Activins as Regulators of Branching Morphogenesis

Developmental Biology 238, 1–12 (2001) doi:10.1006/dbio.2001.0399, available online at http://www.idealibrary.com on

REVIEW Activins as Regulators of Branching Morphogenesis Emma M. A. Ball 1 and Gail P. Risbridger Centre for Urological Research, Monash Institute of Reproduction and Development, Monash University, Melbourne, Victoria, Australia

Development of glandular organs such as the kidney, lung, and prostate involves the process of branching morphogenesis. The developing organ begins as an epithelial bud that invades the surrounding mesenchyme, projecting dividing epithelial cords or tubes away from the site of initiation. This is a tightly regulated process that requires complex epithelial– mesenchymal interactions, resulting in a three-dimensional treelike structure. We propose that activins are key growth and differentiation factors during this process. The purpose of this review is to examine the direct, indirect, and correlative lines of evidence to support this hypothesis. The expression of activins is reviewed together with the effect of activins and follistatins in the development of branched organs. We demonstrate that activin has both negative and positive effects on cell growth during branching morphogenesis, highlighting the complex nature of activin in the regulation of proliferation and differentiation. We propose potential mechanisms for the way in which activins modify branching and address the issue of whether activin is a regulator of branching morphogenesis. © 2001 Academic Press Key Words: branching morphogenesis; organogenesis; embryogenesis; activin; inhibin; follistatin; TGF␤; epithelium; mesenchyme; stroma.

INTRODUCTION Activins and inhibins are members of the TGF␤ superfamily that regulate growth and differentiation via both autocrine and paracrine mechanisms. They perform diverse roles in different tissues and often have opposing effects. Activins were originally isolated as gonadal regulators of follicle-stimulating hormone (Ling et al., 1986), although mesoderm induction was the first main role identified for activin during development (Albano et al., 1990; Smith et al., 1990; Thomsen et al., 1990; van den Eijnden-Van Raaij et al., 1990; Mishina et al., 1995; Oda et al., 1995; Gu et al., 1998; Weinstein et al., 1998). More recently, roles for activins were identified in bone morphogenesis (Merino et al., 1999), the establishment of both anteroposterior and left–right axes (Mitrani et al., 1990; Thomsen et al., 1990; Kondo et al., 1991; Mathews et al., 1992; Oda et al., 1995; Oh and Li, 1997; Nomura and Li, 1998; Waldrip et al., 1998; Song et al., 1999; Kim et al., 2000), and branching morpho1

To whom correspondence should be addressed at Monash Medical Centre, Monash Institute of Reproduction and Development, 246 Clayton Road, Clayton, Vic 3168, Australia. Fax: ⫹61 (3) 9594 7115. E-mail: [email protected]. 0012-1606/01 $35.00 Copyright © 2001 by Academic Press All rights of reproduction in any form reserved.

genesis of glandular organs (references in relevant sections below). The purpose of this review is to examine the evidence that supports the hypothesis that these growth factors modulate branching morphogenesis in the kidney, prostate, and other branched organs. The overview of this topic is necessarily concise and the reader is directed to key references for more detailed information. The patterns of cell– cell interactions during development and the expression of activin-signaling components are compared in different branched organs. Evidence that activins have functional or biological effects (blocking endogenous activin action or addition of exogenous activins) is examined. Finally, the impact of knockout and transgenic models on elucidating the action of activin signaling in vivo during embryogenesis and organogenesis is reviewed.

