normal in lungs, stomach, spleen, and kidneys, even though the HOX-1.5gene is normally expressed during their development, suggesting possible functional overlap at these locations. The HOX1.5 knockout gave primarily deletions of structures and malformations, but no homeotic transformations. Such effects are also seen in Drosophilu for some homeobox gene mutations (McGinnis and Krumlauf, 1992), so these results do not suggest distinct developmental roles for mammalian and Drosophilu homeobox genes. The targeting of the Hox- 1.6 gene has also been very informative. This gene, as predicted from its 3’ most position in the Hox-1 cluster, has a very rostral boundary of expression that correspondsto the rhombomere four region of the presumptive hindbrain at day 7.5 of development. Lufkin et al. (1991) have described the resulting phenotype when the gene is targeted in a manner that is predicted to eliminate all functional transcripts. The homozygous mutants die shortly after birth, apparently of anoxia, with lungs that are never inflated with air. This lethal effect is due to defects in the glossopharyngeal and vagus nerves as well as the hindbrain itself. These mice cannot transmit respiratory stimuli to the hindbrain, which is itself perturbed and may not have functional respiratory centers. It was further observed that neural tube closure was delayed, several bones of the skull were malformed, and the inner ear was distinctly abnormal, with absent semicircular canals and other perturbations. Interestingly, Chisaka et al. (1992) targeted the Hox-1.6 gene in a different manner and observed some distinct phenotypic effects. Alternate RNA processing allows the Hox-1.6 gene to encode two different proteins, one with the homeodomain and one without it. The experiments

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by Lufkin et al. (1991) eliminated both transcripts, but the experiments by Chisaka and Capecchi (1991) only disrupted the synthesis of the homeodomain containingprotein. The homozygotes again died of anoxia shortly after birth and showed similar cranial nerve and hindbrain defects. These mice, however, showed no delay of closure of the neural tube. Furthermore,these mice showed middle ear and external ear defects not reported for the Hux-1.6 null mutants, and there were distinct differences in the development of the hindbrain in the two types of mutants. Some of these contrasting effects may be due to mouse strain differences or differences in observation. Nevertheless, the data suggest significant phenotypic differencesresulting from the two types of targeting. Both the Hux-1.5 and Hux-1.6 mutants show the most severe effects in the general regions of most anterior expression. In these regions cell types of many differentembryonic origins can be affected. It appears that in these areas the change in the combinatorial code of Hux genes expressed has altered developmental programs, resulting in the observed abnormalities. It is clear, however, that a simple combinatorial Hox code is not the complete explanation, since more posterior regions where the Hux-1.5 and -1.6 genes are normally expressed also had their Hux codes altered in the mutants, yet they developed normally. This indicates that some Hox genes can weigh more heavily in the code than others. That is, some Hux genes appear to have more dominant effects than others. In general, there is a tendency for “posterior prevalence.” Hux genes that are expressed only in more posterior regions often have dominant effects over genes with more anterior boundaries of expression. The targeting of the Hux-3.1 gene by Mouellic et al. (1992) yielded the first homeotic transformations. Homozygous mice displayed alterations in vertebral segment identities, with, for example, the first lumbar vertebra transformed into a thoracic segment with 14th pair of ribs. Careful analysis of rib morphologies indicated that a number of thoracic vertebrae were also anteriorized. Interestingly, these gene ablation effects resembled the rib cage abnormalities observed in Hux-3.1 ectopic expression mutants (Pollock et al., 1992). It has been speculated that a transcription complex might be disrupted by the absence of an essential protein, or by its overexpression, which could titrate out other essential components (Pollock et al., 1992).

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Although most Hox-3.1 mutants died within a few days of birth, some survived and were fertile. These adult homozygotes were smaller than normal and showed nervous system disorders, with wavering movements and abnormal reflexes. In addition, the digits of the forelimbs were clenched and some animals had spasmodic contractions of the hindlimbs or back muscles. This brief discussion of targeted Hox mutations illustrates that malformation, deletion, and homeotic transformation can all result. It is likely that all of the cloned mouse homeobox genes will be targeted relatively rapidly as this technique becomes more routine. Careful analysis will reveal how a mutation in one Hox gene alters expression patterns of others, unraveling the genetic circuitry of development. Interbreeding will produce multiple mutants that lack, for example, all Hox genes of a paralogous group. Examination of resulting phenotypes will determine the degree of functional overlap among Hox genes. In addition, use of the “two-step” or “in and out” targeting approach will allow the introduction of subtle mutation into Hox genes. Even single base pair changes in Hox genes can now be generated, making it possible to conduct a fine-tuned in vivo functional dissection of both coding and regulatory sequences. Domains can be deleted, and effects on expression of other Hox genes and resulting phenotype can be assayed. Single amino acids in the DNAbinding domains can be altered and in vivo functional effects determined. Two-step gene targeting can also allow tissue-specific gene ablation. As cis-regulatory elements controlling the expression pattern of Hox genes are further studied, it will become possible to alter these sequences to modulate tissue-specific expression patterns during development. For example, a regulatory element driving expression in the kidney could be deleted, leaving expression patterns in other positions unchanged. Particular effects on kidney development could then be determined in the absence of other developmental perturbations. This may prove a very useful improvement of the current gene targeting techniques, which generally are limited to analysis of the first lethal defect. If the animal could be rescued past this defect, others might become apparent at later times. That is, manipulation of regulatory sequences will facilitate a tissue-by-tissue,position-by-positioncharacterizaticrnof Hox gene function in development.

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VI I I. MAMMALIAN- DROSOPHlf A HOMEOBOX GENE EXCHANGES The most dramatic illustration of the evolutionary conservation of homeobox genes comes from experimentsthat swap Drosophilu and mammalian cognates and test for functional equivalence. McGinnis and co-workers have reported remarkable phenotypic similarities in flies that misexpress Drosophilu homeobox genes or their mammalian counterparts. Indeed, this ectopic expression assay in Drosophila indicates that the Drosophilu and cognate mammalian homeobox proteins are essentially functionally identical. Malicki et al. (1990) connected the Drosophilu Hsp-70 promoter to the mouse Hox-2.2 gene and made transgenic flies. It should be noted that the Hux-2.2 gene is quite similar to the Drosophilu Antennupediu gene, with 57 of 61 amino acids in the encoded homeodomains identical. It had previously been shown that ectopic expressionof the Antennupediu gene, driven by the Hsp-70 promoter, causes the development of thoracic structures in place of head structures (Schneuwly et al., 1987; Gibson and Gehring, 1988). In particular, in larvae there are transformationsthat include the appearance of thoracic dentricles and hairs on the head and transformation of the first thoracic dentricle pattern toward that of the second thoracic segment. In adults the most dramatic effect is transformation of the antenna to second thoracic legs. Strikingly, misexpression of the Hox-2.2 gene results in near-identical phenotypic effects in both larvae and adults. Again there are thoracic dentriclesand hairs that appear on the head of the larvae, and again, expression of Hox-2.2 can cause the formation of legs in place of antennae on the heads of adults. Similar functional similaritieswere found when McGinnis et al. (1990) expressed the human Hox-4.2 gene under the control of the Hsp-70 promoter in transgenic flies. Hox-4.2 is a mammalian cognate of the Drosophilu Deformed gene, and misexpression of either the Drosophilu or mammalian counterpart was found to produce similar phenotypic effects, including loss of the ventral eye and orbital region of the adult head. Furthermore, the Hox-4.2 gene was shown to be able to activate the endogenous Dfd gene. This is of particular interest because the Drosophilu Dfd gene can autostimulate, so it is clear that in these flies the Dfd and Hox-4.2 genes share a common target, the Dfd gene itself. On the other hand, however, this complicates interpretation of the

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phenotypic effects, which could be directly due to Hox-4.2 protein or could simply result from Dfd activation. Zhao et al. (1993) have described a very interesting comparison of the functional specificities of the mouse Hox-1.3 gene and the Drosophilu sex combs reduced (Scr) gene, as measured by misexpression in transgenic flies. Each gene was connected to the Hsp 70 promoter, and effects on both larval and adult development were determined by heat shock induction at 6.5 hours after egg laying or by multiple heat shocks at the second and third larval stages, respectively. The resulting phenotypes of Hsp-Scr flies were largely consistent with those described previously by Gibson et al. (1990). In larvae the second and third thoracic segments are converted toward thoracic segment one identity. Head segments were also affected, with disruption of head involution and reduction of the cephalopharyngeal skeleton. In adults ectopic Scr expression gave a complex constellation of effects that included reduced compound eyes, aristae to tarsal transformation,and disfigured maxillary and labial palps. Heat shock induction of the Hsp-Hox-1.3 transgenic flies was found to give a near-identical set of phenotypic abnormalities. Again, larval thoracic segments 2 and 3 were transformed toward thoracic segment one, and similar defects in the larval head (failed head involution, for example) were observed. Misexpression of Scr and Hox- 1.3 also gave similar malformations of adult structures, indicating a high degree of functional equivalence for the coding sequences of these two genes. Furthermore, the ectopic expression of Hox-1.3 was shown to have no effect on expression levels of the endogenous Scr and Dfd genes, indicating that the Hox-1.3 protein itself was effecting Drosophilu development, and it was not merely activating a Drosophilu homeobox gene which then produced the developmental defects. Of particular interest, Zhao et al. (1993) found that the Hox-1.3 protein could activate fork head (fkh),a known target of the Scr protein. Heat shock of the Hsp-Hox-1.3 transgenic flies induced the same ectopic expression pattern ofjkh as heat shock of hs-Scr flies. Thefkh gene itself encodes a transcription factor that plays a role in development of the gut and salivary gland tissues. Sequence comparisons suggest thatfkh is the Drosophilu counterpart of the mammalian HNF-3 gene family, which is expressed in mammals in organs, such as liver, lung, and intestines, which are derived from the embryonic gut tube (Lai et al., 1991). Since Hox-1.3 expression has also been detected in liver and lung (Odenwald et al., 1987; Garbem et al., 1989), the intriguing suggestion has been

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made that in mammals the Hox- 1.3 gene may regulate HNF-3 expression (Zhao et al., 1993). Indeed, the main conclusion of each of these three studies expressing mammalian Hox genes in flies is that the genetic circuitry of development is much better conserved than one might expect from a comparison of fruit fly and mammalian anatomies. In Drosophila the human Hox-4.2 gene activates a known target of the Drosophila cognate, the Deformed gene itself. And the murine HOX-1.3gene activates t h e m gene, as does Scr. And in each case, including Hox-2.2, misexpression in flies reproduces the phenotype generated by misexpression of the Drosophila cognate homeobox gene, indicating evolutionary conservation of the downstream genetic pathways that are activated. Although the details will surely differ, it appears that we will be able to learn a surprising amount about the genetic circuitry of mammalian developmentby studying Drosophila. In mammals the identification of upstream regulators and downstream targets of homeobox genes has been slow. Perhaps more effort should be devoted to the cloning and characterization of mammalian homologues of known regulators and targets in Drosophila. Just as Drosophila provided the key to the isolation and characterization of mammalian Hox genes, so it might also provide the key to allow the unraveling of the genetic regulatory network in mammals. Indeed in some regards the Drosophila system is the Rosetta stone of development. The functional similarities of mammalian and Drosophila homeobox genes are even more surprising when one considers that almost all of the sequence homology between cognates is restricted to the relatively small homeodomain region. The functional equivalence of mammalian and Drosophila counterpart homeobox genes argues that these conserved amino acids determine functional specificity. And from another perspective it is clear that although the homeodomain itself is a well-conserved unit, only a few amino acids within it confer functional specificity, since most homeodomains are very similar in sequence although functionally distinct. The Drosophila Antennapedia and sex combs reduced homeodomains, for example, are identical at 56 of 6 1 amino acids although they are functionally distinct. That is, some key amino acids within the homeodomain have been preserved over the 600 million years of evolutionary time that separate Drosophila and mammals, allowing them to maintain functional similarity. Yet two different Drosophila homeobox

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genes encode a few key homeodomain amino acid sequence differences that designate functional differences. The above experiments, showing conservation of developmental genetic networks in Drosophilu and mammals, suggest that not only coding regions, but also cis-regulatory elements might be functionally interchangeable (Malicki et al., 1990). Malicki et al. (1992) have demonstrated that at least in some cases this is correct. A 2.8-kb segment from 5’ of the coding sequences of the human Hox-4.2 gene was analyzed for its ability to drive expression patterns in transgenic Drosophila that corresponded to its cognate, the Dfd gene. This segment had previously been shown to direct appropriate Hox-4.2 expression patterns of a reporter gene in the posterior hindbraidcervical spinal cord regions of mouse embryos (Tuggle et al., 1990). This segment and multiple subregions were found to give lac2 expression in the maxillary segments of embryos, closely resembling normal Dfd expression. Immunohistochemistry showed that cells with active lacZ also expressed the Dfd protein. It appeared possible that the human Hox-4.2 control sequencewas actually activated by the Dfd protein itself, as Dfd null mutants showed no maxillary lacZ expression. Alternatively, the appropriate cell types for activating this element may simply fail to form in the absence of Dfd protein. In any event, the regulatory sequences of these human andDrosophiZacognate genes appear to be evolutionarily well conserved in terms of function. It is interesting to note, however, that this functional similarity is not reflected in significant sequence homology. Furthermore, although this experiment worked for HOX-4.2, when performed with a regulatory sequence from Hox-1.3 an appropriate expression pattern (Scr) was not produced (Malicki et al., 1992), suggesting that the conservation of the regulatory network is imperfect, as one might expect. The converse experiment, taking the Drosophila Dfd regulator element and testing it in transgenic mice, has also been reported (Awgulewitsch and Jacobs, 1992). Again the results indicated conservation of function, as the Dfd element directed lac2 expression in the hindbrains of mouse embryos in a manner resembling that of the mouse cognate. Thus the mouse element drives a Drosophila-specific pattern of expression in DrosophiZu, and the Drosophilu element drives appropriate mouse-specific expression in mouse. But it is important to note that the mouse and Drosophilu patterns of expression are very different. While

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Hux-4.2 is active in the neural cells of the hindbrain in the mouse, the Dfd gene is expressed in epidermal cells in Drusuphilu. The clear implication is that the upstream signaling mechanism is evolutionarily conserved, and both Drusuphilu and mouse regulatory elements can respond to these signals, but the spatial distribution of the signal is very different in Drusuphilu and mouse. Hence both elements give the Drusuphilu pattern in Drusuphilu and both elements give the mouse pattern in mouse. The genes upstream of these homeobox genes in the genetic circuitry of mouse and Drusuphilu development have clearly diverged in their expression patterns. This illustrates that although the genetic regulatory network is remarkably similar in mouse and Drusuphilu, there are nevertheless significant differences.

IX. HOMEOBOX GENE CIRCUITRY UPSTREAM AND DOWNSTREAM GENES A. Downstream The homeobox genes represent important nodes in the genetic circuitry of development. They are nevertheless but one set of links in the chain of genetic events, and to understand their function it is necessary to determine h,ow they are regulated and how they in turn regulate downstream genes. The study of the binding of proteins with homeodomains to target sequences has encountered some thorny problems. From structural studies it is clear that homeodomains do assume a helix-turn-helix (HTH) configuration (Otting et al., 1988; Qian et al., 1989), although the homeodomain HTH contacts DNA in a manner different from that of prokaryotic DNA binding proteins (Treisman et al., 1989; Otting et al., 1990; Kissinger et al., 1990; Hanes and Brent, 1991). It is also clear that homeodomain proteins function as regulators of transcription (Biggin and Tjian, 1989; Krasnow et al., 1989; Han et al., 1989; Struhl et al., 1989; Ohkuma et al., 1990).Nevertheless,it has been difficult to discern the mechanism of target gene specificity, as the early DNA binding studies suggested little difference in preferred DNA binding sites for differentAntennupediu-type homeodomains (Treisman et al., 1989;Desplan et al., 1988; Hanes and Brent, 1989). For a time the data indicated that the highly conserved homeodomains encoded by different ho-

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meobox genes could be binding to similar or identical DNA sites, making it difficult to perceive how they obtained functional specificity. More recent studies, however, have found different preferred binding sites for even closely related homeodomains (Ekker et al., 1991; Florence et al., 1991). Dessain et al. (1992), for example, have used full-length proteins and assay their binding to natural target DNA sequences, such as the Drosophila Djb autoregulatory element, and found an excellent correlation between in vitro DNA binding preference and embryonic functional specificity. Some downstream targets of mouse homeobox genes have now been identified. In two cases the homeobox gene itself appears to be a target. Odenwald et al. (1989) reported that the Hox-1.3 protein binds to a sequence 144 bp upstream of the transcription start site of the Hux-1.3 gene. This sequence, identified by DNase I footprinting, was used in gel shift assays along with partially homologous oligonucleotides to determine a consensus DNA binding sequence that had an A'M'AT/G core. This excellent early study of the Hox-1.3 protein did not, however, demonstrate functional significance for the Hox- 1.3 gene binding sequence. Popper1 and Featherstone (1992) used transient expression assays in P 19 embryonal carcinoma cells to demonstrate that the Hox-4.2 protein can autostimulate the Hox-4.2 gene. A block of sequence from -6500 to -393 bp upstream of the Hox-4.2 translational start codon was ligated to a luciferase reporter gene and cotransfected with a Hux-4.2 expression vector carrying the Pgk promoter connected to the Hox-4.2 coding sequences. The presence of the Hox-4.2 expression vector increased luciferase reporter expressionby 20-fold, indicating a strong stimulatory effect. The Hox-4.2 promoter region was then carefully dissected to identify a 217-bp fragment that increased luciferase activity by 13-fold in response to the Hox-4.2 protein expression vector. Inspection of this sequence revealed two TAAT motifs that when mutated reduced activation levels. Oligonucleotides spanning these sequences were shown to bind the Hox-4.2 protein by using electrophoretic mobility shift assays. These experiments demonstrated that in mice, as in Drosophila, the Dfd (or Hox-4.2) gene engages an autostimulatory loop. Such loops are believed to play a role in maintaining the determined state, since once activated they are inclined to stay on. Violette et al. (1992) have reported results implicating homeobox genes in the regulation of the gene encoding the P-amyloid precursor

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protein (APP). The APP gene, which has been associated with Alzheimer's disease, carries 20 TAAT (or A'ITA) core sequences in a 2.2-kb region upstream of the transcription start sites, and seven of these have 8 or 10bases matching the reported Hox-1.3 consensusbinding sequence discussed previously. Violette et al. (1992) therefore performed transfection experiments to determine if homeobox genes could regulate APP gene expression. They first introduced a heat shock promoter-Hox-3.1 construct into HeLa cells and established stable cell lines. It was found that basal levels of Hox-3.1 expression repressedAPP transcription 50% and heat shock induction of Hox-3.1 expression reduced APP transcript levels by 75%. These experiments looked at the endogenous HeLa cell APP gene. Subsequentexperiments used the TAAT-rich upstream region of the APP gene connected to the P-galactosidase reporter, and again increased Hox-3.1 levels were found to repress APP transcription. Electrophoretic mobility shift assays further demonstrated that the Hox-3.1 protein bound to sequences including TAAT comes from the APP upstream sequence. Jones et al. (1992a, 1992b) have also used the cell transfection approach to implicate homeobox genes in the regulation of the cytotactin and neural cell adhesion molecule (N-CAM) genes. Both cytotactin and N-CAM are so-called morphoregulatory molecules that are believed to play a role in cell movement, substrate adhesion, division, and cell-cell communication. Jones et al. (1992a) placed the chicken cytotactin promoter upstream of a CAT reporter and cotransfected into NIH 3T3 cells with the Evx-I or Hox-1.3 coding sequences driven by the CMV or p-actin promoters, respectively. Interestingly, the CMV-Evx-I gene dramatically stimulated the cytotactin promoter, while the P-actin-Hox- 1.3 gene had no effect. Again, the sequences in the cytotactin promoter necessary for this response were dissected out, and footprinting revealed the presence of TAAT cores within the protected region. In a similar set of experiments Jones et al. (1992b) demonstrated that Hox-2.5 protein greatly elevated transcription levels driven by an N-CAM upstream sequence, while the Hox-2.4 protein appeared to have a repressor effect. The above experiments are of considerable interest since they identify downstream targets of Hox genes that may play important roles in morphogenesis and disease. Nevertheless, given the large number of genes in the homeobox family, there is a definite paucity of characterized targets. As the cis-regulatory elements of more downstream genes are

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scrutinizedit is likely that additionalHox gene targets will be found. This is how Olson and colleagues, for example, identified the M-Hox (K-2) gene as a regulator of the muscle creatine kinase gene. But this is an inefficient approach that will only slowly yield small pieces of the genetic network. Another method for target gene identification is to carefully determine consensus DNA binding sequences and then to scan sequence banks for possible downstream genes. The simple TAAT core has already served to identify several possible targets. As PCR approaches are used to determine more extended consensus binding sites for both the clustered and dispersed homeobox genes, one can anticipate that this approach will become increasingly powerful. Mice with “knock out” homeobox genes might also be used to identify targets. First, in situ hybridization can reveal how the absence of one Hox gene influences in vivo expression patterns of other genes. This is a laborious approach, however, and one needs to carefully distinguish primary targets from secondary and tertiary downstream targets, but the strength of this method is that it determines in vivo effects and is therefore relatively artifact free. In addition, mice with a null homeobox gene mutation might serve as sources of cell lines that could be used to make subtraction libraries for the mass selection of gene targets. To facilitate establishment of the cell lines one could introduce a transgene with the targeted Hox gene promoter driving the SV40 large T antigen. This would provide large T expression in cells that normally express the homeobox gene and assist in transforming them, enabling cell line formation, and would assist appropriate cell identification, since cells expressing the large T antigen would presumably be cells that normally express the ablated Hox gene. These cell lines could then be transfected with an inducible promoter driving expression of the coding sequences of the targeted €€ox gene, providing a clean system for generating cells that do and do not express. Subtractionlibraries could then provide access to the normal targets of the Hox gene. B. Upstream Regulation Homeobox genes regulate expression patterns of batteries of target genes and thereby control the developmentaldestinies of groups of cells. But what factors regulate expression of the homeobox genes? Part of the answer is that homeobox genes form a regulatory network that controls

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homeobox gene expression patterns. For some genes there is an autoregulatory mechanism that maintains expression once activated (Popper1 and Featherstone, 1992). And a number of studies have now shown that mammalian homeobox genes can cross-regulate each other. Zappavigna et al. (1991) searched for possible homeoprotein binding sites in the human HOX 4 0 - H O X 4C intergenic region by performing sequence comparisons between mouse, human, and chicken. Such comparisons showed remarkable conservation for human and mouse, even in this intergenic region, and a portion of it was also nearly identical in the chicken. In virro binding assays showed this region to have multiple sites that specificallybound proteins encoded by the human H O X 4 C and HOX 4 0 and mouse Hox-4.3 genes. These binding sites, although distinct, all carried the TAAT core. This region, designated HCR for Hox cross-talk region, could mediate transactivation of a reporter gene when cotransfected with a H O X 4C expression plasmid, suggesting an autoregulatory circuit for HOX 4C. Moreover, the product of HOX 4 0 could also transactivate the HCR reporter gene in cotransfection, suggesting that this neighboring homeobox gene may also modulate HOX 4C expression. Another homeodomain protein, however, that encoded by the murine Hox-4.3 gene, did not activate the HCR, showing that not all homeobox proteins bind and activate equally. Moreover, the Hox4.3 protein had a repressor effect on the activation of the HCR by HOX 4C or HOX 4 0 , and this effect was independent of Hox-4.3 binding to the HCR, since Hox 4 . 3 protein with a deleted homeodomain repressed with equal efficiency. These results suggest protein-protein interactions between the Hox-4.3 and H O X 4C and H O X 4 0 products that could mask, for example, transcription activation domains. The human HOX 30 upstream region was characterized in a similar manner by Arcioni et al. (1992). Again, cross-regulation was examined by cotransfection experiments in cultured cells. Interestingly, proteins encoded by HOX 3C, HOX 4 0 , HOX 4C, and Hox-4.3 (the mouse cognate of HOX 4 3 ) were all able to transactivate the HOX 30 promoter in both human HeLa and mouse NIH 3T3 cells. The authors pointed out that all of these genes are from more 5 ’positions on the homeobox gene clusters, which means they are expressed in more posterior positions at later times in development. But the HOX 3 D protein itself, and those encoded by genes more 3’ on the clusters (HOX 3E and HOX 4B), had

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no activating effects. They interpreted these results to suggest a directional component to cross-regulatory interactions, with genes expressed in more posterior positions able to activate more “anterior”ones. Retinoic acid appears to be another upstream regulator of Hox gene expression. Stornaiuolo et al. (1990) first demonstrated that homeobox genes in NT2PDL embryonal carcinoma cells could be induced by culturing in 10 ph4 retinoic acid. It was observed that 22 homeobox genes, all from the 3’ regions of the clusters, were activated, while 11 monitored homeobox genes from the 5’ ends of the clusters remained silent. In subsequent studies Simeone et al. (1990) found that genes in the 3’ half of the H O X 2 cluster were activated in NI2/D1 cells by 0.01 ph4 retinoic acid, while more 5’ genes required increasing doses of retinoic acid to be activated. A subsequent and more thorough analysis of responses of homeobox genes of all four clusters to varying levels of retinoic acid showed that for each cluster the 3’ genes were activated by low doses, the more 5’ genes were activated by higher retinoic acid doses, and the most 5’ genes were actually slightly down-regulated by retinoic acid. Kessel and Gruss (1991) were able to treat mouse embryos with retinoic acid and induce anterior shifts in Hox gene expression domains that were accompanied by anterior transformations of vertebrae. The altered combination of Hox genes expressed apparently resulted in altered specification of segment identity, supporting the “Hox Code” hypothesis. Marshall et al. (1992) also identified Hox expression pattern changes and segment transformations resulting from exposureto retinoic acid. Their experiments analyzed expression of Hox-B (Hox-2) cluster genes in the hindbrain. In particular, transgenic mice with Hox-BI and Hox-B2 promoters connected to the ZucZ reporter were used to monitor expression of these genes in response to retinoic acid. Again, changes in Hox gene expression were associated with transformation of segment identity, with rhombomere 2/3 assuming a rhombomere 41.5 identity. These experiments, showing that Hox genes can be activated by the putative morphogen retinoic acid, suggest one mechanism by which a specific concentration of morphogen, designating a specific position in a gradient, could be converted into a particular combination of homeobox gene expression, which in turn would determine cell identity.

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X. &-REGULATORY ELEMENTS Sham et al. (1993) in another study of hindbrain development have found that a zinc fiiger gene, Krox 20, plays a role in activating rhombomerespecific expression of the Hox B2 (Hox 2.8) homeobox gene. The Krox 20 gene is expressed in rhombomeres 3 and 5, and Hox B2 is in rhombomeres 3, 4, and 5. In making transgenic mice with upstream sequences from Hox B2 connected to lacZ it was found that reporter expression in the hindbrain first appeared in rhombomeres 3 and 5 and then extended to other regions. This coincidencewith Krox20 expression suggested a role for Krox 20 in regulating Hox B2. Indeed, a number of Krox 20 binding sites were found in this upstream segment by gel shift assays followed by DNase I footprint analysis. These sites were shown to be essential for appropriate rhombomere 3 and 5 expression. Moreover, ectopic expression of Krox 20 in transgenics was able to transactivate the Hox B2 enhancer, strongly suggesting a normal role for Krox 20 in regulating Hox B2 expression. It therefore appears that at least in some cases zinc finger genes are located upstream of homeobox genes in the genetic circuitry of mammalian development. This perhaps should not be surprising, since in Drosophila the zinc finger gap genes hunchback, Kruppel, and Knirps are upstream of the homeotic homeobox genes. It is interesting to note, however, that bicoid, a maternal effect homeobox gene, is upstream of the gap genes. It is clear that the level in the genetic hierarchy at which a transcription factor gene will operate cannot be entirely predicted by the gene family it belongs to. The example described above is a striking success story where the analysis of cis-regulatory sequences resulted in the identification of an upstream tissue specific transcription factor involved in regulation of the homeobox gene. In general this goal has been difficult to achieve. Indeed, there is reason to conclude that the dispersed homeobox genes, outside the clusters, may be particularly good candidatesfor further gene expression studies. The data suggest that most of the Hox cluster genes might share regulatory elements, making it particularly difficult to dissect their control. The strong evolutionaryconservation of the cluster organization argues that it may be functionally significant. Furthermore, the colinearity of the clusters, with all genes transcribed in the same direction and with genes more 3’ expressed earlier and in more anterior positions and more sensitive to retinoic acid, all suggest control as a unit, much like the beta globin gene cluster. In addition, most efforts to identify cis-

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regulatory sequences of cluster genes have only succeeded in reproducing subsets of normal expression patterns in transgenic mice (Zakany et al., 1988; Puschel et al., 1990; Tuggle et al., 1990; Kress et al., 1990; Jegalian et al., 1992). Indeed, it has been reported that extensive efforts to reconstruct normal expression patterns in transgenics with Hox cluster genes have generally failed (Whiting et al., 1991). Of course, on the positive side, it is extremely important to identify the enhancers of the homeobox gene clusters, and several of the above studies were able to re-create significant portions of the normal expression pattern in transgenics. Moreover, in a few cases it has been possible to obtain expression patterns that closely mimic the endogenous genes (Whiting et al., 1991; Bieberich et al., 1990; Puschel et al., 1991). Nevertheless, for dispersed homeobox genes the control elements are more likely within the vicinity of the gene and not spread throughout a cluster. Understanding their cis-regulatory elements may therefore be a simpler task. In searching for regulatory elements it is clear that particular attention must be given to intron sequences. The homeobox gene intron sequences are often well conserved, with the mouse and human Hox-4.4 introns and the mouse and human Hox-1.3 introns being almost identical (Renucci et al., 1992; Tournier-Lasserve et al., 1989). The use of transgenic animals to dissect cis-regulatory elements has serious disadvantages; chief among them are time and cost. Generally, multiple transgenic lines must be made for each DNA construct tested, to average out variations in expression due to the random chromosome insertion sites. F l animals are usually examined instead of the original transgenics so that a number of individuals from each line can be tested, and to avoid mosaicism problems associated with FO mice. This means that a few months are needed to examine each construct, and hence the “turnaround” time is slow. That is, when one is defining the limits of cis-regulatory sequences one needs the results from one series of constructs before proceeding to the next. Despite these problems there are also considerable advantagesto using transgenic mice. First and foremost, it represents an in uivo situation, with all cell types and all developmental stages available for analysis of gene expression. The results obtained represent what is happening in living, developing mice and are therefore relatively artifact free. Although cell culture systems are faster and easier, when expressing cells are available, it is important to note that control of gene expression is a complex multifactorial process generally involving a number of cisregulatory elements and interacting transcription factors. For each ex-

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pressing tissue a different combination of cis-elements and trans-factors are involved, so at best a cell line is appropriate for understanding gene expression in one tissue only. The problems of slowness can be partly overcome by using transient transgenics. With this approach the FO mice are sacrificed at some stage in development to X-gal stain and examined for transgene expression (when using the lacZ reporter). The great advantage is that expression results are obtained 2-3 weeks after microinjection. It therefore greatly cuts the turnaround time and allows the analysis to proceed with some rapidity. The disadvantages are that a significant fraction of FO mice are mosaic, and with this approach only one time point can be analyzed for each transgenic. It is therefore more brute force at the microinjectionend of the experiments, requiring a larger number of FO transgenic mice. In any event, by using both transgenic mice and cell culture systems, the work of defining the regulatory elements of the Hox genes is moving ahead rapidly (Renucci et al., 1992; Galliot et al., 1989; Z w a r t h i s et al., 1991). Enhancers that at least specify subsets of normal expression patterns are being identified, and gel shift assays and DNase I footprints are defining precise sequences that interact with upstream transcription factors. And at least in a few cases the upstream genes encoding these proteins regulating Hox gene expression are being characterized.

XI. CONCLUSIONS Within the DNA of the zygote is encoded a genetic program that drives appropriate temporal and spatial patterns of gene expression during development. It is becoming abundantly clear that homeobox genes will play a central role in this process. In some respects the job of today’s scientists attempting to understand the genetic regulatory network of mammalian development resembles that of the biochemists of decades past that worked out the pathways of intermediary metabolism. Indeed, in time it may become possible to generate a wall chart of mammalian developmental genetic interactions that parallels the current wall chart of intermediary metabolism. At prominent positions on the developmental wall chart one will find homeobox genes serving as master switches that control the developmental destinies of groups of cells. Their gene products will directly influence expression levels of batteries of target genes, at least some of which will be other homeobox genes. It is clear from work with Drosophila that homeobox genes can sometimes initiate

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genetic cascades that determine the identities of developing structures. The final picture of mammalian developmental genetic interactions that will in time emerge will be quite complex, in some ways more resembling a neural network than a simple diagram of intermediary metabolism because of its three-dimensional rather than linear nature. That is, each gene can be influenced by multiple upstream inputs and each transcription factor gene can in turn modulate expression patterns of a number of downstream targets. Nevertheless, our eventual understanding of the molecular genetic interactions involved in generating specific structures of the mammal will be deeply satisfying and will serve as a basis for our understanding of inherent as well as environmentally induced developmental defects in children.

ACKNOWLEDGMENTS Thanks to the members of the laboratory for helpful discussions and thanks to Jan Hagedorn for excellent preparation of the manuscript. Support has been provided by NIH HD2599-01.

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EXPRESSION AND FUNCTION OF C - ~ O SIN MAMMALIAN GERM CELLS

Geoffrey M. Cooper

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Introduction.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Tissue Specificity of c-mos Expression . . . . . . . . . . . . . . . . . . . . 111. Expression and Function of c-mos in Oocytes . . . . . . . . . . . . . . . . Iv. Expression and Potential Function of c-mos in Spennatocytes . . . . . . . V. Regulation of c-mos Transcriptionin Mouse Oocytes . . . . . . . . . . . . VI. Negative Regulation of c-mos Transcription in Somatic Cells . . . . . . . . VII. Summary and Future Directions . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . , . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

127 128 129 131 136 137 140 143 144 144

PREFACE The c-mos proto-oncogene is unique in being specifically expressed in male and female germ cells, where it appears to play a central role in regulating the meiotic cell cycle. Its function is best understood in

Advances in Developmental Biochemistry Volume 3, pages 127-148. Copyright 0 1994 by JAI Press InC. All rights of reproduction in MY form reserved. ISBN: 1-55938-865-X

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oocytes, where it has been studied by microinjection of antisense oligonucleotides to ablate maternal stores of c-mos M A . Such experiments in both Xenopus and mouse oocytes have shown that the c-mos gene product (Mos) is required to maintain maturation promoting factor (MPF) activity by stabilizing cyclin B during progression from meiosis I to meiosis 11, as well as acting as a component of cytostatic factor to maintain subsequent metaphase I1 arrest. In Xenopus, c-mos translation is also required for initiation of meiosis I. This is not the case in the mouse, but it is possible that Mos protein is already present in mouse oocytes and functions at this stage. In spermatocytes, c-mos appears to be expressed prior to meiosis, which is consistent with the possibility that it may also function in the initiation of meiosis and progression from meiosis I to meiosis 11, although it clearly does not cause metaphase I1 arrest in male germ cells. In somatic cells, c-mos is either silent or possibly expressed at very low levels, indicating that expression of this proto-oncogeneis subjectto stringent tissue-specificregulation. We have identified two distinct regulatory elements that activate c-mos transcription in oocytes and repress its transcription in somatic cells, respectively. Because aberrant c-mos transcription is sufficient to result in either cell death or transformation, repression of c-mos in somatic cells may constitute a particularly important regulatory mechanism.

1. INTRODUCTION The c-mos gene was first discovered as the normal cell homolog of the oncogene (v-mos) of Moloney sarcoma virus, a highly oncogenic retrovirus that induces sarcomas in infected mice. Like other retroviral oncogenes, v-mos originatedby the incorporationof c-mos into a retroviral genome in such a way that its resulting aberrant expression led to abnormal proliferation and neoplastic transformation of virus-infected cells. In this respect, c-mos is similar to the 70 or so other proto-oncogenes that have been identified in mammalian genomes. However, the normal function of c-mos differs considerably from those of the many other proto-oncogenes that have been studied. In particular, c-mos appears to play a unique role in germ cells, so studies of its expression and function offer the promise of novel insights into the mechanisms that control germ cell development and the meiotic cell cycle. The proteins encoded by most proto-oncogenesare normally expressed in a variety of differentiated cell types, where they generally function as

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components of signaling pathways that regulate cell growth and differentiation. The majority of these genes encode growth factors, growth factor receptors, and proteins involved in intracellular signal transduction (protein-tyrosine and protein-serinekhreonine kinases, guanine nucleotide-binding proteins, and transcription factors). The c-mos gene also encodes a protein-serinekhreonine kinase (Mos). However, it displays a highly restricted pattern of specific expression in male and female germ cells, where it appears to function during meiosis, rather than as an intracellular signaling molecule involved in the control of somatic cell proliferation.Both the regulation and function of c-mos thus pose important issues with respect to understanding the molecular mechanisms that control mammalian development.