ACTIVIN BIOLOGY Activins and inhibins are dimeric proteins (Fig. 1). Activin consists of two ␤ subunits, ␤A and ␤B, that form the hetero- or homodimers, activin A (␤A:␤A), activin B (␤B: ␤B), or activin AB (␤A:␤B). Inhibin consists of an ␣ subunit

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FIG. 1. Schematic diagram of activin and inhibin dimers. Activin isoforms consist of two ␤ subunits linked by disulfide bonds. The nomenclature of activin dimers is such that ␤A:␤A is designated activin A, ␤B:␤B is activin B, and ␤A:␤B is activin AB. Inhibin isoforms consist of a common ␣ subunit linked to a ␤ subunit. The ability of ␤C, ␤D, or ␤E subunits to dimerize with other ␤ or ␣ subunits is currently unknown.

that dimerizes with one of the activin ␤ subunits to form either inhibin A (␣:␤A) or inhibin B (␣:␤B). Three other ␤ subunits (␤C, ␤D, and ␤E) have the potential to form new activins and inhibins (Hotten et al., 1995; Oda et al., 1995; Fang et al., 1996) but the existence of all the possible dimer combinations remains to be confirmed. Activin ␤C subunit did not dimerize with the ␣ subunit to form inhibin C in vitro (␣:␤C), although it dimerized with the other ␤ subunits, ␤A and ␤B (Mellor et al., 2000). Activins signal through membrane-bound receptors as shown in Fig. 2 (see recent reviews: Heldin et al., 1997; Datto and Wang, 2000; Itoh et al., 2000; Miyazono, 2000; Weinstein et al., 2000; Wrana and Attisano, 2000). The activin receptors share homology with the receptors for other TGF␤-superfamily ligands. In vitro studies demonstrated an interaction between the activin ligands and activin receptors and between the TGF␤ ligands and TGF␤ receptors. Bone morphogenetic proteins (BMPs) signal through activin receptors (Yamashita et al., 1995) and activin and TGF␤ receptor subunits form complexes in vitro (ten Dijke et al., 1994), making it difficult to predict the outcomes and significance of ligand bioactivity. Furthermore, because the pathways converge, it is theoretically possible that different ligands signal through the same set of Smads, resulting in the same bioactivity. However, subtle discrete differences in the recruitment and activation of different receptors and Smads lead to distinct biological activities (Dennler et al., 1998; Labbe et al., 1998; Dennler et al., 1999; Zhu et al., 1999). It remains to be determined how the different combinations of ligands, receptors, and Smad proteins lead to distinct biological effects and whether there are some combinations that lead to the same biological outcomes. One of the key features to distinguish activin signaling from TGF␤ and other members of the superfamily is that activin ligand access to the receptors

can be blocked by follistatin (FS) or inhibin (Nakamura et al., 1990; Xu et al., 1995; de Winter et al., 1996; Iemura et al., 1998). The issue of activin binding proteins such as follistatin is another complicating factor and, in interpreting the data described in this review, one must also bear in mind that follistatin can also bind other members of the TGF␤ superfamily such as the BMPs, albeit at much lower affinity than that of activin (Fainsod et al., 1997; Iemura et al., 1998). The interaction between follistatin and activin has a significant effect on the biological action of activin. Follistatins are glycoproteins that bind activins, neutralizing their activity. Follistatins are highly conserved between species and are structurally unrelated to the activin and inhibin subunits (Phillips and de Kretser, 1998). Differential mRNA splicing generates several functional isoforms, such as the ECM-bound FS-288 and the secreted FS-315 (Sugino et al., 1993). Another associated protein, follistatin-related protein (FSRP), was shown to bind activin

FIG. 2. Schematic diagram of activin signaling. The extracellular activin ligand binds to the activin type II (ActRII A or B) receptor. The activin type I (ActRI A or B) receptor is recruited and phosphorylated by the ActRII. This then triggers a signaling cascade mediated by intracellular Smad proteins. After phosphorylation of ActRI, Smad-2 or Smad-3 (pathway-restricted, receptor-regulated Smads or R-Smads) bind to the type I receptor in the cytoplasm and are phosphorylated. Heteromeric complexes consisting of two types of Smads are formed. They then bind Smad-4 (common Smad or Co-Smad) and translocate with Smad-4 to the nucleus. The nuclear Smad complex binds to Fox HI (Fast-1) DNA-binding protein forming an “activin-responsive factor” (ARF). The ARF binds to activin-response elements (AREs) in promoters to regulate activin-responsive genes.