II. TISSUE SPECIFICITY OF c-mos EXPRESSION The first insights into the function of c-mos were provided by understanding its highly restricted pattern of tissue-specific expression. In contrast to other proto-oncogenes, which are generally expressed in a range of different cell types, early studies failed to detect c-mos transcription in a variety of tissues and cell lines. The first indication of normal c-mos activity was thus provided by detection of c-mos transcripts in testes and ovaries of adult mice (Propst and Vande Woude, 1985). Subsequent studies established that c-mos was specifically transcribed in both male and female mouse germ cells (Goldman et al., 1987; Mutter and Wolgemuth, 1987; Propst et al., 1987). Extensions of these findings have similarly documented expression of c-mos in gonads and germ cells of several other species, including primate ovaries and testes (Paules et al., 1988), chicken ovaries and testes (Schmidt et al., 1988), Xenopus oocytes (Sagata et al., 1988), rat spermatocytes (Van der Hoorn et al., 1991), and human oocytes (Pal et al., 1994). It thus appears that c-mos is generally expressed in reproductive tissues, in particular in both male and female germ cells, of a variety of vertebrate species. This germ cell-specific expression provided the initial suggestion of the role of c-mos as a regulator of the meiotic cell cycle. Although testicular and ovarian germ cells clearly represent the principal sites of c-mos transcription, lower levels of c-mos expression have been reported in some somatic tissues and cell lines. Propst et al. (1985, 1987) reported low levels of c-mos transcripts in mouse embryos and

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somatic tissues of adult mice, although these transcripts were not further characterized. Herzog et al. (1989) have also reported detection of Mos protein in a variety of somatic cell types, including N M 3T3 mouse fibroblasts, and Leibovitch et al. (1990) have reported c-mos transcripts and protein in rat muscle cells. However, Paules et al. (1992) failed to confirm detection of Mos protein in NIH 3T3 cells, nor were these investigators able to detect c-mos RNA by PCR amplification in either NIH 3T3 cells or mouse skeletal muscle. It is possible that the divergent results of these groups, at least at the protein level, are due to differences in antisera, which may lead to the detection of cross-reactive proteins in addition to authentic Mos. At present, the issue of whether c-mos is expressed and functions in somatic cells remains an area of some controversy. Whether or not there is some expression of c-mos in somatic cells, it is clear that germ cells represent the major sites of c-mos transcription. It is interesting to note that the consequences of aberrant c-mos expression suggest that suppressing inappropriate c-mos expression in somatic cells may be a particularly important aspect of c-mos regulation. High levels of Mos are toxic to fibroblasts (Papkoff et al., 1982) possibly because Mos induces metaphase arrest (Sagata et al., 1989) (see below). Furthermore, only low levels of expression of c-mos are sufficient to induce cell transformation (Papkoff et al., 1982; Wood et al., 1983). In this respect, c-mos differs from other proto-oncogenes encoding protein kinases, which require structural mutations for their conversion to oncogenes. For example, the c-ruf proto-oncogenes (which also encode protein-serinelthreonine kinases) require deletions or mutations of an amino-terminalregulatory domain for oncogenic activity (Stanton et al., 1989; Heidecker et al., 1990). In contrast, the activation of c-mos as an oncogene only requires unregulated expression of the normal c-mos protein (Blair et al., 1981). It is noteworthy in this regard that Mos is a relatively small protein (343 amino acids) that encompasses little more than the protein kinase catalytic domain (-300 amino acids) (Blair et al., 1984; Hanks et al., 1988). The absence of an apparent Mos regulatory domain suggests the possibility that much of the regulation of c-mos may be at the level of gene expression, rather than control of Mos kinase activity. The drastic consequences of even low levels of aberrant c-mos expression in somatic cells emphasize the importance of transcriptional regulation of this proto-oncogene, as discussed further below.

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111. EXPRESSION AND FUNCTION OF C - ~ O SIN OOCYTES In mouse oocytes, c-mos is transcribed during oocyte growth and high levels of c-mos transcripts (estimated on the order of 100,000molecules per oocyte) accumulate in fully grown germinal vesicle stage oocytes arrested at the diplotene stage of meiotic prophase (Goldman et al., 1987; Mutter and Wolgemuth, 1987) (Fig. 1). The c-mos transcripts in these oocytes lack detectable poly(A) tails but become polyadenylated following resumption of meiosis (Goldman et al., 1988; Mutter et al., 1988). Like other maternal mRNAs in the mouse, c-mos mRNAis subsequently degraded by the two-cell stage of embryo development, when transcription of the embryonic genome is initiated (Goldman et al., 1988; Mutter et al., 1988; Keshet et al., 1988). In Xenopus oocytes, c-mos mRNA is similarly polyadenylated following resumption of meiosis, although (like other Xenopus maternal RNAs) it is not degraded until the blastula stage (Sagata et al., 1988). The polyadenylation of c-mos mRNA in maturing oocytes is similar to that of other maternal RNAs that are accumulated during oocyte

Oocyte growth

c-rnos transcription

Fertilization and cleavage

c-mos d e g r a d a t i o n

Figure 7. Expression of c-mos in mouse oocytes. c-mos is transcribed during oocyte growth and transcripts with short poly(A) tails are accumulated in fully-grown germinal vesicle (GV) stage oocytes. These transcripts are polyadenylated and translated following the resumption of meiosis and then degraded following fertilization and cleavage to the two-cell stage.

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growth and subsequently translated following the resumption of meiosis. Stored maternal mRNAs in both Xenopus and mouse oocytes contain short poly(A) tails of 30-40 nucleotides that are elongated by up to several hundred nucleotides during meiotic maturation, and it has recently been shown that such cytoplasmic polyadenylation in fact controls translation of these maternal mRNAs (McGrew et al., 1989; Vassalli et al., 1989). Consistent with this, c-mosmRNAis translated following resumption of meiosis inbothxenopus and mouse oocytes (Sagataet al., 1988;Paules et al., 1989), although Paules et al. (1989) have also reported a low level of Mos synthesis in mouse oocytes arrested at the germinal vesicle stage. The behavior of c-mos mRNA as a maternal message recruited for translation during maturation of mouse oocytes suggested the potential function of Mos in meiosis, fertilization, or the initial cleavage of fertilized eggs to two-cell embryos. The function of c-mos in meiosis of both Xenopus and mouse oocytes was initially demonstrated using microinjection of antisense oligonucleotidesto ablate the maternal pool of c-mos mRNA (Sagata et al., 1988; O’Keefe et al., 1989; Paules et al., 1989). In Xenopus, Mos synthesis was required for initiation of meiosis I, since oocytes injected with antisense c-mus oligonucleotides failed to undergo germinal vesicle breakdown (GVBD) in response to hormonal stimulation (Sagataet al., 1988).In contrast, mouse oocytes injected with antisense c-mos oligonucleotides underwent GVBD but failed to progress normally through meiotic maturation (O’Keefe et al., 1989; Paules et al., 1989). In particular, antisense c-mos injected oocytes completed meiosis I but failed to initiate meiosis 11, instead reforming a nucleus containing decondensed chromatin within a few hours following polar body extrusion (O’Keefe et al., 1989) (Fig. 2). Although different, the effects of ablating c-mos mRNA coincide with the requirements for protein synthesis during meiosis in Xenopus and mouse oocytes. In Xenopus, but not in mouse, protein synthesis is required for GVBD, whereas protein synthesis is required for progression from meiosis I to meiosis I1 in both species (Wassennan and Masui, 1975;Siracusaet al., 1978;Clarke andMasui, 1983;Gerhartet al., 1984). The lack of requirement for c-mos translation for GVBD in the mouse may suggest that Mos protein is required for GVBD in Xenopus but not mouse oocytes. Alternatively, mouse (but not Xenopus) oocytes might contain some Mos protein synthesized prior to resumption of meiosis. The detection of a low level of Mos protein in germinal vesicle stage oocytes (Paules et al., 1989) is consistent with the latter possibility. The synthesis of Mos protein in mouse oocytes prior to resumption of meiosis

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Figure 2. Role of c-mos in meiosis of mouse oocytes. Mouse ooc:ytes microinjected with antisense c-mos oligonucleotides'fail to progress from meiosis I to meiosis 11, instead reforming nuclei following polar body extrusion (O'Keefe et al., 1989). has been confirmed by Zhao et al. (1990, 1991), who have further reported that microinjection of mouse oocytes with anti-Mos antibody blocks GVBD. Although these results indicate arole for Mos in initiation of meiosis in mouse as well as Xenopus oocytes, it must be noted that the antibody used by Zhao et al. (1990, 1991) also detects Mos protein in somatic cells, including NM 3T3 fibroblasts. As discussed above, the failure to confirm these results with other antibodies or to detect c-mos transcription in NM 3T3 cells with RT-PCR (Paules et al., 1992) raises the possibility of antibody cross-reaction with cell proteins other than authentic Mos. Thus, questions concerning the possible translation of c-mos in germinal vesicle stage mouse oocytes, and the function of the Mos protein kinase during initiation of meiosis I in the mouse, have not yet been unambiguously resolved. That Mos is required for progression from meiosis I to meiosis I1 in Xenopus as well as mouse has been demonstrated by allowing Xenopus

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oocytes to initiate meiosis I prior to injection of antisense c-mos oligonucleotides (Kanki and Donoghue, 1991). Under these conditionsXenopus oocytes complete meiosis I but fail to progress to meiosis 11, similar to the results obtained in the mouse (O’Keefe et al., 1989). Thus, although the role of Mos in initiation of meiosis I in the mouse is unclear, studies in both Xenopus and the mouse indicate that Mos is required for normal progression of oocytes from meiosis I to meiosis 11. Further studies in both Xenopus and mouse oocytes have related the activity of Mos to the cell cycle regulator, maturation promoting factor (MPF). MPF, originally identified in amphibian eggs, is a proteinserine/threonine kinase composed of catalytic and regulatory subunits, cdc2 and cyclin B, respectively (Kirschner et al., 1992; Murray, 1992). Its activation induces chromosome condensation, nuclear membrane breakdown, and the cytoplasmic reorganization associated with entry into M phase of either mitosis or meiosis. The activity of the cdc2 kinase is controlled both by association with cyclin B and by phosphorylation. Entry of cells into the M phase of either mitosis or meiosis is induced by dephosphorylation of cdc2 complexed to cyclin B. During mitosis, inactivation of MPF by the proteolytic destruction of cyclin B then signals the metaphaseto anaphasetransition (Draettaet al., 1989;Murray et al., 1989), which is followed by chromosome decondensation,reformation of anucleus, and cytokinesis. In contrast, oocyte meiosis involves two successive M phases without reformation of a nucleus, and vertebrate eggs remain arrested at metaphase I1 for up to several days prior to fertilization, suggesting that distinct mechanisms regulate MPF during oocyte meiosis. Studies of both Xenopus and mouse eggs have indicated that MPF activity increases upon the resumption of meiosis and remains high until the metaphase/anaphasetransition of meiosis I, when it declines to low levels (Gerhart et al., 1984; Hashimoto and Kishimoto, 1988) (Fig. 3). Following polar body extrusion, however, MPF activity again increases and remains high while the egg is arrested at metaphase 11. The maintenance of high levels of MPF activity involves an additional factor first detected in amphibian eggs (cytostatic factor or CSF), which appears to stabilize MPF during metaphase I1 arrest. Mos has been identified as a component of CSF in Xenopus and thus appears to be involved in maintaining MPF and holding the egg at metaphase I1 until fertilization (Sagata et al., 1989). Mos is also required for maintenance of MPF activity during progression of mouse eggs from meiosis I to meiosis 11, apparently acting to stabilize cyclin B and allow the increase in MPF

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Mos Required

I

GVBD

Polar body extrusion

Metaphase I I

arrest

figure 3. Activity of maturation promoting factor (MPF) during oocyte meiosis. MPF activity increases upon the resumption of meiosis, leading to germinal vesicle breakdown (GVBD). Its activity remains high until the metaphase/anaphase transition of meiosis I, when it declines to low levels. Following completion of meiosis I, indicated by polar body extrusion, MPF activity again increases and remains high while the egg is arrested at metaphase 11. In both Xenopus and mouse, newly synthesized Mos is required for maintenanceof MPF activity during progressionfrom meiosis I to meiosis 11. In Xenopus, but not in mouse, new Mos synthesis is also required for GVBD, although it is possible that preexisting Mos protein is active during GVBD in the mouse.

activity required for progression to meiosis I1 (O’Keefe et al., 1991). It thus appears that Mos contributes to the unique regulation of the meiosis of vertebrate oocytes by inhibiting the degradation of cyclin B that would otherwise occur after the metaphase/anaphase transition of meiosis I, thereby allowing progression to meiosis I1 and maintaining metaphase I1 arrest. The biochemical mechanism of Mos action is not yet established. Mos has been found to phosphorylate cyclin B in v i m , and it is possible that this phosphorylation directly inhibits cyclin B proteolysis (Roy et al., 1990). However, such a direct effect of phosphorylation on cyclin B stability remains to be demonstrated, and it is alternatively possible that Mos inhibits (directly or indirectly) the proteolytic pathway responsible for cyclin B degradation. Mos has recently been found to stimulate mitogen-activated protein kinase (MAP kinase) in Xenopus oocytes,

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suggesting the possibility that MAP kinase may mediate Mos activity (Posada et al., 1993; Nebreda and Hunt, 1993). Also unresolved is the relationship between the function of Mos as a regulator of meiosis in oocytes and its activities in somatic cells. As noted above, high levels of Mos expression in fibroblasts are cytotoxic (Papkoff et al., 1982). It is likely that this cytotoxicity is a manifestation of the CSF activity of Mos, leading to the M phase arrest of mitotically proliferating cells (Sagataet al., 1989;Okazaki et al., 1992). On the other hand, cell transformation resulting from lower levels of Mos expression is exerted in the GI phase of the cell cycle (Okazaki et al., 1992), so the relationship between the transforming activity of Mos and its normal function in regulating the M phase of oocyte meiosis is unclear. One possibility is that transforming activity of Mos might result from activation of MAP kinase (Posada et al., 1993;Nebreda and Hunt, 1993). MAP kinase represents a common target of oncogenes (e.g., Ras and Raf) involved in intracellular signaling pathways leading to somatic cell proliferation and is believed to play acentral role in conveying mitogenic signals from growth factor receptors to the nucleus (Wood et al., 1992; Thomas et al., 1992; de Vries-Smits et al., 1992; Dent et al., 1992; Kyriakis et al., 1992;Howe et al., 1992).It is thus an attractive possibility that a low level of Mos, insufficient to induce metaphase arrest, leads to cell transformation via activation of MAP kinase signaling pathways.

IV. EXPRESSION AND POTENTIAL FUNCTION OF C - ~ O SIN SPERMATOCYTES While the general role of Mos in oocyte meiosis now appears to be established, its potential activity in male germ cells is less clear. In the mouse, the highest levels of c-mos RNAare detected in round spermatids, which represent the earliest haploid postmeiotic cell type, although lower levels of c-mos mRNA may be present in pachytene spermatocytes, which have not yet undergone meiosis (Goldman et al., 1987;Mutter and Wolgemuth, 1987; Propst et al., 1987). Mos protein has also been reported in both mouse pachytene spermatocytes and round spermatids, as well as in later postmeiotic stages of spermiogenesis (Herzog et al., 1989). In the rat, c-mos RNAis present in both pachytene spermatocytes and postmeiotic spermatids,with the highest levels of Mos protein found at the pachytene stage (Van der Hoorn et al., 1991).

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Although the studies of c-mos protein are subject to caveats related to concerns about antibody specificity (as discussed above), it appears most likely that c-mos is expressed prior to spermatocyte meiosis, consistent with its possible function during meiosis of spermatocytes as well as oocytes. It must be recognized, however, that there are substantial differences in the meiotic cycles of these two cell types-in particular, spermatocytes progress rapidly through two meiotic divisions without arresting at metaphase 11. Thus, while Mos may be involved in the initiation of spermatocyte meiosis and in progression from meiosis I to meiosis 11, it clearly does not induce metaphase I1 arrest. Indeed, postmeiotic round spermatids have reformed a nucleus even though they contain high levels of c-mos mRNA and protein, indicating that additional regulatory mechanisms must be involved. It should be noted that Mos is not the only protein required for progression ofXenopus oocytes from meiosis I to meiosis I1 (Yew et al., 1992). Moreover, it has recently been shown that the cdc2-related protein kinase cdk2 is required in addition to Mos for metaphase I1 arrest in Xenopus (Gabrielli et al., 1993). It is thus possible that an absence of cdk2 could be responsible for the ability of male germ cells to progress through metaphase I1 even in the presence of Mos. It is also interesting to note that cyclin B mRNA (like c-mos) is found at high levels in postmeiotic male germ cells (Chapman and Wolgemuth, 1992), suggesting the possibility that Mos and cyclin B may play some role in spermiogenesis other than cell cycle regulation. Unfortunately, in vitro culture systems that support meiosis and differentiation of male germ cells are not available, so functional studies of Mos in spermatocytes have not been possible.