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Activins and Branching Morphogenesis

and was localized to the nuclear membrane (Prof. Alan Schneyer, personal communication) but its function is currently unknown.

a role of activin as a universal regulator of branching morphogenesis.

Kidney

ACTIVINS AND BRANCHING MORPHOGENESIS Branching morphogenesis is a distinctive process that follows a repetitive pattern common to branched glandular organs. Organogenesis of branched glandular organs begins by the budding of preexisting epithelial structures into the surrounding mesenchyme. Arborization of glandular epithelia occurs by bifurcation of termini, producing two tips and to a lesser extent, by formation of new branches from the shaft of older established branches (reviewed in Davies and Davey, 1999). During branching, differentiation and ductal canalization occur in a proximal-to-distal manner. Signals from the mesenchyme initiate and modulate branching events in the developing gland. Branching is a highly complex three-dimensional process that is tightly regulated by cell– cell communications, cell–matrix interactions, and signal transduction pathways, resulting in the ordered fate-determination, remodeling, proliferation, and/or differentiation of specific cell populations. Opposing gradients of inhibitory and stimulatory growth factors are postulated to tightly regulate branching. Geometric features such as the length along the ducts, the angle of branching, and the number of branches are all highly coordinated. Subtle alterations in any of these features would have profound effects on the final number of ducts, the spatial organization, and the overall shape and function of the fully developed organ. The importance of activin action during mesoderm induction, bone morphogenesis, and left–right asymmetry formation is consistent with previous studies showing that activins are pleiotrophic proteins with diverse effects over a range of tissues. Activins are key molecules required during early embryogenesis and organogenesis, where they provide information to cells on their spatial position and developmental fate. Given activin’s role in spatial organization during development, and by analogy to other members of the TGF␤ superfamily, activin is a candidate factor for a role in regulating the formation of complex branched organs. In Drosophila larvae, the development of the highly branched tracheal system is regulated by activin/TGF␤. In this system the activin/TGF␤ homologue, decapentaplegic (Dpp), signals through its receptor, thickveins (tkv). Dpp controls cell migration out of the tracheal placode followed by formation of the tracheal system, independent of its role in embryonic patterning (reviewed in Hogan and Yingling, 1998; Metzger and Krasnow, 1999). The indirect, direct, and correlative lines of evidence that activin is involved in more complex mammalian branching morphogenesis are discussed below. For clarity, each organ is considered separately, followed by general discussion of the evidence for

Signaling between the metanephric mesenchyme and the ureteric bud induces kidney development. The ureteric bud forms as an outgrowth from the Wolffian duct [embryonic day (ED) 11 in the mouse] (Barasch et al., 1997). For recent reviews on collecting duct morphogenesis see Davies and Davey (1999), Pohl et al. (2000b), and http://cpmcnet. columbia.edu/dept/genetics/kidney/ for time-lapse movies of ureteric bud growth of in vitro organ cultures (Srinivas et al., 1999). The strongest evidence for a role of activins in branching morphogenesis emerged from studies in the kidney. Activin is expressed during kidney development, in vitro it reduces branching of kidney explants, and in vivo disruption of ActRII caused a range of renal abnormalities. In the developing kidney, several components of the activin-signaling pathway are expressed: the ␤A and ␤B subunits, the type II activin receptors, and follistatin (Hilden et al., 1994; Roberts and Barth, 1994; Tuuri et al., 1994; Ritvos et al., 1995). Indirect evidence demonstrated that blocking activin action induced branching tubulogenesis in Madin–Darby canine kidney (MDCK) cells, with either transfection of a dominant-negative ActRII or addition of exogenous follistatin (Kojima et al., 2001). Further, mouse kidney rudiments cultured in the presence of activin showed delayed ureteric epithelial branching: the distances between branches and the number of secondary branches were diminished (Ritvos et al., 1995). Follistatin inhibited the effect of activin but inhibin A had no effect on branching in kidney explants (Ritvos et al., 1995). This illustrates the importance of the interplay between activin ligand and follistatin binding protein, rather than between activin and inhibin ligands during branching. The mechanism of action of activin A during kidney development involves other growth factors, for example, hepatocyte growth factor (HGF). HGF is required for the initial budding event and branching. Following treatment with HGF in vitro, branching tubulogenesis was stimulated by downregulation of activin A (Maeshima et al., 2000b). This effect was blocked by follistatin. Although activin A is considered a downstream effector of HGF, the details of the cascade are unknown. Data generated in vivo support the in vitro data that demonstrate an inhibition of branching morphogenesis by activin. Mice expressing a dominant-negative ActRII, preventing activin signaling, displayed an increase in the number of glomeruli in vivo (Maeshima et al., 2000a). Overall, in the kidney there is reasonable evidence that activin is expressed during the development of the organ, that it has a biological effect both in vitro and in vivo, and that its mechanism may involve other growth factors such as HGF.

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Prostate Development of the rodent prostate begins late in gestation when epithelial budding occurs as a result of intercell signaling in the urogenital sinus between the mesenchyme and epithelium (Sugimura et al., 1986; Hayashi et al., 1991; Timms et al., 1994). Initiation of budding in the developing prostate is androgen regulated (Cunha et al., 1987, 1995; Donjacour and Cunha, 1988) but because androgen receptors are located in the mesenchyme, the epithelium responds indirectly via stromal signals (Cunha, 1972; Cunha and Young, 1991; Takeda and Chang, 1991). Candidate factors responsible for signaling events between the epithelium and mesenchyme include members of the FGF and TGF␤ superfamily. Evidence of a role for activins in prostatic development is preliminary. Activin production in the adult prostate is androgen regulated (Risbridger et al., 1996); it is not known whether activin synthesis is androgen regulated during development. Whereas the adult prostate produces ␣, ␤A, and ␤B subunits, during development ␤B or ␣ subunit production is negligible (Cancilla et al., 2001). The activin ␤A subunit was localized to the undifferentiated mesenchymal cap adjacent to prostatic epithelium (Cancilla et al., 2001), consistent with a role for activin A ligand in branching morphogenesis. Functional studies in which activin A was added to cultured prostatic explants in the presence and absence of testosterone showed an inhibition of branching morphogenesis (Cancilla et al., 2001). Fewer branches developed and epithelial tips and surrounding mesenchyme were expanded. Although activin expression and action in vitro were demonstrated during branching morphogenesis of the prostate, there are currently no data to implicate inhibins. In contrast, the activin-binding proteins (follistatins) are powerful modulators of activin action. Follistatin induced proliferation and branching of the organ as well as differentiation of smooth muscle surrounding the epithelial ducts (Cancilla et al., 2001). Therefore the interplay between activin and follistatin, rather than activin and inhibin, is implicated in these studies on prostate, similar to those in kidney. There are several mouse models to study the effects of a disruption of activin signaling, for example, the ␤A subunit and activin receptor knockout models and mice expressing dominant-negative activin receptors (Matzuk et al., 1995a, b; Song et al., 1999; Maeshima et al., 2000a). No prostate phenotypes were reported in these models, suggesting that prostate development occurred, but either phenotypic alterations do not exist or closer examination is required to determine whether the number of branches, length of branches, and other features of prostate growth are changed. At this stage the detailed studies necessary to identify these changes have not yet been performed. Therefore, in the prostate, the expression and in vitro data provide preliminary evidence of a role for activins during development. Whether the mechanism of activin

action in the prostate is similar to that in the kidney is unknown.