V. REGULATION OF C - ~ O STRANSCRIPTION IN MOUSE OOCYTES The highly restricted tissue-specific transcription of c-mos poses an interesting problem in gene regulation during germ cell development. Moreover, c-mos is transcribed from different promoters in mouse spermatocytes and oocytes (Fig. 4) (Propst et al., 1987). In spermatocytes, transcription initiates approximately 280 nucleotides upstream of the c-mos ATG (Propst et al., 1987), whereas the transcription start site in oocytes has been mapped to 53 base pairs upstream of the ATG (Pal et al., 1991). Neither the spermatocyte nor oocyte promoter regions are

NRE

llnr

I

I ATG

Figure 4. Transcriptional regulatory sequences of the mouse c-mos gene. Transcription in oocytes and spermatocytes initiates 53 and approximately 280 base pairs upstream of the c-mos ATG, respectively. Efficient transcription in oocytes requiresan initiator (Inr)-likesequence located downstream of the transcription start site. A negative regulatory element (NRE) located upstream of the spermatocyte promoter acts to suppress c-mos transcription in somatic cells.

associated with TATA, CCAAT, or GC boxes, so it has been of interest to determine the regulatory elements involved in tissue-specific c-mos transcription. Our initial studies of c-mos regulation employed transient expression assays to identify sequences required for c-mos transcription in mouse oocytes (Pal et al., 1991). Microinjection of growing oocytes with c-moslCAT reporter constructs indicated that the c-rnos 5’ flanking sequences promoted transcription with an efficiency similarto that of the SV40 early promoter. Surprisingly, deletion analysis revealed that 5’ flanking sequences up to 20 base pairs upstream of the major transcription start site could be deleted without any significant reduction of promoter activity. On the other hand, sequences within 20 nucleotides downstream of the transcription initiation site were required for efficient expression in microinjected oocytes. These sequences included an element with the pyrimidine-rich consensus sequence of initiator (Inr) elements, first defined in the gene encoding terminal deoxynucleotidyltransferase (Smale and Baltimore, 1989). Point mutations within or immediately adjacent to this Inr-like sequence dramatically reduced c-mos promoter activity in oocytes, confirming its functional importance. It thus appears that efficient c-mos transcription in mouse oocytes requires only a simple promoter, consisting of sequences immediately surroundingthe transcription start site and a downstream Inr-like element (Fig. 5). It should be noted that not all Inr-like elements overlap transcription start sites; downstream Inr-like elements in other genes have also been shown to be required for efficient transcription (Nakatani et al., 1990. The lack of requirement for enhancers in order to achieve efficient c-mos transcription in oocytes is consistent with previous studies indi-

c-mos Expression and Function

GATGA

139

C T C AT T T C T

Figure 5. The oocyte c-mos promoter. Efficient transcription in oocytes requires only a minimal promoter fragment, extending from -21 to +20. This fragment includes an Inr-like element, which is required for efficient gene expression. A potential binding site for the oocyte-specific transcription factor OSP-1 is located further upstream.

cating that the SV40 and thymidine kinase promoters are transcribed efficiently in oocytes without enhancer sequences(Chalifour et al., 1987; Martinez-Salas et al., 1989).The only other oocyte-specificgenes whose transcription has been studied are those encoding the zona pellucida (ZP) proteins which, like c-mos, are transcribed at high levels in growing oocytes. Unlike c-mos, the ZP promoters contain TATA boxes, and upstream elements are required for expression in oocytes (Lira et al., 1990; Millar et al., 1991; Schickler et al., 1992). Analysis of mouse ZP2 and ZP3 promoters has further identified an element located approximately 200 base pairs upstream of the TATA box, which is both necessary and sufficient for efficient expression in oocytes (Millar et al., 1991). This element contains an E box consensus binding site (CACGTG), which is recognized by members of the helix-loop-helix family of transcriptional activators. Members of this group of transcription factors, which includes immunoglobulin enhancer binding proteins, Myc, MyoD, and USF, regulate the expression of a variety of genes during development and differentiation(Murre and Baltimore, 1992).Although the E box sequence of the ZP promoters does not appear to be directly related to the Inr-related element of c-mos, it is important to note that a USF-related factor, TFII-I, specifically binds to Inr elements (Roy et al., 1991). Interestingly, TFII-I binds with similar affinities to both Inr and E box sequences. USF also binds to Inr sequences, although with lower affinity than to the E box. In addition, TFII-I and USF interact cooperatively at both the E box and Inr sites. The structural basis for the ability of USF and TFII-I to bind to two different sequence elements is not yet understood, although competition experiments suggest that TFII-I binds to E box and Inr sites through distinct DNA binding domains (Roy et al.,

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1991).In any case, the ability of USF and TFII-I to recognize both E box and Inr-like elements clearly has interesting implicationsfor the potential role of a member of this transcription factor family in oocyte-specific gene expression. Namely, it is possible that a USF-related factor could be involved in oocyte-specificexpression of both the c-mos and ZP genes via interaction with Inr and E box elements, respectively. Identification and characterization of the oocyte factors that bind to these sequences will obviously be needed to test this possibility. An additional sequence element within the mouse ZP3 promoter has been identified as the binding site of an oocyte-specific protein called OSP-1 (Schickler et al., 1992),although the possible effect of mutations of this element on transcriptional activity has not been established. Nonetheless, it is interesting that a similar sequence (GATGA) is present 40 nucleotides upstream of the c-mos transcription initiation site (Fig. 5), although deletion of this element had no discernible effect on the activity of the c-mos promoter in microinjectedoocytes (Pal et al., 1991).

VI. NEGATIVE REGULATION OF C - ~ O S TRANSCRIPTION IN SOMATIC CELLS As discussed above, suppression of aberrant c-mos expression in somatic cells may be a particularly important aspect of c-mos regulation, especially in view of the ability of unregulated Mos synthesis to induce either cell death or transformation.We therefore analyzed a series of upstream c-mos deletions to attempt to identify negative regulatory elements involved in repression of c-mos transcription in somatic cells (Zinkel et al., 1992). These experiments identified a regulatory region extending from approximately 100to 200 nucleotides upstream of the spermatocyte c-mos promoter, deletion of which resulted in transcription from the c-mos spermatocyte promoter in NIH 3T3 and other somatic cell types (Fig. 6). The c-mos negative regulatory element (NRE) sequences are highly conserved in the mouse, rat, and human genes, consistent with the fact that the mouse NRE was also active in suppressingc-mos transcription in rat and human cell lines. In addition to its activity in association with the c-mos promoter, the NRE also suppressed transcription when inserted upstream of a heteroiogous promoter, indicating that it acted as a transcriptional silencer in somatic cells. In contrast, this sequence had no effect on transcription of c-mas in microinjected oocytes, so it appears

CCAAGTT CACTGT

CTAGCACTAA

,

Prot

T

I

CACTGT

CTAGCACTA

t

T

-764

Pgk2 E3/E4 region

CTCTTC I\

-501

Mouse Human

CCAAATC TCAGATC

figure 6. The c-mos negative regulatory element (NRE). Nucleotide positions of the NRE are shown relative to the spermatocyte transcription start site, taken as 280 base pairs upstream of the c-mos ATG (see Fig. 4). The endpoints of the NRE are defined by deletions that allow c-m5s expression in NIH 3T3 and other somatic cells. Mutations of the sequences designated by boxes 1,2, and 3 also allow c-mos transcription in NIH 3T3 cells, indicating that these sequences representfunctional elements within the NRE. Boxes 1 and 2 are similar to sequences upstream of the protamine (Prot) promoter that inhibit in vitro transcription in HeLa cell extracts. A sequence just upstream of box 2 is also similar to a putative repressor-binding site in the regulatory region of Pgk2.

that these sequences specifically inhibit c-mos transcription in somatic cells. It should be noted that the NRE defined in these experiments is distinct from the previously described c-mos UMS sequence (Blair et al., 1984; Wood et al., 1984 ). The UMS is located approximately 1.4 kb upstream of the c-mos spermatocyte promoter and was identified because it blocked activation of c-mos transforming potential by insertion of retroviral promoters. It is thought to act as a transcriptional terminator, blocking transcription of c-mos initiated at upstream sequences. However, both the spermatocyte and oocyte transcription initiation sites are substantially downstream of the UMS. Moreover, the presence or absence of the UMS does not affect c-mos expression in either microinjected oocytes (Pal et al., 1991) or transfected NIH 3T3 cells (Zinkel et a]., 1992). It thus appears unlikely that the UMS functions as a negative regulator of c-mos transcription from either the spermatocyte or oocyte promoters in somatic cells.

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Sequence comparisons indicated that the c-mos NRE contained three regions of similarity with 5’ flanking sequences of the mouse protamine gene that had been found to inhibit in vitro transcription in HeLa cell, but not male germ cell, extracts (Bunick et al., 1990). Point mutations or deletions of these c-mos sequences (defined as c-mos box 1 and box 2; Fig. 6) and of sequences immediately downstream of these regions (box 3) were sufficient to relieve suppression of c-mos promoter activity in transfected NIH 3T3 cells, indicating that they represent functional components of the NRE (Zinkel et al., 1992). In addition, a putative repressor binding site in the regulatory region of the spermatocyte-specific Pgk-2 gene (Gebara and McCarrey, 1992) is also similar to c-mos sequence just upstream of and possibly overlapping box 2. It is thus an intriguing possibility that these conserved sequences contribute to a general mechanism by which transcription of germ cell-specific genes is repressed in somatic cells. The c-mos NRE defined by the deletions and point mutations described above spans at least 60-100 base pairs, suggesting that it consists of multiple interacting elements. We have begun to investigate protein binding sites within this region by gel-shift analysis and have identified at least three complexes that form with nuclear proteins from N M 3T3 cells (Wenhao Xu and G. M. Cooper, manuscript submitted).One of these complexes involves specific interaction with the NRE box 2 sequences, which have been established as a functional component of the NRE because mutations in box 2 activate the c-mos promoter in NIH 3T3 cells (Zinkel et al., 1992). Proteins extracted from testicular germ cells also form complexes with the NRE but do not appear to interact with the box 2 element. The box 2-binding protein identified specifically in N M 3T3 extracts therefore appears to be a good candidate for a somatic cell repressor of c-mos transcription. The multiple proteins binding to the NRE may well cooperate to achieve tissue-specific c-mos expression. It is important to note that although we identified the NRE by its ability to suppress transcription in somatic cells, tissue-specific transcription may also involve the binding of spermatocyte transcriptional activators to this region. A well-studied example of the complex cooperative interactions responsible for tissuespecific transcription of other genes is provided by the immunoglobulin heavy chain enhancer, which contains binding sites for at least eight proteins, some of which repress transcription in nonlymphoid cells and others of which specifically activate transcription in B lymphocytes (Libermann et al., 1990; Ruezinsky et al., 1991). Further analysis of the

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c-mos NRE may well reveal similar interactions between activators and repressors of c-mos transcription, contributing to the highly specific transcription of c-mos in male germ cells.

VII. SUMMARY AND FUTURE DIRECTIONS The novelty of the c-mos proto-oncogene resides in its specific expression in germ cells, where it appears to play a unique role in regulating the meiotic cell cycle. In oocytes, the general role of Mos appears to be established: it acts to stabilize cyclin B, thereby maintaining MPF activity as required for progression from meiosis I to meiosis I1 and subsequent metaphase I1 arrest. However, the biochemical basis for Mos action is not yet known. Although Mos can phosphorylate cyclin B in v i m , it is not clear whether this is responsible for inhibiting cyclin B degradation. It is alternatively possible that Mos stabilizes cyclin B by inhibiting (either directly or indirectly) the enzymes responsible for cyclin B proteolysis. In addition, the role of Mos in initiation of meiosis I in Xenopus, and possibly in mouse, remains to be determined. Also in need of further study is the function of Mos in male germ cells. It appears that Mos is expressed prior to meiosis of spermatocytes, consistent with the possibility that it acts in the initiation of meiosis and/or during progression from meiosis I to meiosis 11. However, c-mos expression continues in postmeiotic spermatids,where it does not induce metaphase I1 arrest. This may be due to the absence of other components of cytostatic factor, but the role of c-mos expression in postmeiotic male germ cells remains unclear. The importance of tissue-specific regulation of c-mus transcription is indicated by the consequences of aberrant Mos expression in somatic cells. High levels of Mos are cytotoxic, apparently as aresult of arresting cell division at metaphase. However, low levels of expression of structurally normal Mos protein are sufficient to induce cell transformation, possibly as a result of stimulating MAP kinase. The potential relationships between the activities of Mos as a cytostatic factor and as a transforming oncogene are not understood. It is possible that phosphorylation of a common target (e.g., MAP kinase) is involved in both transformation and metaphase arrest, with variations in Mos expression determining the outcome. Alternatively, transformation and arrest might result from phosphorylation of distinct targets, also dependent on levels of Mos activity.

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The ability of unregulated c-mos expression to induce cell transformation contrasts with other protein kinase proto-oncogenes, which require structural mutations in their gene products for oncogenic activity. It thus appears that control of c-mos transcription is a critical aspect of its regulation. Two sets of cis-acting transcriptional regulatory sequences have now been identified: an Inr-like element that activates c-mos transcription in oocytes, and a negative regulatory element that represses c-mos transcription in somatic cells. Identification of the transcription factors that interact with these sequences may contribute to understanding the regulation not only of c-mos, but also of other genes that are specifically expressed in germ cells.

ACKNOWLEDGMENTS I am pleased to acknowledge the collaboration of Ann Kiessling in many of the experiments from our laboratory, as well as the contributions of Debra Goldman, Katsuhisa Kogawa, Stephen O’Keefe, Subrata Pal, Donald Torry, Heiner Wolfes, Wenhao Xu, and Sandra Zinkel to these studies. Work in our laboratory has been supported by National Institutes of Health grant HD26594.

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Dent, P., Haser, W., Haystead, T. A. J., Vincent, L. A., Roberts, T. M., and Sturgill, T. W. (1992). Activation of mitogen-activated protein kinase kinase by v-Raf in NIH 3T3 cells and in vitro. Science 257: 1404-1407. de Vries-Smits, A. M. M., Burgering, B. M. Th., Leevers, S. J., Marshall, C. J., and Bos, J. L. (1992). Involvement of p21msin activation of extracellular signal-regulated kinase 2. Nature 357: 602-604. Draetta, G., Luca, F., Westendorf, J., Brizuela, L., Ruderman, J., and Beach, D. (1989). cdc2 protein kinase is complexed with both cyclin A and B: evidence for proteolytic inactivation of MPF. Cell 56 829-836. Gabrielli, B. G., Roy, L. M., and Maller, J. L. (1993). Requirement for Cdk2 in cytostatic factor-mediated metaphase I1 arrest. Science 259: 1766-1 769. Gebara, M. M., and McCarrey, J. R. (1992). Protein-DNA interactions associated with the onset of testis-specific expression of the mammalian Pgk-2 gene. Mol. Cell. Biol. 12: 1422-1431. Gerhart, J., Wu, M., and Kirschner, M. (1984). Cell cycle dynamics of an M-phasespecific cytoplasmic factor in Xenopus laevis oocytes and eggs. J. Cell Biol. 98: 1247-1255. Goldman, D. S., Kiessling, A. A., Millette, C. F., and Cooper, G. M. (1987). Expression of c-mos RNA in germ cells of male and female mice. Roc. Natl. Acad. Sci. USA 8 4 45094513. Goldman, D. S., Kiessling, A. A,, and Cooper, G. M. (1988). Post-transcriptional processing suggests that c-mos functions as a maternal message in mouse eggs. Oncogene 3: 159-1 62. Hanks, S. K., Quinn, A. M., and Hunter, T. (1988). The protein kinase family: conserved features and deduced phylogeny of the catalytic domains. Science 241: 42-52. Hashimoto, N., and Kishimoto, T. (1988). Regulation of meiotic metaphase by a cytoplasmic maturation-promoting factor during mouse oocyte maturation. Dev. Biol. 126 242-252. Heidecker, G., Huleihel, M., Cleveland, J. L., Kolch, W., Beck, T.W., Lloyd, P., Pawson, T., and Rapp, U. R. (1990). Mutational activation of c-rafl and definition of the minimal transforming sequence. Mol. Cell. Biol. 10: 2503-2512. Herzog, N. K., Ramagli, L., and Arlinghaus, R. B. (1989). Somatic cell expression of the c-mos protein. Oncogene 4: 1307-1315. Howe, L. R., Leevers, S. J., Gomez, N., Nakielny, S., Cohen, I?, and Marshall, C. J. (1992). Activation of the MAP kinase pathway by the protein kinase raf. Cell 71: 335-342. Kanki, J. P., and Donoghue, D. J.(1991). Progression from meiosis I to meiosis I1 in Xenopus oocytes requires de novo translation of the mos protooncogene. Proc. Natl. Acad. Sci. USA 88: 5794-5798. Keshet, E., Rosenberg, M. P., Mercer, J. A., Propst, F., Vande Woude, G. F., Jenkins, N. A,, and Copeland, N. G. (1988). Developmental regulation of ovarian-specific Mos expression. Oncogene 2: 235-240. Kirschner, M. (1992). The cell cycle then and now. Trends Biochem. Sci. 17: 28 1-285. Kyriakis, J. M., App, H., Zhang, X., Banerjee, P., Brautigan, D. L., Rapp, U. R., and Avruch, J . (1992). Raf-1 activates MAP kinase-kinase. Nature 358:417421.

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Leibovitch,S. A., Lenormand, J. L.,Leibovitch,M. P.,Guiller, M., Mallard, L.,andHarel, J. (1990). Rat myogenic c-mos cDNA: cloning sequence analysis and regulation during muscle development. Oncogene 5: 1149-11 57. Libermann, T. A., Lenardo, M., and Baltimore, D. (1990). Involvement of a second lymphoid-specific enhancer element in the regulation of immunoglobulin heavychain gene expression. Mol. Cell. Biol. I 0 3155-3162. Lira, S. A., Kinloch,R. A.,Mortillo, S . , and Wassman, €? M.(1990). An upstreamregion of the mouse ZP3 gene directs expression of firefly luciferase specifically to growing oocytes in transgenic mice. Proc. Natl. Acad. Sci. USA87 7215-7219. Martinez-Salas,E., Linney, E., Hassell, J., and DePamphilis, M. L. (1989). The need for enhancers in gene expression first appears during mouse development with formation of the zygotic nucleus. Genes Dev. 3: 1493-1506. McGrew, L. L., Dworkin-Rastl, E., Dworkin, M. B., and Richter, J. D. (1989). Poly(A) elongation during Xenopus oocyte maturation is required for translational recruitment and is mediated by a short sequence element. Genes Dev. 3: 803-815. Millar, S . E., Lader, E., Liang, L-E, and Dean, J. (1991). Oocyte-specificfactors bind a conserved upstream sequence required for mouse zona pellucida promoter activity. Mol. Cell. Biol. 11: 6197-6204. Murray, A. W. (1992). Creative blocks: cell-cycle checkpoints and feedback controls. Nature 359: 599-604. Murray, A. W., Solomon, M. J., and Kirschner, M. W. (1989).The role of cyclin synthesis and degradation in the control of maturation promoting factor activity. Nature 339: 280-286. Murre, C., and Baltimore, D. (1992). The helix-loop-helixmotif: structure and function. In: TranscriptionalRegulation (McKnight, S. L., and Yamamoto, K. R., eds.), pp. 861-879. Cold Spring Harbor Laboratory, Plainview, NY. Mutter, G. L., and Wolgemuth, D. J. (1987). Distinct developmental patterns of c-mos proto-oncogene expression in female and male mouse germ cells. Proc. Natl. Acad. Sci. USA 88: 7869-7872. Mutter, G. L., Grills, G. S . , and Wolgemuth, D. J. (1988). Evidence for the involvement of the proto-oncogene c-mos in mammalian meiotic maturation and possibly very early embryogenesis. EMBO J. 7: 683-689. Nakatani, Y., Horikoshi, M., Brenner, M., Yamamoto,T., Besnard,F., Roeder, R. G., and Freese, E. (1990). A downstream initiation element required for efficient TATA box binding and in vitro function of TFIID. Nature 348: 86-88. Nebreda, A. R., and Hunt, T. (1993). The c-mos proto-oncogene protein kinase turns on and maintains the activity of MAP kinase, but not MPF, in cell-free extracts of Xenopus oocytes and eggs. EMBO J. 12: 1979-1986. Okazaki, K., Nishjzawa,M., Furuno, N., Yasuda, H., and Sagata, N. (1992). Differential occurrence of CSF-like activity and transforming activity of Mos during the cell cycle in fibroblasts. EMBO J. 11: 2447-2456. O'Keefe, S. J., Wolfes, H., Kiessling, A. A., and Cooper, G. M. (1989). Microinjection of antisense c-mos oligonucleotidesprevents meiosis I1 in the maturing mouse egg. Proc. Natl. Acad. Sci. USA 86 7038-7042. OKeefe, S. J., Kiessling, A. A., and Cooper, G. M. (1991). The c-mos gene product is required for cyclin B accumulation during meiosis of mouse eggs. Proc. Natl. Acad. Sci. USA 88: 7869-7872.

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Pal, S. K., Torry, D., Serta, R., Crowell, R. C., Siebel, M. M., Cooper, G. M., and Kiessling, A. A. (1994). Expression and potential function of the c-mos protooncogene in human eggs. Fertil. Steril. 61:496-503. Pal, S. K., Zinkel, S. S., Kiessling, A. A., and Cooper, G. M. (1991). c-mos expression in mouse oocytes is controlled by initiator-related sequences immediately downstream of the transcription initiation site. Mol. Cell. Biol. 11: 5190-5196. Papkoff, J., Verma, I. M., and Hunter, T. (1982). Detection of a transforming gene product in cells transformed by Moloney murine sarcoma virus. Cell 29: 417-426. Paules, R. S., Propst, F., Dunn, K. J., Blair, D. G., Kaul, K., Palmer, A. E., and Vande Woude, G. F. (1988). Primate c-mos proto-oncogene structure and expression: transcription initiation both upstream and within the gene in a tissue-specific manner. Oncogene 3: 59-68. Paules, R. S., Buccione, R., Moschel, R. C., Vande Woude, G. F., and Eppig, J. J. (1989). Mouse mos proto-oncogene product is present and functions during oogenesis. Proc. Natl. Acad. Sci. USA 86 5395-5399. Paules, R. S., Resnick, J., Kasenally, A. B., Emst, M. K., Donovan, P., and Vande Woude, G. F. (1992). Characterization of activated and normal mouse mos gene in murine 3113 cells. Oncogene 7 2489-2498. Posada, J., Yew, N., Ahn, N. G., Vande Woude, G. F., and Cooper, J. A. (1993). Mos stimulates MAP kinase in Xenopus oocytes and activates a MAP kinase kinase in vitro. Mol. Cell. Biol. 13: 2546-2553. Propst, F., and Vande Woude, G. F. (1985). Expression of c-mos proto-oncogene transcripts in mouse tissues. Nature 315: 516-518. Propst, F., Rosenberg, M. F?, Iyer, A., Kaul, K., and Vande Woude, G. F. (1987). c-mos proto-oncogene transcripts in mouse tissues: structural features, developmental regulation, and localization in specific cell types. Mol. Cell. Biol. 7 1629-1637. Roy, L. M., Singh, B., Gautier, J., Arlinghaus, R. B., Nordeen, S. K., and Maller, J. L. (1990). The cyclin B2 component of MPF is a substrate for the c-moSxe proto-oncogene product. Cell 61: 825-831. Roy, A. L., Meisterernst, M., Pognonec, P., and Roeder, R. G. (1991). Cooperative interaction of an initiator-binding transcription initiation factor and helix-loop-helix activator USE Nature 354: 245-248. Ruezinsky, D., Beckmann, H., and Kadesch, T. (1991). Modulation of the IgH enhancer’s cell type specificity through a genetic switch. Genes Dev. 5: 29-37. Sagata, N., Oskarsson, M., Copeland, T., Brumbaugh, J., and Vande Woude, G. F. (1988). Function of c-mos proto-oncogene in meiotic maturation in Xenopus oocytes. Nature 335 519-525. Sagata, N., Watanabe, N., Vande Woude, G. F., and Ikawa, Y. (1989). The c-mos proto-oncogene product is a cytostatic factor responsible for meiotic arrest in vertebrate eggs. Nature 342: 512-51 8. Schickler, M., Lira, S. A., Kinloch, R. A., and Wassarman, P. M. (1992). A mouse oocyte-specific protein that binds to a region of mZP3 promoter responsible for oocyte-specific mZP3 gene expression. Mol. Cell. Biol. 12: 120-127. Schmidt, M., Oskarsson, M. K., Dunn, J. K., Blair, D. G., Hughes, S., Propst, F., and Vande Woude, G. F. (1988). Chicken homolog of the mos proto-oncogene. Mol. Cell. Biol. 8: 923-929.

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Siracusa, G., Whittingham, D. G., Molinaro, M., and Vivarelli, E. (1978). Parthenogenetic activation of mouse oocytes induced by inhibitors of protein synthesis. J. Embryol. Exp. Morphol. 43: 157-1 66. Smale, S. T., and Baltimore, D. (1989). The “initiator” as a transcriptional control element. Cell 57: 103-113. Stanton, V. I?, Jr., Nichols, D. W., Laudano, A. P., and Cooper, G. M. (1989). Definition of the human raf amino-terminal regulatory region by deletion mutagenesis. Mol. Cell. Biol. 9 639-647. Thomas, S. M., DeMarco, M., D’Arcangelo, G., Halegoua, S . , and Brugge, J. S. (1992). Ras is essential for nerve growth factor- and phorbol ester-induced tyrosine phosphorylation of MAP kinases. Cell 68: 1031-1040. Van der Hoorn, F. A., Spiegel, J. E., Maylie-Pfenninger, M. F., andNordeen, S . K. (1991). A43 kd c-mos protein is only expressed before meiosis during rat spermatogenesis. Oncogene 6 929-932. Vassalli, J. D., Huarte, J., Belin, D., Gubler, I?, Vassalli, A,, O’Connell, M. L., Parton, L. A., Rickles, R. J., and Strickland, S . (1989). Regulated polyadenylation controls mRNA translation during meiotic maturation of mouse oocytes. Genes Dev. 3: 2 163-21 71. Wasserman, W. J., and Masui, Y. (1975). Effect of cycloheximide on acytoplasmic factor initiating meiotic maturation in Xenopus oocytes. Exp. Cell Res. 91: 381-388. Wood, T. G., McGeady, M. L., Blair, D. G., and Vande Woude, G. F. (1983). Long terminal repeat enhancement of v-mos transforming activity: identification of essential regions. J. Virol. 46 726-736. Wood, T. G., McGeady, M. L., Baroudy, B. M., Blair, D. G., and Vande Woude, G. F. (1984). Mouse c-mos oncogene activation is prevented by upstream sequences. Proc. Natl. Acad. Sci. USA81: 7817-7821. Wood, K. W., Sarnecki, C., Roberts, T. M., and Blenis, J. (1992). ras mediates nerve growth factor receptor modulation of three signal-transducing protein kinases: MAP kinase, Raf-1, and RSK. Cell 68: 1041-1050. Yew, N., Mellini, M. L., and Vande Woude, G. F. (1992). Meiotic initiation by the mos protein in Xenopus. Nature 355:649-652. Zhao, X., Batten, B., Singh, B., and Arlinghaus, R. B. (1990). Requirement of the c-mos protein kinase for murine meiotic maturation. Oncogene 5: 1727-1730. Zhao, X., Singh, B., and Batten, B. E. (1991). The role of c-mos proto-oncoprotein in mammalian meiotic maturation. Oncogene 6 43-49. Zinkel, S . S . , Pal, S . K., Szeberenyi, J., and Cooper, G. M. (1992). Identification of a negative regulatory element that inhibits c-mos transcription in somatic cells. Mol. Cell. Biol. 12: 2029-2036.

REGULATION OF PIGMENTATION DURING MAMMALIAN DEVELOPMENT

Friedrich Beermann, Ruth Gang, and Gunther Schutz

.

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 11. Melanocytes Originate from the Neural Crest . . . . . . . . . . . . . . . . 151 111. Role of the Dominant spotting Locus and the Steel Locus in the Migration and Proliferation of Melanocytes . . . . . . . . . . 154 IV. Genes Affecting MelanocyteDermis Interaction and the Morphology of Melanocytes . . . . . . . . . . . . . . . . . . . . . 156 V. Melanin Synthesis-The Role of Tyrosinase and Related Proteins . . . . . 158 VI. The Mouse Tyrosinase Gene . . . . . . . . . . . . . . . . . . . . . . . . . 161 VII. Expression Analysis of the 'Qrosinase Gene in Transgenic Mice . . . . . . 163 A. Rescue of the Albino Phenotype . . . . . . . . . . . . . . . . . . . . . 163 B. Identification of Sequences Required for Expression in Pigment Cells . 167 C. Expression of the Tyrosinase Gene During Development . . . . . . . . 167

Advances in DevelopmentalBiochemistry Volume 3, pages 149-177. Copyright 0 1994 by J A I Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-865-X 149

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VIII. In Search of Pigment Cell-Specific Factors: Which Elements Determine Expression in Melanocytes? . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. 169 171 171

1. INTRODUCTION It was at the beginning of the century that the inheritance of coat color became the first mammalian trait to be analyzed by genetic studies. One of the first mutations analyzed that demonstrated Mendelian inheritance in mice was the albino phenotype (Castle and Allen, 1903). Albino and a second mutation, pink-eyed, represented the f i s t example of genetic linkage in the mouse (Haldane et al., 1915; in Hogan et al., 1986). Subsequent investigations elucidated the cellular basis of pigmentation, providing evidence that melanogenesis occurs in melanocytes and that tyrosinase is the key enzyme required for production of melanin (for review see Hearing and JimCnez, 1989; Hearing and Tsukamoto, 1991). A large number of genes and mutations affecting coat color have been identified in mice (Silvers, 1979; Lyon and Searle, 1989). These studies have been aided by the fact that dominant and recessive mutations are rarely lethal and produce an easy, identifiable phenotype. Some of these mutations have their origins in the mouse fancy stock, kept for their aesthetics by mouse breeders and collectors of pet mice. Other mutations have arisen spontaneously in mouse colonies or were induced following mutagenesis by radiation or chemicals. Overall, these mutations have Table 7. Pigmentation Genes Act at Different Levels Level

LocudGene

I. Migration and proliferationof melanocyteslmelanoblasts Wlc-kit SUkit ligand 11. Interaction between hair follicle and melanocytes ufligand? 111. Shape and ultrastructure of melanocytes dh4HC plmembrane protein IV. Enzymes involved in melanin synthesis cltyrosinase slurylDOPAchrome tautomerase Notes: Well-characterizedexamples for mutant loci and their respective gene products (when known) in the mouse are given (see text for references). W = Dominant sporting; Sl = Steel; a = nonagouti; d = dilute; c = albino; p = pink-eyed dilution: MHC = myosin heavy chain.

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been mapped to more than 50 distinct loci throughout the mouse genome that act at different levels in the melanogenic process (Silvers, 1979; Jackson, 1991). These loci encode structural genes as well as regulatory genes that affect the expression and/or establishment of the phenotype (see Table 1 for a set of well-characterized mutations). In this review, we will confine our studies on mouse pigmentation solely to the production of melanin in melanocytes. Other pigments like neuromelanin or lipofuscin are not the subject of this review. We will mainly focus on neural crest-derived melanocytes, and less on the pigment-producing cells of the retinal pigment epithelium. Despite their different origins (neural crest versus neuroectoderm) and functions (secretion of pigment granuledmature melanosomes versus retention), the two types of pigment cells share the same biosynthetic pathway. With respect to skin pigmentation several possible roles have been attributed to melanin (Hill, 1992). These include its function as a sunscreen, thermoregulator, camouflage, or radical scavenger. For example, melanins can absorb different forms of energy and dissipate them as heat. If the energy input is too great, the capacity of the pigment to detoxify the radicals is exceeded and potentially damaging radical specieshydroxyl radicals-are produced.

11. MELANOCYTES ORIGINATE F R O M THE NEURAL CREST The neural crest is a vertebrate structure and consists of a transient population of ectodermally derived cells that are in the angle between the neural tube and the body ectoderm and appear upon closure of the neural tube (see Le Douarin, 1982). Neural crest cells disperse from their location at the dorsal surface of the neural tube and migrate extensively through the embryo. They give rise to a great variety of different cell types including most neurons of the peripheral nervous system, adrenomedullary cells, and skeletal and connective tissue of the face and head. With the exception of the retinal pigmented epithelium, neural crest cells are the sole source of all pigment cells of the body. The majority of analyses on neural crest cell migration, proliferation, and differentiation have been carried out in avian and amphibian embryos because of the relative ease of experimental manipulation (see Le Douarin, 1982). These experimentsdemonstrated that pigment cells arise from the neural crest. When pieces of neural foIds were grafted to the

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ventral side of an embryo or explanted in vitro, this resulted in the appearance of melanophores. The most compelling evidence was provided by Rawles when he transplanted neural crest from black mouse embryos into the coelom of unpigmented chick (Rawles, 1947). Pigmentation was obtained in the otherwise unpigmented chick, thus demonstrating that mouse neural crest cells can give rise to melanoblasts. Neural crest cells follow two major migratory pathways once they detach from the neural tube. Cells migrating ventrally through the rostra1 portion of each somitic sclerotome give rise to structures such as dorsal root ganglia, sympathetic ganglia, the adrenal medulla, and Schwann cells. The second migratory pathway used by melanoblasts in the mouse is rather dorsolateral,between the dermamyotomeand the epidermis,just below the basement membrane (Serbedzija et al., 1990; 1992). Once melanoblasts have reached their presumptive location they may penetrate the basement membrane and invade the epidermal ectoderm, where they further proliferate (see Hirobe, 1992) and become incorporated into developing hair follicles. A major focus of pigment research in the mouse was to determine the number of precursor cells that finally give rise to melanoblasts and melanocytes. Pioneering work in this area was performed by Mintz using the then rather novel approach of generating chimeras (Mintz, 1967; 1970). These chimeras were generated by fusing (or aggregating) fouror eight-cell embryos of different mouse strains carrying dissimilar coat color genes. The resulting mice therefore have four parents (tetraparental) and represent a mixture of cells of different genotype. In these experiments Mintz obtained a high incidence of patterns with transverse bands of alternate colors on the head and body. At the borders some intermixing between the two colors was observed. The dorsal midline represented a sharp boundary, and the stripe patterns on either side of the mice seemed to be established independently. From these experiments, Mintz concluded that only 17 pairs of cells originating from the neural crest constitute the precursors or clonal initiators of melanoblasts.Three pairs contribute to the head, six pairs to the body, and eight to the tail, giving rise to the maximum of 34 bands found (Mintz, 1967, 1970). At boundaries descendantsof adjacent clones are mixed, which is seen when these clones are of different genotype (i.e., show different pigmentation). Therefore, in the mouse, a limited subpopulationof neural crest cells are committed to becoming pigment cells, and these melanocyte precursors yield clones of identical cells. Which factors or genes determine these events during differentiation in the mouse remains to be elucidated.

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A quite different approach was taken by Jaenisch (1985). Mammalian neural crest cells were isolated from explants of 8.5-day-old C57BL/6J embryos and injected back into 9-day-old mouse embryos derived from an unpigmented albino strain (BALBk). These cells can then migrate over considerable distances and participate in normal development. The resulting chimeric mice show pigmentation in hair and iris. The fraction of chimeric mice obtained was strongly dependent on the age of the neural crest cell cultures, and on the age of donor embryos. The most effective pigment contribution was observed when neural crest cells were derived from 8.75- to 9.25-day-old embryos and cultured for 3-5 days (Jaenisch, 1985; this was recently rectified to 2 days, Huszar et al., 199la). Pigmented cells were also detected in the eye but, consistent with earlier findings, were restricted to the neural crest-derived pigmented cell layers (i.e., the choroid and the iris). Interestingly, coat pigmentation was restricted to the head region and posterior trunk but never found in anterior trunk and forelimbs (Jaenisch, 1985). More recent experiments have demonstrated the dependence on the genotype of the host embryo (Huszar et al., 1991a). These investigators used embryos carrying a mutation at the W locus (see below) characterized by the absence of functional endogenous melanoblasts. Consequently, migration of introduced melanoblast precursors was not limited to the head, for example, but instead melanoblasts spread through an quasiempty space that was free of host. melanoblasts. W mutant neural crest chimeras displayed extensive pigmentation, often exceeding 50% of the coat. The pattern of pigmentation was, at least in 90%of the animals, consistent with an entry of cells at the dorsal midline, followed by subsequent proliferation and migration dorsoventrally and laterally. The same technique was used to analyze the clonal history of melanocytes (Huszar et al., 1991b), which should be consistent with the earlier findings of Mintz (1967; 1970; see above). Neural crest cells were infected with a tyrosinase-expressing retrovirus, thus marking the otherwise unpigmented BALBk- or FVB/N-derived cells (Huszar et al., 199lb). Mosaic animals obtained following the in utero microinjection procedure showed pigmented bands. The width of these bands was consistent with that seen in aggregation chimeras, thus verifying the contention that the stripes seen represent clonal descendants of a single progenitor melanoblast. The clonal origin of pigment is also obvious in mice with a heritable striped pigment pattern. In addition to mutations like chinchilla-mottled(Silvers, 1979), these mice occasionally occur as a by-product of transgenic experiments where the transgene is expressed

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in melanocytes. Mice with such a phenotype have been described expressing the mouse tyrosinase gene (Mintz and Bradl, 1991) or the SV40 tumor antigene (Bradl et al., 1991a,b). Expression of the SV40 transforming sequences led to diminished differentiation of melanized pigment granules concomitant with a lighter hypopigmented coat. One transgenic mouse showed a completely striped coat similar to that of allophenic mice. This pattern of light transverse stripes on a dark (C57BL/6J) background is autosomally inherited by all transgenic offspring (Bradl et al., 1991b). These transgenic mice therefore represent phenotypic mosaics, since the transgene present in all cells is expressed differently in different pigment cell clones. A striped pattern supports the conclusion that the original difference is determined in the clonal initiators. The color pattern in this line therefore exemplifies the concept of phenoclones, which are defined as phenotypically different clones among cells of the same type that correspond to the developmental lineages of which that type comprises.