Mammary Gland Mammary gland development begins with the extension of a duct from the nipple. Branching increases dramatically with the onset of ovarian function. Unlike other organs discussed in this review, full development of the rodent mammary gland occurs later in life; however, the processes are similar to those witnessed in other ductal organs (reviewed in Silberstein, 2001). Hormones such as glucocorticoids, progesterone, estrogen, prolactin, and other factors are required for mammary alveolar outgrowth (Topper and Freeman, 1980). The localization of activins to the developing mammary gland is largely unknown, but a role for activins is implied because the adult gland and some mammary epithelial cell lines expressed activin and activin receptors types I, II, and IIB (Ying et al., 1995; Liu et al., 1996; Ying and Zhang, 1996). Functional studies demonstrated that addition of exogenous activin A inhibited HGF-induced growth of mammary epithelial and tubule formation (Liu et al., 1996). Although there are very little data on the role of activins, preliminary studies reveal an association between activin A and HGF in mammary development. The question remains as to whether this paradigm is similar to that of the kidney. The in vivo studies using transgenic mice with various deletions of genes involved in activin signaling implicate activins in the branching morphogenesis of mammary epithelium. Rather than demonstrating a role in inhibition of branching morphogenesis, tissue recombination techniques demonstrated a requirement for activin in mesenchyme. Mice lacking the activin ␤B subunit showed abnormal mammary ductal elongation and alveolar morphogenesis during puberty and pregnancy (Robinson and Hennighausen, 1997). Therefore in mammary development, the inhibitory role of activin A in vitro contrasts with the apparent requirement for the ␤B subunit in vivo. One explanation for the discrepancy may lie in the multiple roles of individual activin subunits in forming different activin and inhibin dimers (Fig. 1).

Lung Murine lung development commences with the primary lung buds budding from the ventral foregut at ED9.5 followed by dichotomous branching events. Localization of activin and its downstream effectors suggests a role of activin in lung development. Activin ␤A and ActRIIB mRNAs were expressed in rat bronchiole tubes, whereas follistatin was expressed in the mesenchyme (Roberts and Barth, 1994). Similarly, in the baboon lung, type I and II activin receptor transcripts were expressed and developmentally regulated (Zhao et al., 1996). In mouse, both ActRI and RIB were highly expressed (Verschueren et al., 1995) and Smads -2, -3, and -4 were detected predominantly in the

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distal epithelium (Zhao et al., 1998). Abrogation of Smads -2 and -3 together, and Smad-4 alone, stimulated lung branching morphogenesis in culture, resulting from blockade of activin and/or TGF␤ ligand signaling (Zhao et al., 1998). Functional data generated in embryonic rat lung explants suggested that TGF␤, rather than activin A, regulated pattern formation in the developing rat lung because activin A had no effect in this system (Liu et al., 2000). However, activin A stimulated fibroblast proliferation and differentiation in human lung fibroblasts (HFL1 cells) by stimulating ␣-smooth muscle actin expression (Ohga et al., 1996). Under three-dimensional culture conditions, activin A enhanced fibroblast-mediated contraction, the effect of which was abolished by follistatin (Ohga et al., 2000). These data imply a role for activin A in pulmonary connective tissue morphogenesis. In vivo, mice deficient in follistatin were defective in lung function, although the specific details of this abnormality are unknown (Matzuk et al., 1995c). Alone, the in vivo effects of follistatin deficiency do not rule out an effect via BMPs and in vitro it is difficult to distinguish between the effect of TGF␤ and activin by altering Smad function. However, follistatin does not interact with TGF␤, and BMPs signal through a different set of Smads. Therefore, overall the data do not exclude activin in lung morphogenesis, as demonstrated by the in vitro studies using activin A.