111. ROLE OF THE DOMINANTSPOTTINC LOCUS AND THE S T E L LOCUS IN THE MIGRATION AND PROLIFERATION OF MELANOCYTES Mice with mutations at either the Dominant spotting locus (W) or the Steel ( S o locus produce a very similar range of phenotypic effects, causing impairments in germ cells, hematopoietic cells, and neural crest-derived melanocytes (Silvers, 1979). The genes encoded by these loci have recently been identified. The W locus is a tyrosine kinase receptor encoded by the c-kit oncogene. Its ligand, a growth factor, has been identified as the product of the Steel locus also known as kit ligand (KL), mast cell growth factor (MGF), or stem cell factor (SCF) (Geissler et al., 1988; Witte, 1990; Zsebo et al., 1990; Nocka et al., 1990; Wagner and Alexander, 1991).This receptorfligand interaction explains why the W gene product (as a receptor) acts cell-autonomously, in contrast to the Sl product (as a growth factor). Melanoblasts migrate from the neural crest to skin, hair follicle, and choroid. Several mutations including Sl and W affect this migration and can generate white patches or spots in the coat because of a disturbed migration and/or proliferation andor differentiation of melanoblasts. These effects are even obvious in heterozygous animals, and W/+ mice show a characteristic white belly spot with some unpigmented hairs. The

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range of mutations varies from severe (W2)with no pigment in heterozygous and lethality when homozygous, to a milder phenotype (W7)having a white belly spot in heterozygotes and larger white patches in homozygotes (see Jackson, 1991). These phenotypical analyses have unequivocally proven the involvement of c-kit in melanocyte developmentin embryonic and postnatal life. However, it was unclear at which stage of melanocyte development c-kit is required. In situ hybridization analyses first showed direct evidence for c-kit expression during early development (Manova and Bachvarova, 1991). According to these data, c-kit is expressed along the presumptive path of melanoblast migration at day 10-1 1 of development. Recently, it was questioned whether these cells are really melanoblasts or rather other c-kit-positive cells like mast cell precursors (Steel et al., 1992). Nishikawa et al. (1991) used a monoclonal anti-c-kit antibody to interfere with proper development of melanocytes during embryonic and postnatal life. When injected intradermally into pregnant mice migration of melanocytes was inhibited in the offspring. Unexpectedly, antibody injections in newborn or adult mice also led to alterations of coat colors. The authors of this study therefore conclude that two major c-kitdependent processes occur in melanocyte development.During mid-gestation, at around day 14.5, c-kit is necessary for proliferation of melanoblasts in the dermal layer and their entry into the epidermis. In adult life, c-kit is required for activation of melanocytes in the fully developed hair follicles. This latter finding is quite attractive; however, it remains to be elucidated how activation of the hair growth cycle, c-kit expression, and melanocyte proliferation are related. Evidence for a c-kit function in early melanogenesis was also provided by a recent transgenic experiment (Ray et al., 1991). A number of data had indicated that the most severe W mutant, W2,encodes a c-kit protein product that contains a missense mutation and lacks tyrosine kinase activity (Tan et al., 1990).This protein nonethelessgets incorporated into receptor heterodimers and possibly interferes with c-kit ligand-induced signal transduction. To investigate the dominant-negative character of the V 2c-kit gene product, Ray et al. (1991) generated transgenic mice in which the c-kit v2 cDNA was expressed under control of the constitutive human p-actin promoter. Ectopic expression of the W 2c-kit gene product recapitulates some W phenotypes and shows that it can act in a dominant fashion. The coat color phenotypes observed are not characterized by stripes but rather by a more splotchy appearance. In contrast to allophenic mice (Mintz 1967, 1971), unpigmented areas in W mutant

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mice are devoid of melanocytes. Thus, consistent with the experiments described above (Huszar et al., 199la), viable pigmented melanoblasts/melanocytesmay have entered the adjacent nonpigmented regions at some sites. In conclusion, the data provide further evidence for a function of c-kit during early development in melanoblast precursors. c-kit, which was originally described as the cellular homologue of the transforming viral gene (v-kit) of a feline retrovirus, is conserved in other mammalian species. Amutation in c-kit affecting,among other cell types, melanocytes has recently been described in rat (Tsujimura et al., 1991). In human, the autosomal dominant genetic disorder piebaldism-characterized by patches of skin and hair devoid of melanocytes-is most probably caused by mutations of c-kit. Deletions and point mutations affecting the tyrosine kinase domain have recently been identified in piebald patients (Fleischman et al., 1991;Giebel and Spritz, 1991;Spritz et al., 1992).

IV. GENES AFFECTING MELANOCYTE/DERMIS INTERACTION A N D THE MORPHOLOGY OF MELANOCYTES Once melanocytes have reached their proper location (e.g., the hair follicle) they begin to synthesize melanin. In addition to the genes involved in melanin synthesis, other genes are also required for a correct pigmented phenotype. The importance of such genes was highlighted by the occurrence of mouse mutations, which affect the type of pigment made and the morphology of the melanocyte. Wild-type mice have two types of pigment: eumelanin, which is black and colors the tip and base of dorsal hair, and phaeomelanin, a yellow or brown pigment coloring the middle of the hair (Silvers, 1979). Mutations at the a-locus (nonagouri) and e-locus (extension) indicate that these two loci are involved in determining the relative amount of eumelanin and phaeornelanin made. The site of action of the genes encoded by these loci is different: a acts outside of the melanocyte (Siracusa, 1991; Bultman et al., 1992), whereas the e locus product functions within the melanocyte. Both products have, similar to W and SI, properties of signaVligand and receptor, where a acts nonautonomously and e autonomously. The e locus has recently been shown to code for the receptor of the a-melanocyte-stimulating hormone (Robbins et al., 1993).

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The interaction of the a-locus product with melanocytes is quite interesting because of its action by virtue of the surroundinghair follicle cells. This has been demonstrated in mixing experimentsand in chimeras where only the genotype of follicle cells and not melanocytes was responsible for the resulting agouti phenotype (Silvers, 1979;Mayer and Fishbane, 1972; Mintz, 1970). Thus, chimeras generated between BALB/c mice (genotyped as Abc, i.e., Agouti brown albino) and C57BL/6J (genotyped as aBC, i.e., nonagouti Black) not only give patches of nonagouti coat but also agouti coat caused by a combination of BALB/c follicle cells and C57BL/6J melanocytes. A much finer striped pattern in chimeras is also observed, substantiating the fact that the gene products of the a-locus act through hair follicles. According to Mintz (1970) the approximate number of hair follicle cell precursors is around 85 on both sides. The gene product of the a-locus has recently been identified (Bultman et al., 1992). The gene is expressed as a 0.8-kb mRNA in skin of wild-type mice (MA), whereas the transcript seems to be absent in nonagouti mice (a/a). Other genes that interfere with the melanocyte morphology and affect the deposition of pigment have been identified. Perhaps the best known are those affecting the d locus (dilute) and the p locus (pink-eyed dilution). Many dilute mutations have recently been identified (see Silvers, 1979; Rinchik et al., 1985). The recessive dilute coat color mutation, which is present in some inbred mouse strains (e.g., DBA/2), produces a lightening of the coat color, caused by abnormal melanocyte morphology. These melanocytes do not extend their dendrites normally, which disturbs proper secretion of melanin granules into the hair. The gene product encoded by this locus has recently been identified as a novel type of myosin heavy chain; its C terminus shares elements of both type I1 (a-helical coiled coil) and type I (no-coiled coil) myosin heavy chains (Mercer et al., 1991). The lack of melanocyte dendrites in dilute mutations suggests an important role of the dilute gene product in correct function of melanocytes. The dilute coat color of mice homozygous for dilute and two similar mutations (leaden and ashen) is partially restored by another locus, the so-called dilute suppressor (dsu) (Moore et a]., 1988, 1990). Phenotypically, the melanocytes again become dendritic, which facilitates the transportation of melanosomes into the hair shaft and produces a darkened coat color. It was suggested that dsu interacts intracellularlywith the dilute gene product and compensatesfor its defect (Jackson, 1991).

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Many mutations at the pink-eyed dilution locus (p) have been recovered in mutagenesis experiments (see Silvers, 1979; Lyon et al., 1992). Homozygotes have pink eyes with reduced but not absent pigmentation. The black eumelanin of the hair is diluted, whereas yellow phaeomelanin is only slightly affected. The melanosomes appear smaller and abnormal in shape. Several of the radiation-inducedmutations at the locus showed pleiotropic effects, affecting not only pigmentation, but also development, reproduction, and behavior. Extensive complementation analyses provided evidence that these mutations represent physical deletions, and at least four neighboring loci were postulated to be involved (Lyon et al., 1992). Molecular analyses finally resulted in characterization and identification of the mouse p gene (Brilliant et al., 1991; Brilliant, 1992; Gardner et al., 1992;Rinchik et al., 1993).The gene presumably encodes an integral membrane protein and thus may be a component of the melanosomal membrane. Expression of the 3.4-kb mRNA generated by the gene is strongly reduced in homozygous and heterozygous mutants for the p mutation (Gardner et al., 1992; Rinchik et al., 1993). Furthermore, analyses of the human homologue ( P ) have suggested that mutations in the gene may cause tyrosinase-positive(type 11)oculocutaneous albinism (Rinchik et al., 1993). Another melanocyte-specific gene, Pmell7, which cross-reacts with anti-tyrosinase antibodies (Kwon et al., 1987a), was tentatively assigned near the silver (so locus on mouse chromosome 10 (Kwon et al., 1991). Silver induces hypopigmentation involving a reduction in the number of melanocytes rather than a deficiency in melanin synthesis. To provide more information on a possible defect intrinsic to silver, immortal si/si melanocyte lines were established (Spanakis et al., 1992). Although otherwise appearing normal, si/si melanocytes show reduced growth and proliferation in culture. This supportsthe idea that the silver gene product functions as a growth, survival, or attachment factor synthesized by melanocytes (and, according to experiments using aggregationchimeras, possibly by other skin cells (Mintz, 1971)).

V. MELANIN SYNTHESIS-THE ROLE OF TYROSINASE A N D RELATED PROTEINS The pathway of melanin synthesis starts from the amino acid tyrosine (Fig. 1). The first two reactions are catalyzed by the copper-containing enzyme tyrosinase (EC 1.14.18.1). Tyrosine is hydroxylated to 3,4-dihy-

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LeucoDOPAchrome DOPAchrome

/w

DHI

I3

/

Indole-qure

DHICA >\e-quinoneca oxylic acid

CysteinylDOPA

4 4 1 Alanyl-hydroxybenzothiazine

Figure 7. The biosynthetic pathway from tyrosine to melanin (according to Hearing and Tsukamoto, 1991; Tsukamoto et al., 1992). Tyrosinase

catalyzes three different reactions in this pathway (1, 2, 3). The reaction catalyzed by the product of TRP-2, DOPAchrome tautomerase, is indicated by 4. DOPA = 3,4-dihydroxyphenylalanine; DHICA = 5,6-dihydroxyindole-2-carboxylic acid; DHI = 5,6-dihydroxyindole.

droxyphenylalanine (DOPA), and DOPA itself is oxidized to DOPAquinone (Hearing and Tsukamoto, 1991). Apart from these two starting reactions this enzyme catalyses a third reaction, the oxidation of 5,6-dihydroxyindole (DHI) to indole-quinone. The first of these reactions is the key reaction for melanin biosynthesis, because follow-up reactions can proceed spontaneously at physiological pH. Thus, tyrosinase is the rate-limiting and therefore the key enzyme for melanin biosynthesis (see below in the next chapter). Two types of melanin are found in mammals, the black eumelanin and the brown or yellow phaeomelanin. Eumelanins are derived from DOPAchrome, whereas phaeomelanins are derived from cysteinylDOPA (see Fig. 1). The two pathways diverge from DOPAquinone, and it was suggested that the availability of sulfhydryl groups influences binding of glutamine or cysteine to DOPAquinoneand thereby the switch to phaeomelanin (Jara et al., 1988; Hearing and Tsukamoto, 1991).

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Two tyrosinase-related proteins initially suspected as candidates for tyrosinase have been identified in recent years. These have been isolated as cDNA clones when antisera raised against tyrosinase were used (Shibahara et al., 1986; Jackson, 1988). Tyrosinase, TRP-1 (tyrosinaserelated protein l), and TRP-2 (tyrosinase-related protein 2) show approximately 40% amino acid homology (see Hearing and JimCnez, 1989; Jackson, 1991). These proteins share several features: they possess a transmembrane region at the carboxyterminus, two potential copper binding sites, and multiple glycosylation sites. They probably localize to the lumenal surface of the melanosome membrane, where they may form a multienzyme complex. Fifteen of the 16 cysteine residues have been conserved in all three proteins, as well as seven of eight tryptophans, and it was suggested that these residues are important for structure, stability, and enzymatic activity (see Hearing and Tsukamoto, 1991). All three proteins were reported to be pigment cell-specific (Beermann et al., 1990; Steel et al., 1992). Some expression of TRP-2 seems also to be present in the developing telencephalon and the endolymphaticduct. The meaning of this finding is not clear (Steel et al., 1992). Whereas the function and localization of tyrosinase and expression of its gene have long been well known, the function of the related proteins, TRP-1 and -2, has been studied in detail only recently. The TRP-1 gene is encoded at the mouse brown locus on chromosome 4. Mutations at this locus cause the production of brown rather than black eumelanin. When melanocytes of the immortal melan-b line, which are homozygous for brown, were infected with a retrovirus carrying and expressing TRP-1, 20-50% of the cells had a black to dark brown color characteristic of wild-type mouse melanocytes (Bennett et al., 1990). Thus, TRP-1 complements brown, which is the final proof that TRP-1 is the product of the b locus. On comparingthe TRP- 1coding region of a brown mutant mouse and a revertant from brown to black, it was shown that only one nucleotide exchange in which cysteine replaces tyrosine causes the brown phenotype (Zdarsky et al., 1990). In skin, both TRP-1 RNA and protein are more abundant than tyrosinase RNA and protein (Jackson et al., 1990;Hearing and Tsukamoto, 1991).The specific function of TRP-1 is not clear, and it has been postulated to be a melanosomal catalase, named catalase B (Halaban and Moellmann, 1990). However, different properties have been suggested by others (see Hearing and Tsukamoto, 1991), and the enzymatic function of the protein still remains to be elucidated.

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The TRP-2 gene was mapped to mouse chromosome 14 in the region of the coat color mutation slaty (Jackson et al., 1992).Homozygous s2aty mice are characterized by a change in eumelanin leading to a dark-grey fur (see Silvers, 1979). Jackson et al. (1992) could identify a point mutation in TRP-2 in mice homozygous for slaty, which makes it very likely that TRP-2 is encoded at this locus. In a parallel effort, Tsukamoto et al. (1992) used specific antibodies to isolate, characterize, and purify tyrosinase, TRP-1, and TRP-2 by immuno-affinity to finally study them with respect to their melanogenic catalytic function. As previously mentioned, tyrosinase catalyzed all three reactions ascribed to this enzyme (Fig. 1). TRP-I was only very weakly active in hydroxylating tyrosine and producing melanin. However, most surprising, TRP-2 had none of these activities but instead functioned as DOPAchrome tautomerase (EC 5.3.2.3), another enzyme critical in the synthesis of eumelanin. In the presence of this enzyme DOPAchrome is converted to DHICA (5,6-dihydroxyindolecarboxylicacid) rather than DHI (Fig. 1 ;see Hearing and Tsukamoto, 1991).The data suggestthat the mouse slaty locus encodes TRP-2 as the likely candidate for DOPAchrome tautomerase.

VI. THE M O U S E TYROSINASE GENE Mice with null mutations at the albino locus (c) have no pigment in skin, hair, or eyes, but they possess a full complement of pigment cells, so-called amelanotic melanocytes (Silvers, 1979). Some c-alleles result in pigmentation intermediate between black and white, while maintaining tyrosinase activity. It was an open question whether the c locus encoded tyrosinase or a trans-acting protein, which regulates tyrosinase activity andlor synthesis. This issue has now been resolved by appropriate rescue experiments. Tyrosinase cDNAs and genomic clones have been identified (Kwon et al., 1987b; Yamamoto et al., 1987; Ruppert et al., 1988; Porter and Mintz, 1991). The structural gene encodes five exons separated by large introns and spans a chromosomal region of about 70 kb (Ruppert et al., 1988). Comparison of the single copy structural gene and various cDNAs showed that different tyrosinase cDNA clones are generated by alternative splicing of a single transcript (Ruppert et al., 1988; Porter and Mintz, 1991). Some of these cDNAs were cloned in an expression vector and electroporated into tyrosinasenegative cells. When protein extracts were tested for tyrosinase enzyme activity it became clear that only the full-lengthtyrosinase cDNAconfers

F. BEERMANN, R. GANB, and G. SCHUTZ

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C C

5'

exon 1

2

3

4

--G G>C

5

A

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u -

A >G

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CCh

Cm

G *A

- -

figure 2. Alterations of the tyrosinase gene have been found in c-locus mutations of the mouse. In c (albino)a change from G to C at +391 causes an amino acid exchange from cysteine to serine at position 85 (see Jackson, 1991). In himalayan (ch), an exchange was found at +I 341 substituting a histidine to arginine at residue 402 (Kwon et al., 1989a). In chinchilla (cCh),a mutation at +1526 results in an alanine to threonine change (position 464; Beermann et al., 1990). The mutation chinchilla-mottled (P)is due to a rearrangement in the 5' region upstream of -5 kb (Porter et al., 1991; indicated by dashed lines). D a r k - e y e d a l b i n o ( ~is~caused ~ ~ ) by a point mutation in exon 1, leading to a serine-to-isoleucine exchange at position 128 (not shown; Schmidt and Beermann, 1994).

tyrosinase activity (Muller et al., 1988). This early experimental evidence that the c-locus does encode tyrosinase was strengthened when it was shown that cloned tyrosinase cDNA indeed maps to the c-locus region (Kwon et al., 1987b; Ruppert et al., 1988). When albino mutant tyrosinase cDNA was sequenced and compared to the wild-type (C57BL/6J) sequence, one significant difference was observed (Kwon et al., 1989b; Shibahara et al., 1990; Yokoyama et al., 1990). A point mutation in exon 1 of the mouse tyrosinase gene changes a cysteine residue into a serine (at position 85), thus leading to a nonfunctional protein (Fig. 2). This mutation is common to all tyrosinase-negative albino mice analyzed so far (Yokoyamaetal., 1990; Jackson and Bennett, 1990).Atyrosinase gene carrying this substitution is functional in neither cells (Shibahara et al., 1990) nor transgenic mice (Yokoyama et al.,

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1990). Furthermore, a melanocyte line that has reverted in culture from albino to wild type has restored the cysteine codon at position 85 (Jackson and Bennett, 1990). In vivo studies (see below) proved that the c-locus encodes tyrosinase and showed that expression of a tyrosinase minigene rescues the albino phenotype (Yokoyama et al., 1990; Tanaka et al., 1990; Beermann et al., 1990). In humans, the known genetic disorder type I oculocutaneousalbinism (OCA) is characterized by absent or reduced activity of tyrosinase (Giebel and Spritz, 1992). Whereas type IA OCA is tyrosinase-negative, type IB OCA is associated with greatly reduced tyrosinase enzymatic activity and little or no pigment at birth. DNA analyses of patients with type I OCA have established that mutations in the tyrosinase structural gene constitute the molecular basis of these disorders, and a large series of different allelic mutations of the tyrosinase gene have been identified in patients with type I OCA (King et al., 1991; Giebel and Spritz, 1992). In mouse, the situation is different. Apart from radiation-induced deletions of the c-locus (e.g., c14c0s; Russell et al., 1982) only a few specific mutations have been analyzed (albino,chinchilla, himalayan, dark-eyed albino) which are caused by point mutations in the coding region (Fig. 2; Jackson, 1991, Schmidt and Beermann, 1994). Recent results suggest that the c-locus mutation chinchilla-mottledis caused by a rearrangement in the tyrosinase gene upstream sequence that affects the level of transcription of the gene (Porter et al., 1991).