Pancreas Pancreas development begins with specification of the foregut endoderm followed by the budding of digestive tract epithelium. The pancreas consists of two main components, the endocrine islet cells (secreting internally into the circulation) and exocrine acini (secreting outwardly via a lumen). Transcripts of the type II activin receptor and both ␤A and ␤B activin subunits were detected in the developing human pancreas (Hilden et al., 1994; Tuuri et al., 1994). In the rat, activin A protein was detected in primitive islet cells (Furukawa et al., 1995) and in the mouse, ␤B, ActRI, ActRII, and ActIIRB mRNA were localized to the pancreatic epithelium and islet cells (Ritvos et al., 1995; Yamaoka et al., 1998; Maldonado et al., 2000). Initially, follistatin was expressed in the mesenchyme and later downregulated in the murine pancreas (Miralles et al., 1998; Maldonado et al., 2000). In cultured pancreatic rudiments in vitro differentiation of exocrine components occurred in the presence of mesenchyme, whereas in epithelial rudiments cultured in the absence of mesenchyme, differentiation of endocrine cells was favored (Miralles et al., 1998). In vitro activin A inhibited branching and elongation of epithelia in cultured mouse pancreatic rudiments, blocking development of the exocrine pancreas. Follistatin alone did not affect the branching process but counteracted the effects of activin (Ritvos et al., 1995). In epithelial rudiments cultured in the absence of mesenchyme, exogenous follistatin mimicked the effect of mesenchyme by inducing exocrine and repress-

ing endocrine development (Miralles et al., 1998). Altogether, the pattern of expression and the functional studies show that follistatin blocks activin and hence allows exocrine differentiation while suppressing endocrine differentiation. Later, the subsequent downregulation of follistatin allowed activin-mediated endocrine cell differentiation and formation of islets. In vivo a requirement for activin in endocrine pancreatic development is implied because two independently generated mouse models expressing a dominant-negative ActRII, blocking activin signaling, showed evidence of pancreatic islet hypoplasia and various abnormalities of endocrine cells (Yamaoka et al., 1998; Shiozaki et al., 1999). In mice expressing the transgene in all cell types, the exocrine epithelial acinar cells were disrupted (Shiozaki et al., 1999); therefore, there was consistency between the in vivo and in vitro data that demonstrated inhibition of pancreatic branching morphogenesis and development. As in the kidney, there is reasonable evidence that activin is expressed in the pancreas during development and that it has a biological effect both in vitro and in vivo.

Salivary Gland The salivary gland develops with budding of the epithelium into surrounding mesenchyme in a manner similar to that of other glandular organs. Expression of activin ␤A and ␤B was detected in the developing human, mouse, and rat salivary gland (Tuuri et al., 1994; Ritvos et al., 1995; Blauer et al., 1996). In the mouse, expression of the activin ␤A subunit was further localized to the smooth muscle cells and in the rat, it was expressed in the ductal cells (Ritvos et al., 1995; Blauer et al., 1996). It is not clear whether these differences in localization reflect interspecies variation, differences in the detection techniques, or the time of investigation. Follistatin mRNA was expressed in human salivary gland (Tuuri et al., 1994): FS protein was localized to mouse mesenchymal cells surrounding the branching acinar epithelia as well as to the epithelium itself (Ritvos et al., 1995). ActRII mRNA was detected in human, mouse, and rat (Hilden et al., 1994; Roberts and Barth, 1994; Ritvos et al., 1995) and ActRIB in mouse (Verschueren et al., 1995), although the location of the receptors themselves is not yet known. Activin A inhibited salivary gland branching morphogenesis (Ritvos et al., 1995). Specifically, rudiments of embryonic mouse salivary gland showed fused adenomeres and disappearance of interlobular clefts, so that globular bulges appeared in individual lobules after treatment with activin A (Ritvos et al., 1995). Abnormalities in salivary gland function or appearance in vivo have not been reported using any of the animal models with perturbations in activin signaling. Therefore, as in the prostate, there is preliminary evidence that elements of the activin-signaling pathway are present and activin A has an inhibitory effect in vitro, but little else in known.