VII. EXPRESSION ANALYSIS OF THE TYROSINASE GENE IN TRANSGENIC MICE In the following chapter we will discuss recent experiments we have performed using transgenic mice (see Beermann et al., 1992b). This approach allowed us to address several questions that could not have been followed by using in v i m methods only. A. Rescue of the Albino Phenotype The albino mutation (c) is due to a point mutation in exon 1 of the mouse tyrosinase gene (Fig. 2). Thus, mice carrying mutations at the c-locus still possess a full complement of so-called amelanotic pigment cells. We therefore expected that a tyrosinase transgene would rescue the albino phenotype. Since the tyrosinase gene extends over 70 kb (Ruppert et al., 1988), we constructed a tyrosinase minigene (prrTyr4;see Fig. 3),

-3.4 kb

P PP

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P

P

exon 1

-270 bp *

ptrTyr4

exons 2-5

$

-2.25 - - kb i

P P

pt rTyr5 TyBS

exons 1-5

Figure 3. Different tyrosinase minigenes used for generating transgenic mice. The constructs ptrTy4 and 5 contain 5.5 kb and 270 bp of 5‘ sequence, first exon, first intron, and exons 2-5 fused to a SV40 splice and polyadenylation signal (open box) (Beermann et al., 1990, 1991; Kluppel et al., 1991). The construct TyBS(bottom)contains mousetyrosinasecDNA sequence, including the tyrosinase polyA signal and 2.25 kb of 5’flanking sequence (Yokoyama et al., 1990; Overbeek et al., 1991). Hindlll sites are indicated as open rectangles.

Figure 4. Rescue of the albino phenotype in transgenic mice. The photograph shows a nontransgenic albino mouse, two transgenic pigmented mice (ptrTyr4; agouti coat color), and a nontransgenic C57BV6J mouse (nonagouti black). 164

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which was injected into fertilized eggs of the albino mouse strain NMRI (Beermann et al., 1990). We obtained transgenic mice that showed pigmentation in skin and eyes (Fig. 4). This experiment conclusively demonstrates that we had rescued the albino mutation by introduction of and expression from a functional tyrosinase gene. Analysis of RNA of different organs shows that the expression of the transgene mimics the endogenous gene in retinal pigment epithelial cells and melanocytes. It also shows that the 5’ flanking sequencescontain the signals required for specific expression in the skin and expression in the neuroectoderm-derived pigment epithelium (Beermann et al., 1990). These results have been corroboratedby others using expression of a tyrosinase cDNA from the tyrosinase promoter (Yokoyama et al., 1990;Tanaka et al., 1990) but also from the metallothionein gene promoter (Mintz and Bradl, 1991). Specific expression from the tyrosinase promoter was also reported in transgenic mice c a v i n g the SV40 early region including the T antigen gene under control of 2.5 kb of tyrosinase gene sequences. These mice showed tumor formation in pigmented cells (Bradl et al., 1991a,b; Klein-Szanto et al., 1991).

A

B

Figure 5. Tyrosinase gene expression in melanocytes of the hair follicle. Bright field photographs show sections of skin of 4-day-old albino mice. Bar represents 50 pm.

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B. Identification of Sequences Required for Expression in Pigment Cells

The previous experiments showed that 5.5 kb of tyrosinase 5’ flanking sequence faithfully mimics cell type-specific expression as displayed by the endogenous gene (see Fig. 5; Beermann et al., 1990). We therefore attempted to define the sequence requirements for specific expression (Kliippel et al., 1991). A deletion series of tyrosinase-chloramphenicol acetyltransferase fusion genes was constructed and electroporated into tyrosinase-expressingand nonexpressing cell lines (see Fig. 8A). Melanoma cell-specific expression was initially obtained with constructs containing 6.1-kb and 3.7-kb upstream sequences. Further deletion showed that this expression pattern could even be obtained with as little as 270 bp of 5’ sequence. We then asked if the elements required for melanoma cell-specific expression in vitro are sufficient to direct celltype specific expression in vivo. A tyrosinase minigene was therefore constructed that contained only 270 bp of 5’ sequence (ptrTyr.5;Fig. 3; Kliippel et al., 1991; Beermann et al., 1991). Transgenic mice were obtained that were pigmented in both skin and eyes. In situ hybridization analyses confirmed cell type-specific expression from the transgene in neural crest-derived melanocytes and the pigment epithelium of the retina. The results indicate that 270 bp of the tyrosinase promoter is sufficient to specify pigment cell-specific expression. However, only a few animals showed intense pigmentation. This suggested the importance of more upstream sequences and a cell-specific enhancer has recently been identified at -12 kb of the mouse tyrosinase gene (GanB et al., 1994c; Porter and Meyer, 1994). C. Expression of the Tyrosinase Gene During Development

The temporal expression pattern of the mouse tyrosinase gene during embryonic development was not known, and therefore we exploited the

Figure 6. Transgene and tyrosinase gene expression in early eye development. Sections are derived from embryonic day 10.5 (A, B), 12.5 (C, D), 13.5 (E), and 17.5 (F) (for details see Beermann et al., 1992a,b). Transgenespecific expression (ptrTyr.5) is depicted in A, B, E, F, and tyrosinase gene expression in C and D. In A-D, the same section was photographed both in dark field (A, C) and in bright field (B, D). Bar represents 150 pm in A, B; 250 ym in C, D, E; and 200 ym in F.

F. BEERMANN, R. GANB, and G. SCHUTZ

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figure 7. Expression pattern of the mouse tyrosinase gene during embryonic development and its recapitulation in transgenic mice as determined by in situ hybridization (Beerrnann et al., 1992a). Black box, mouse tyrosinase; open box, transgene ptrTyr4; striped box, ptrTyr5. Interrupted boxes indicate variations between lines. RPE, retinal pigment epithelium; e, days of gestation; d0.5, newborn. availability of probes to investigate the temporal regulation of expression during early eye development and in the developing hair follicle of the mouse by in situ hybridization. Furthermore, we asked whether the transgenes would precisely mimic this pattern (Figs. 6 and 7; Beermann et al., 1992a). Development of the eye starts at about day 8-9 with the appearance of the optic vesicle from the forebrain. The optic vesicle then invaginates to form the bilayered optic cup between day 9.5 and day 10.0. At this stage the prospective pigment epithelium and the neuroretina are prominent. Tyrosinase gene-specific transcriptswere first detected at this stage. At day 16.5 of gestation melanocytes expressing tyrosinase mRNA were detected both in dermis and epidermis, but rarely in the developing hair follicles. The hair follicles then differentiaterapidly, and at days 17.5and 18.5 of gestation all hair follicles contain expressing melanocytes. Thus, tyrosinase mRNA is detected in melanocytes before melanin is produced and secreted into the growing hair, which emerges only after birth. This cell type-specific and temporal regulation is largely reproduced in transgenic mice carrying tyrosinase minigenes either using 5.5 kb (ptrTyr4) or 270 bp (ptrTyr-5) of 5' sequence (Figs. 6 and 7). At day 10.5 of gestation, transgene-specific transcripts derived from ptrTyr4 and ptrTyr.5 are detected in the developing pigmented epithelium of transgenic embryos. At day 16.5, melanocytes expressing ptrTyr4 are only rarely seen within developing hair follicles or the epidermis and are restricted mainly to the dermis. Expression is much more pronounced at later stages. Expression ofprrTyr.5 is first detected in melanocytes of the

~

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hair follicle at day 18.5 but is prominent only after birth. These results demonstrate temporal regulation of the tyrosinasegene in the developing eye and hair follicle and furthermore show that tyrosinase 5' sequences are sufficient to provide this cell type-specific and developmentally regulated expression. In a recent report by Steel et al. (1992) the developmental profile of expression of TRP-2 was compared with that of TRP-1 and tyrosinase. Using in situ hybridization they detected tyrosinase gene transcripts somewhat later (see above; Beermann et al., 1992a), both in retinal pigment epithelium and in melanoblasts, which might be explained by differences in the level of sensitivity. TRP-1 was found at around the time tyrosinase was expressed. They show that TRP-2 detects presumptive pigment cells at day 9.5 in the retinal pigment epithelium and at day 10.5 in melanoblasts, that is, before they can be detected with TRP-1 or tyrosinase (Steel et al., 1992). Unexpectedly, they also report on TRP-2 expression in developing forebrain and endolymphaticduct, the function of which is not known.

VIII. IN SEARCH OF PIGMENT CELL-SPECIFIC FACTORS: WHICH ELEMENTS DETERMINE EXPRESSION IN MELANOCYTES? The restricted expression of genes suggests the presence of cis-regulatory elements and trans-acting tissue-specific factors. The aforementioned studies with transgenic mice as well as gene transfer experiments (see Fig, 8A) demonstrated that the region 270 bp upstream of the transcriptional start site contains sufficient information to induce tissuespecific and developmentally regulated tyrosinase expression (Kluppel et al., 1991; Beermann et al., 1992a). To define crucial cis-acting sequences a refined deletion analysis with CAT (chloramphenicol acetyltransferase) fusion genes containing 5' flanking sequences of different extent was performed (see Fig. 8B). Expression of these fusion genes is limited to melanoma cells, and no CAT activity is detected in fibroblasts. The results obtained allow the conclusion that at least three important domains exist within 270 bp of the tyrosinase minimal promoter. Two appear to represent positively acting elements (-270 to -230 bp; -1 30 to -80 bp), whereas one sequencebetween -195 and -1 30 bp seems to harbor a negative element. A more detailed analysis revealed no cell specifity for the three elements characterized so far (GanB et al., 1994a,b).

F. BEERMANN, R. CAN& and G.SCHUTZ

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Figure 8. Characterization of pigment cell-specific gene expression. A. Determination of the transcriptional activity of tyrosinase 5' deletion sequences (-6.1 to -0.08 kb) fused to the CAT-reporter gene (Kluppel et al., 1991). B. Refinement of the deletion constructs from -270 to -80 bp (Gang et al., 1994a,b). CAT activity was corrected for luciferase activity and is given as relative activity by arbitrarily setting a CAT reporter without a promoter, pBLCAT6, to 1. Expression of the fusion genes is restricted to tyrosinase-expressing cell lines, which has been demonstrated by transfection in NlH3T3 mouse fibroblast cell lines (data not shown).

Ponnazhagan and Kwon (1992) reported a putative tissue-specific cis-element (TE-1) located at -236 bp of the mouse tyrosinase promoter. They partially purified a TE-1 binding protein (approximately 49 kDa in size), but tissue specificity remains to be confirmed by a more detailed analysis. For the human tyrosinase promoter, Shibata et al. (1992) identified a 200-bp pigment cell-specific enhancer, located between -2.0 and -1.8 kb. A minimum core sequence of 39 bp was shown to be sufficient to confer the specific activity, although other regions (not identified so far) within the 200-bp fragment are required for more efficient expression in melanoma cells (Shibata et al., 1992).

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For the TRP-I gene it has been demonstrated that only 44 bp of upstream sequence are sufficient to confer pigment cell-specific transcription in the mouse melanoma cell line B16 (Shibahara et al., 1991; Lowings et al., 1992). A detailed analysis of the TRP-I promoter using deletion, point, and linker scanning mutations identified a positive and a negative regulatory element between positions -332 and -299 together with a second positive acting &-element ( 4 3 to -34), termed M-box. None of the three characterized promoter elements confers melanocytespecific gene expression (Lowings et al., 1992). The striking feature of the M-box is the complete conservation of an 11-bpelement in the mouse and human tyrosinase promoter, which are located about 90 and 140 bp upstream of the major transcriptional start sites, respectively. The 11-bp sequence of the M-box contains a consensus binding site, CATGTG, for members of the basic helix-loop-helix family of transcription factors. A possible candidate may be the product of the microphthalmia gene locus (Hodgkinson et al., 1993). Moreover, a melanocyte-specific factor (MSF) has been characterized, which may act as an antirepressor in regulating TRP- 1 transcription (Yavuzer and Goding, 1994). This brief summary shows that our understanding of the basis of melanocyte-specific expression of the tyrosinase gene and the tyrosinase-related genes TRP-1 and TRP-2 is still limited. Novel approaches such as the introduction of the 70-kb-long mouse tyrosinase gene with extensive 5’ flanking sequences as recently achieved (Schedl et al., 1993) allow an experimental attack of this important question.

ACKNOWLEDGMENTS Thanks are due to E. Hummler and A. P. Monaghan for comments on the manuscript. We also acknowledge the contributions of S. Ruppert, E. Hummler, M. Kluppel, G. Muller, A. Schmidt, and E. Schmid to the work on the mouse tyrosinase gene. The work was supported by the Deutsche Forschungsgemeinschaft (SFB 229; Leibniz Programm), the Fonds der Chemischen Industrie and the Swiss National Science Foundation.

REFERENCES Beermann, F., Ruppert, S., Hummler, E., Bosch, F. X., Miiller, G., Riither, U., andSchiitz, G . (1990). Rescue of the albino phenotype by introductionof a functional tyrosinase gene into mice. EMBO J. 9: 28 19-2826.

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Beermann, F., Ruppert, S., Hummler, E., and Schiitz, G. (1991). Tyrosinase as a marker for transgenic mice. Nucleic Acids Res. 19: 958. Beermann, F., Schmid, E., and Schutz, G. (1992a). Expression of the mouse tyrosinase gene during embryonic development: recapitulation of the temporal regulation in transgenic mice. Proc. Natl. Acad. Sci. U S A 8 9 2809-2813. Beermann, F., Schmid, E., G a d , R., Schiitz, G., and Ruppert, S. (1992b). Molecular characterization of the mouse tyrosinase gene: pigment cell-specific expression in transgenic mice. Pigment Cell Res. 5: 295-299. Bennett, D. C.,Huszar, D., Laipis, P. J., Jaenisch, R., andJackson, I. J. (1990). Phenotypic rescue of mutant brown melanocytes by a retrovirus carrying a wild-type tyrosinase-related protein gene. Development 110 471475. Bradl, M., Klein-Szanto, A., Porter, S., and Mintz, B. (1991a). Malignant melanoma in transgenic mice. Proc. Natl. Acad. Sci. U S A 8 8 164-168. Bradl, M. L., Larue, L., and Mintz, B. (1991b). Clonal coat color formation due to a transforming gene expressed in melanocytes of transgenic mice. Proc. Natl. Acad. Sci. USA 88: 6447-645 1. Brilliant, M. H. (1992). The mouse pink-eyed dilution locus-a model for aspects of Prader-Willi-syndrome, Angelman syndrome, and a form of hypomelanosis of Ito. Mamm. Genome 3: 187-191. Brilliant, M. H., Gondo, Y., and Eicher, E. M. (1991). Direct molecular identification of the mousepink-eyed unsfublemutation by genome scanning. Science 252: 566-569. Bultman, S. J., Michaud, E. J., and Woychik, R. P. (1992). Molecular characterization of the mouse ugouti locus. Cell 71: 1195-1204. Castle, W. E., and Allen, G. M. (1903). The heredity of albinism. Proc. Am. Acad. Arts. Sci. 38: 603-621. Fleischman, R. A,, Saltman, D. L., Stastny, V., and Zneimer, S. (1991). Deletion of the c-kit protooncogene in the human developmental defect piebald trait. Proc. Natl. Acad. Sci. USA 88: 10885-10889. GanS, R., Schiitz, G., and Beermann, F. (1994a). The mouse tyrosinase gene: promoter modulation by positive and negative regulatory elements. J. Biol. Chem. (in press). GanB, R., Schmidt, A., Schutz, G., and Beermann, F. (1994b). Analysis of the mouse tyrosinsase promoter in vitro and in vivo. Pigment Cell Res. (in press). GanB, R., Montoliu,L., Monaghan, A. P., and Schiitz,G. (1994~).Acell-specific enhancer far upstream of the mouse tyrosinase gene confers high level and copy numberrelated expression in transgenic mice. EMBO J. 13: 3083-3093. Gardner, J. M., Nakatsu, Y.,Gondo, Y., Lee, S., Lyon, M. F., King, R. A,, and Brilliant, M. H. (1992). The mouse pink-eyed dilution gene: association with human PraderWilli and Angelman syndromes. Science 257: 1121-1124. Geissler, E. N., Ryan, M. A., and Housman, D. E. (1988). The dominunr-white sponing (W) locus of the mouse encodes the c-kit protooncogene. Cell 55: 185-192. Giebe1,L. B.,andSpritz,R. A. (1991). MutationoftheKIT(mast/stemcellgrowthfactor receptor) protooncogene in human piebaldism. Proc. Natl. Acad. Sci. USA 88 8696-8699. Giebe1,L. B.,andSpritz,R.A. (1992).The molecularbasisoftypeI(tyrosinase-deficient) human oculocutaneous albinism. Pigment Cell Res. Suppl. 2: 101-106.