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CONCLUSIONS Based on the data reviewed in the preceding sections, we conclude that there is sufficient evidence for a role of activins in branching morphogenesis. The activin subunits or proteins are expressed at appropriate times and locations during branching and, in many instances, there are functional data to demonstrate that activins regulate branching morphogenesis. Limited studies from knockout mice show functional consequences of targeted disruption of activin and associated signaling pathways. However, these results cannot be consolidated into a single concept or effect of activins on branching morphogenesis in general. In terms of the localization and expression of activins in each of the organs considered in this review, there was no obvious pattern whereby expression was restricted to epithelia or mesenchyme. Instead, several studies reported activin subunit and follistatin expression in both cell types. There was an apparent predominance of activin ␤A subunit expression in prostate and lung epithelia, whereas ␤B was the main activin subunit expressed in salivary and pancreatic epithelia. Follistatin expression was detected in both epithelia and mesenchyme and there was a common lack of inhibin ␣ subunit expression in branched organs. The patterns of expression were consistent with functional data and emphasized the interplay between activin and follistatin, rather than between activin and inhibins. The functional in vitro experiments yielded consistent results across most organs studied. The addition of exogenous activin A inhibited branching morphogenesis and both ductal branch formation and elongation were reduced, suggesting that activin acted to constrain the degree of branching and govern the final number and threedimensional structure of the ducts (Maeshima et al., 2000a). A notable exception to these findings was lung, in which activin A stimulated branching morphogenesis (Ohga et al., 1996, 2000). Almost exclusively, activin A was the ligand used in these studies and little is known about the effects of the other activin homo- or heterodimers. It would be appropriate to compare the effects of activin A with activin B and AB, especially in pancreas and salivary gland where localization data implicate ␤B subunit homo- or heterodimeric ligands (Ritvos et al., 1995; Blauer et al., 1996; Maldonado et al., 2000) and to determine whether other activin dimers stimulate branching in the lung as reported for activin A. Generally, follistatin blocked the effects of activin A but the effects of other activin-binding proteins, for example FSRP, are unknown and require examination to determine whether the interplay between activin and follistatin includes other binding proteins. The functional consequences of targeted disruption of elements of the activin-signaling pathway in vivo showed abnormalities consistent with an inhibitory role for activins as shown in vitro, at least in the kidney and pancreas. In other organs the consistency between in vivo and in vitro studies was not determined because the mouse models were not exam-

ined in sufficient detail or were embryonic lethal (Sirard et al., 1998; Weinstein et al., 1998). The construction of new mouse models will contribute to this area of knowledge. The use of technologies such as the cre-lox system with temporal and tissue-specific promoters will enable dissection of the effects of reducing or increasing expression of the gene of interest and overcome the problems associated with the embryonic lethality of some of the existing models (Sirard et al., 1998; Weinstein et al., 1998). Complex breeding strategies between existing models will enable dissection of multifaceted phenotypes and eliminate problems associated with redundancy between proteins. Better tools of analysis will also contribute to the generation of data from existing and future animal models. Assessment of subtle phenotypic changes in animal models was hindered by a lack of adequate description of the “normal” developmental pattern of branching organs. This requires construction of detailed three-dimensional spatiotemporal maps. Confocal microscopy of whole-mounted microdissected tissues can be used to digitally reconstruct and analyze three-dimensional structures. The digital images can be converted to numerical data and mathematical algorithms of branched organs constructed to allow documentation and comparison of features such as branch number or duct diameter. Lack of appropriate techniques of analysis may also explain, at least in part, why so few changes in branching morphogenesis were detected in the branched organs from animal models carrying defects in the activin-signaling pathway. Limited studies described upstream regulators and downstream effectors of activin A in different branched organs. Activin A did not induce apoptosis in the developing kidney, pancreas, salivary gland, or prostate (Ritvos et al., 1995; Cancilla et al., 2001), although the mechanisms by which activin inhibited branching morphogenesis are speculative. There is insufficient information regarding factors that regulate activin A synthesis and action. Using the kidney and prostate as examples, a preliminary model for the action of activin A in branching of these organs is presented in Fig. 3. In the kidney, and perhaps the mammary and salivary glands, activin A acted downstream of HGF signaling through its receptor c-met and was downregulated by HGF during branching (Santos et al., 1994; Woolf et al., 1995; Liu et al., 1996; Sato and Takahashi, 1997; Furue et al., 1999; Maeshima et al., 2000b). In lung and prostate, retinoic acid (RA) inhibited branching (Cardoso et al., 1995; Aboseif et al., 1997; Oshika et al., 1998; Mollard et al., 2000) and activin A was postulated to act downstream of RA signaling because both ActRIIA and ActRIIB were upregulated by RA (Wan et al., 1995; Ameerun et al., 1996). Other factors such as FGFs (Green et al., 1992; Ferguson et al., 1998) and androgens (Risbridger et al., 1996) were also upstream regulators of activin A ligand in at least some organs. Whether these pathways operate during branching morphogenesis and whether these factors