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INDEX a-locus, 156157 product with melanocytes, 157 A-raf, 34 AADC, 77 expression in vertebrates, 59 mRNAs, 79 Aberrant c-mos expression, 140 Abl, phosphorylation of, 32 Acrosome reaction, 34 p-actin promoter, 95,96 Active MPF kinase and chromosome condensation, 25 and GVBD, 25 and spindle formation, 25 Adfl, 68-69,70,71,72 and DNA binding, 72 agouti, 156, 157 Alanyl-hydroxybenzothiazine,159 albino locus, 161 Albino mutation, 150, 163 Albino phenotype, 163, 164 Altered Ddc genes, 75 Alternative splicing, 77 Alzheimer’s Disease, 111 Amino acid decarboxylase (AADC), 57 expression, 59 in tumors, 60

Amino acid sequence comparison, 78 P-Amyloid precusor protein (APP), 110-111 and expression, 111 Aniridia, in humans, 94 Antennapedia, 89,91 Antennapedia cluster, 101 Antennapedia gene, similarity to Hoxgene, 106 Antennapedia mutation, in Drosophila, 94 Antennapedia-type homeodomains, 109 Anti-Cdc25A antibodies, 18 Antisense, c-mos oligonucleotides, 27,132,133 Antisense cycA, 24 Antisense cycB1,24 Antisense cycB2,24 APP gene in Alzheimer’s Disease, 111 ATBFl gene, 71 Autostimulatory loop, 110 Axonal projections, 64 B lymphocytes, and immunoglobulin heavy chain enhancer, 142 179

180

B-type cyclins, 10 Basic fibroblast growth factor (bFGF), 16 bicoid, 115 Binding factors, Ddc, 68-69 Biogenic amines, 56 biosynthetic pathways, 57 Bithorax, 91 cluster, 101 brown locus, 160 Budding fission, 4 C-kit, 155, 156 and melanoblasts, 155 expression, 15, 155 female, 16 proto-oncogene, 15 c-mos, 4, 26 aberrant expression, 140 and oocyte maturation, 27 and thymidine kinase promoters, 139 and zona pellucida (ZP) proteins, 139 antisense, 132, 133 as maternal message, 132 comparison to proto-oncogenes, 128 expression, 27,28 expression, in mammalian germ cells, 127-148 expression in spermatocytes, 136137 expression, somatic tissues, 129 expression, tissue specificity, 129130 expression unregulated, 144 function, in mammalian germ cells, 127-1 48 gel-shift analysis, 142 in pachytene spermatocytes, 136 in postmeiotic spermatids, 136 in postmeiotic spermatocytes, 136

INDEX

negative regulatory element (NRE), 138,141 NRE interactions with immunoglobulin heavy chain enhancer, 142 NRE tissue specificity, 142 potential function in spermatocytes, 136-137 proto-oncogene, 143 regulator of meiotic cell cycle, 129 repressor, 142 reproductive tissues, 129 RNA, 26 role in meiosis of mouse oocytes, 133 sites of transcriptions, 129, 130 somatic tissues, 130 transcription in somatic cells, negative regulation, 140-143 transcription, oocytes, 128 transient expression assays, 138 c-raf, 4 proto-oncogenes, 130 c-rafl, 34 in mouse, 34 proto-oncogene, 22 CAT fusion genes, 169 Cdc2 cell cycle, 7 dephosphorylation, 12 hypophosphorylated,24,5 kinase, 143 phosphorylated, 6 regulators, 8 CdcUCycB complex, 17-1 8 cdc25, 18 activity, 19 expression in liver, 19 murine homologue, 19 Cdc25 class of phosphatases, 30 dephosphorylation in egg extracts, 19

index

in mitosis, 12 cdk2,5,29 Cell cycle events eukaryotic cycle, 4 primary regulation, 4 regulator, 8,4, 134 Cell cycles M phase, 9 S phase, 9 Cell division control, 4 Cell transformation, 144 cfla, 68-69, 70, 71 Chimeras, 152, 153, 157 Chinchilla, 162 Chinchilla-mottledmutation, 153, 162 Chromatin, 32 cis-regulatory elements, 73, 108, 115 disadvantages and advantages, 116 in Ddc promoter, 64-68 of gene, cis-regulatory sequences, as identifier of homeobox regulation, 115 CNS monoamines, 60 Coat color, and genetic studies, 150 Colocalization, Ddc and ZFH-2, 73 Conservation of developmental genetic networks, 108 Corpus luteum, 3 1 CSF, cytostatic factor, 25-26 Cyc B mRNA, 18 zygotic transcription of, 18 cycB1 expression, 25 cycB2 expression, 25 Cyclin B, 17 as regulatory component, 24 degradation of, 135 during meiosis, 128 in oocytes, 143 in S. cerevisiae, 24

181

in testis, 25 phosphorylation of, 135 Cyclin function, during meiosis or mitosis, 28 Cyclin in early spermatids, 33 Cyclin, regulatory subunit, 6 Cyclin-dependent kinases (cdk) Drosophila, 7 goldfish, 7 humans, 7 mouse, 7 Xenopus, 7 Cyclins, 6 , 9 Cyclins A, 9, 10 and Cdc2, 10 Cyclins B, 9 Cyclins C, 9 Cyclins GI, 9 Cyclins D, 9 Cyclins E, 9 Cyclin, GI, 9 CysteinylDOPA, 159 Cytoplasmic domain, 4 Cytostatic factor (CSF), 26, 134 and calcium, 26 Cytotactin promoter, 111 regulation of, 111 D. melanogaster, 65,66,68 D. virilis, 65, 66, 68 d-locus dilute, 157 pink-eyed dilution, 157, 158 Ddc allele, 64 Ddc, and immunofluorescence,64 Ddc expression, 58,64 in the CNS, 58 Ddc genes altered, 75 glial expression, repressing function, 65 mRNA splicing, 74

182

promoter, 64 regulation in Parkinson’s Disease, 81 regulatory regions, 65 tissue-specificexpression, 65 transcriptional regulatory elements, 66 DDC comparison of amino acid sequences, 78 enzyme activity, 58 expression of, 82 hypodermal isoform, 75 immunoreactivity, 68 in vivo expression, 67 Deformed genes, 107 Dephosphorylation,5 of proteins, 2 Dfd expression, 108 Dfd protein, 108 Differentiation, germ cell, 12 Diplotene stage, 13 Dispersed homeobox genes, 92, 115 Distal enhancer, 66 Distribution of DDC protein in Drosophila, 6 1-64 Distribution of monoamines in Drosophila CNS, 6 1 Dlx genes, 92 DNA synthesis, (S) 4 Dominant spotting locus (W), 154156 DOPA, 159 DOPA decarboxylase (DDC), 56 activity, 59 DOPA decarboxylase gene (DDC), 55 DOPAchrome, 159, 161 Dopamine, 56 Dopamine cells, 63 types of, 63 Dopamine, expression of, 82 Dopamine-specific antiserum, 63

INDEX

DOPAquinone, 159 Downstream genes, homeobox gene circuitry, 109, 110 Drosophila, 18,56,57, 89, 91,92 Ddc gene, comparison to AADC gene, 76-88 Ddc gene, post-transcriptional regulation, 73-76 Ddc gene, transcriptional regulation, 64-73 DDC protein distribution, 61 Dfd expression, in mice, 108 melanogaster Ddc gene, 80 monoamines distribution, 61 opsin genes, 71 ventral ganglion, 62 Dual-specificity kinases, 10 Elfl, 68-69,70,71 and binding, 69 and lethal mutations, 70 and transcription, 70 En genes, 92,97 as target, 101 Eumelanin, 156,159 Expression of c-mos mammalian germ cells, 127-148 Expression of c-mos, oocytes, 131136 Extracellular signal regulated kinases (ERKs), 20 Eye development, 168 Female mammalian germ cells, mouse, 14 Fertilization, 34-35 Fission yeast, 4 cell cycle, 5 Gdh4 phase, 5 START cycle, 5 Function of c-mos mammalian germ cells, 127-148 oocytes, 131-136

Index

Functional equivalence, of genes between species, 105 G-protein-coupled receptors, 60 Gametogenesis,35 mammalian, 1-53 Gel-shift analysis, 142 Gene expression, hierarchy, 89 Gene, leaky expression, 101 Gene pyramid, 88,89 Gene targeting, 100 Gene transfection, 82 Genetic cascades from homeobox genes, 117 Germ cell development, stages, 13 Germ cell differentiation, 12 early, 14 male, 13 mammalian 2 migration, 14 proliferation, 14 role of c-mos, 128 Germainal vesicle breakdown (GVBD), 13,132,135 and Mos. protein, 28, 132, 133, 135 in mouse oocytes, 23 Glutamate, 76 Gsh genes, 92,101 GTP-binding protein, Ras,22 Haploid spermatids, 32 Heatshock promoter, 94 HeLa cells, 8 himulayan gene, 162 Histidine, 76 Histone phosphorylation, 33 Homeobox genes, 87-125 circuitry, 109-1 14 clusters, mouse, 90 exchanges, mammalianDrosophila, 105-109 dispersed, 92-93

183

family, 89 in Drosophila, 101 knockout, 112 modified in mice, 101 promoter exchanges, 98 proteins, 90 targeted, 100-104 targeting experiments, 102 upstream gene regulation, 112-1 14 Homeodomain genes, in mice, 110 Homeodomain proteins, as regulators of transcription, 109 Homeodomains, and protein binding, 109 Homeoprotein binding sites, human, 113 Homeotic gene complex, Drosophila, 101 Homozygous mice, 100 Hox clusters, 92,93 Hox genes, 1.5 and 1.6 mutants, 103 3.1 mutants, 103-104 alterations to generate misexpression, 98 and Hsp-70 promoter, 105 and phenotypic effects, 102 ectopic expression, 106 functional overlap of, 98 murine transgenic misexpression, 99 normal function, 99 null mutants, 101-102 promoter exchange, 98-99 relationships, 99 retinoic acid regulation, 114 role in dedifferentiated state, 100 role in disease, 111 role in morphogenesis, 111 similarity to Antennapedia gene, 106 similarity to Scr gene, 105

184

transgene construct, 98 Hox-4.2 gene, 110 Hsp-70 promoter, 105 Human amino acid decarboxylase (HUMAADC), 78 Human Cdc2 kinase, 29 Human cdc2.5 homologues, 18 hunchback, 115 Hybridization, in situ, 167 Hydroxyl radicals, 151 Immunoglobulin heavy chain enhancer, 142 Immunoglobulin heavy chain enhancer, and B lymphocytes, 142 Indole-quinone, 159 Indole-quinone-carboxylic acid, 159 INH, 20 Inhibition of cdcUCyc B kinase activity, 20 Initiator (Inr) elements, 138 Intermediate MAPK activator, 21 Intron sequences, 116 IPOU, and heterodimers, 7 1 K-2 gene, 92,93 Kinases, dual-specificity, 10 Kit, 14, 16 kit genes, 150 Knirps, 115 Krox 20, as Hox regulator, 115 Krox 20, expression of, 115 Kruppel, 115

lacZ expression, 108 Leptotene stage, 13 Lethal mutations, 70 LeucoDOPAchrome, 159

M phase, cell cycles, 9 M-box, 171

INDEX

M-Hox gene, as regulator of muscle creatine kinase gene, 112 mak, 30-31 Male germ cells, mouse, 13 Mammalian development, 87-125 pigmentation, 149- 177 transcription factors, 87-1 25 Mammalian developmental genetic interactions, 118 Mammalian genome, 88 Mammalian germ cells expression of c-mos, 127-148 function of c-mos, 127-148 Raf- 1 function, 23 Mammalian homeobox gene complexes, 91 Mammalian Hox genes, flies, 107 Mammalian-Drosophila homeobox gene exchanges, 105-109 MAP kinase (MAPK), 20,136 and re-entry of quiescent cells, 20 and microtubule-associated protein-2 (MAP-2), 20 and myelin basic protein (MBP), 20 and structural proteins, 20 mRNA, 20 signal transduction pathway, 21 signal transduction pathway, and S. cerevisiae, 21 targets, 20 Mast cell growth factor, (MGF), 15 Mat alpha 2 gene, 90Ubx protein, 90 Maternal message, 132 Maternal mRNAs, 131 Maturation promoting factor (MPF), 128,134 activation of, 134 inactivation of, 134, 135 oocyte meiosis, 135 Megacolon, in mice, 95 Meiosis, role of c-mos, 129

Index

Meiotic arrest, 25 Meiotic stages, 23-3 1 Melanin, 168 mouse, 151 Melanin synthesis, 158-161 Melanoblasts, 152, 153 and chimeras, 152 and precursor cells, 152 Melanocyte/dennis interaction, 156-158 Melanocytes, 152, 155, 168 a-locus product, 157 clonal history, 153 migration, 154 morphology, 156-158 neural crest-derived, 151 of hair follicle, 165 origin, 151-154 proliferation, 154 Melanophores, 152 Metaphase 11-arrestingMos, 28 Mice, S t / S f , 15 mikl, 10 Misexpression studies, 94-100 Mitogen-activatedprotein (MAP) kinase, and Mos, 135-136 Mitogens, and Raf-l,22 Mitosis (M), 4, 11 onset, 6 Mitotic stages, 16-23 M015,29 as a negative regulator, 29 isoIation, 29 Moloney sarcoma virus, 128 Monoamines, 60 Morphogenetic differentiation, 33 Mos, 26 and autophosphorylation,28 and meiosis, 133-134, 136 and phosphorylation, 27 expression, 143 function, 35 in oocytes, 143

185

kinase, role in oocyte maturation, 27 MPF stabilization, 28 regulator of meiosis, 136 toxicity, 130 Mouse c-mos gene, 138 FSH, 17 genome, 10 male germ cell, 13 oocytes, c-mos transcription, 137-140 oocytes, expression of c-mos, 131 oogenesis, 13 pgene, 158 spermatogenesis, 13 tyrosinase gene, 161-163 Mox genes, 92 MPF (M phase promoting factor), 23-24 and oocytes, 24 MPF activity, 9 msh genes, 92, 101 Murine c-abl, 32 Murine cycBI, 33 Murine cycBI, expression in testis, 33 Murine germ cell development, 25 Murine homeobox genes, 87-125 Murine rux 33 Murine transgenic misexpression studies, 99 Mutations extension, 156 nonagouti, 156 paired box genes, 93 Myelin basic protein (MBP), 20 Negative regulatory element W E ) , 141 Nekl ,30-3 1 and corpus luteum, 31 Neural AADC mRNAs, 79

186

Neural cell adhesion molecule (NCAM), regulation of, 111 Neural crest, 151, 153 analyses, 151 and melanocytes, 151-154 Neural crest cells infection with tryosinase-express ing retrovirus, 153-154 pathways, 152 Neuroblastoma cells, 79 NIH 3T3 fibroblasts, 133 NIMA protein kinases, 3 1 Non-neural AADC mRNAs, 79 Non-receptor tyrosine kinases, 4 cytosolic families, 4 transmembrane, 4 Norepinephrine, 60 Northern blots, 95 Novel kinases, 30 NTFl, 70 Null mutants, 100 Oculocutaneous albinism (OCA), 158,163 and tyrosinase, 163 Okadaic acid, 19 Oocyte c-mos promoter, 139 Oocytes, 12, 13, 128 c-mos function and expression, 131-1 36 c-mos transcription, 128 maturing, 13 Oogenesis, mouse, 13 Ovarian germ cells, 129 Pachytene stage, 13 Paired box sequence, 93-94 and DNA binding, 93 and mutation, 93 Parkinson’s disease, 8 1 Pax genes, 92,93 PDGF receptor, 22 Phaeomelanin, 156, 159

INDEX

Phenotypic analysis, dominant mutants, 97 Phenylalanine, 76 Phosphatases, 18 Cdc25,12 in mammalian gonadal function, 19 Phosphorylation, 5 and cell cycle regulation, 19 of proteins, 2 Pigment cell-specific gene expression, 170 Pigment cells, 165 Pigment epithelium, 167 Pigmentation genes, 150 Pigmentation, regulation of during mammalian development, 149-177 Pigments, and hydroxyl radicals, 151 Pink-eyed mutation, 150 Pkg promoter, 110 Polar body extrusion, 133 Polyadenylation of c-mos, 131 Polymerase chain reaction (PCR), 112 Post-meiotic events, 31-34 Post-transcriptional regulation, Drosophila Ddc gene, 73-76 POU proteins, 70 POU-domain family of transcription factors, 70 Pre-leptotene stage, 13 Primordial germ cells (PGC), mouse, 14 Progesterone, and protein phosphorylation, 23-24 Prokaryotic repressors, 89-90 Protein expression, during mammalian gametogenesis, 1-53 Protein function, during mammalian gametogenesis, 1-53 Protein kinase activities, 4

Index

Protein kinase regulators, 11 Protein phosphorylation, 12 history, 12 Proto-oncogenes, 128 Proximal promoter, 65 Pseudohomeodomain, 7 1 Pyridoxal-dependentdecarboxylases, 76 Quinone metabolism, 58 Raf, 22-23 Raf serindthreonine kinase, 22 Raf- 1 and PDGF receptor, 22 and signal transduction, 22 expression in testis, 23 Receptor tyrosine kinase (RTK), 3, 22 Recognition helix, 90 Regulation of c-mos transcription, mouse oocytes, 137-140 Regulation of DOPA decarboxylase gene, 55-86 Regulators, Cdc2 protein kinase, 11 Retinal pigment epithelium, 169 Retinoic acid, as Hox gene regulator, 114 Rhombomeres, 114 S phase, cell cycle, 9 S. pombe, 11 Scr gene, similarity to Hox gene, 105 Serine kinases, 4 Serindthreonine protein kinases, 20 Serotonin, 56 Serotonin cells, 63 DDC expression in, 68 in grasshopper, 72 Serotonin, expression of, 82, Sertoli cells, 17 mutant, 17

187

treated with FSH, 17 Signal transduction cascade, 23 Signal transduction pathways, 20 silver (si) locus, 158 Sl expressing cells, 16 SZ mutation, 15 Small eye mutation, in mouse, 94 Somatic cells, negative regulation ofc-mos, 140-143 Somatic tissues, c-mos expression, 129,130 Spermatocytes, 136- 137 Spermatogenesis,mouse, 13 Spermiogenesis,32 Splotch mutation, 94 Steel (Sl), 15 Steel (Sl) locus, 154-156 c-kit, 154 Steel, 14, 16 Steel factor (SLF), 15 Stem cell factor (SCF), 15 string expression, 30 string transcripts, 30 string mRNA, 18 SV40 DNA, 95 SV40 large Tantigen, 112 SV40 promoter, 138 T antigen gene, 165 Target gene expression, 88 Targeted homeobox genes, 100-104 Targeted Hox mutations, 104 TDC, Tryptophan decarboxylase,78 Temporal regulation, 168 Terminal differentiationproducts, 88 Testicular germ cells, 129 TH, expression of, 82 Threonine kinases, 4 Tissue specific expression, 129 Tissue specificity, c-mos expression, 129 Tissue-specific c-mos expression, 142,143

188

Tissue-specificcis-element, 170 trans-acting factors Ddc expression, 68 DNA binding, 68 Transcription factors, mammalian development, 87-1 25 Transcriptional regulation, 64-73 Transfection, cdk2,8 Transgene, expression, 166- 167 Transgenic animals, 116 advantages, 116 disadvantages, 116 Transgenic flies, heat shock induction, 106 Transgenic mice, 154, 155, 167 tyrosinase gene, 163, 164 Transient expression assays, 110 TRP-1,171 Tryptophan, 76 Tryptophan decarboxylase (TDC), 78 Tryptophan, decarboxylated,56 twine expression, 30 twine transcripts, 30 Two-step gene targeting, 104 Tyrosinase, 158, 159, 161, 168 Tyrosinase cDNA, 165 Tyrosinase gene alterations, 162 development, 167 expression, 165, 166-167 expression in transgenic mice, 163-1 69 hydridization, 169 mouse, 161 163 Tyrosinase in melanin synthesis, 158-161 minigenes, 163, 164 promoter, 169 sequencing, 167

INDEX

Tyrosinase-chloramphenicolacetyltransferase, 167 Tyrosinase-related protein 1 (TRPI), 160 Tyrosinase-related protein 2 (TW2), 160,161 Tyrosine, 76 decarboxylated, 56 Tyrosine hydroxylase (TH), 63 Tyrosine phosphorylation, 35 Upstream genes, homeobox gene circuitry, 109 v-kit, 156 v-mos, 128 Vertebrate AADC gene, 76-88

Wlocus, 154 W mutation, 15 Waardenburg’s syndrome, 94 Wee 1 gene, 10 weel, 10 Xenopus oocytes, 12, 13 @-2,71 ZFH-2,68-69,70,7 1,72 expression profile, 72 Zinc finger gene, Krox 20, 115 Zinc fingers, 71 Zona pellucida (ZP) binding sites, 34 mouse, 34 promoters, 139 glycoprotein (ZP3), 34 Zygote, development, 88 Zygote DNA, 117 Zygotene stage, 13 Zygotic transcription of Cyc B, 18

Advances in Neural Science Edited by Sudarshan Malhotra, Department of Zoology, University of Alberta Volume 1, 1993,228pp. ISBN 1-55938-356-9

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