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FIG. 3. Schematic diagram showing the process of branching morphogenesis and a proposed model for the role of activin. This simple model is based on the current evidence that activins are involved in branching morphogenesis. Specifically it proposes that the epithelial bud invades the surrounding mesenchyme, which condenses to form a cap. Branching commences with cleft formation in the mesenchymal cap and bifurcation of the epithelial ducts. We propose that activin expression and bioactivity may be modulated by factors such as hepatocyte growth factor (HGF), fibroblast growth factors (FGF), retinoic acid (RA), and follistatin. This in turn leads to either branching or mesenchymal expansion via ECM remodeling, depending on the degree and location of activin signaling. The ECM remodeling may result from either the direct effects of activins or indirectly via regulation of factors such as Sonic hedgehog.

regulate other activin ligand levels remains to be established. Little is known about the downstream effectors of activin ligands during branching morphogenesis. Sonic hedgehog (Shh) is a secreted signaling factor involved in patterning and in development, and its expression was downregulated by activin receptor-mediated signaling (Kim et al., 2000) and by activin ␤B ligand subunit (Hebrok et al., 1998). Shh was required for development and branching of the prostate and lung (Pepicelli et al., 1998; Podlasek et al., 1999) but inhibited pancreatic development and endocrine differentiation (Hebrok et al., 2000; Kim et al., 2000). Overall, these data implied that Shh acted downstream from activin but the nature of its role in branching morphogenesis is not consistent across the different organs and may be influenced by the predominant activin subunit expressed in each organ. A feature of branching morphogenesis is the extensive remodeling of extracellular matrix (ECM) involving matrix deposition and specific disruption of matrix surrounding budding tips (Hisaoka et al., 1993; Fata et al., 1999; Pohl et al., 2000a). At the bud tips and alongside the basement membrane, epithelial cells contact the mesenchymal ECM. The ECM provides the necessary scaffold along which branching normally occurs and therefore a nonspecific disruption of the matrix inhibited branching (Sorokin et al.,

1990; Falk et al., 1996; Kadoya et al., 1997). In other cells and tissues activin regulated the expression and bioactivity of ECM components (Caniggia et al., 1997; Ogawa et al., 2000), although it is not clear whether ECM remodeling is associated with activin ligand bioactivity during branching morphogenesis. Factors that degrade the basement membrane had effects similar to those described when activin A was exogenously added to salivary gland explants (Banerjee et al., 1977; Nakanishi et al., 1986). Branching morphogenesis also depends on cell–ECM and epithelial cell– cell adhesion. Integrins are expressed by many cell types in branched organs and commonly mediate cell–ECM adhesion in morphogenesis (reviewed in Pohl et al., 2000b). In other systems, activin A played a role in cell adhesion during development (Whittaker and DeSimone, 1993; Ramos et al., 1996), but it is not clear whether activins have similar effects during branching. Further studies using three-dimensional gel matrices could examine the effect of activin on branching in vitro, and particularly on the expression and activity of ECM components and adhesion molecules. Such systems could screen for factors that modify branching by regulating activin ligands and identify effectors of activin action. A molecular description of branching morphogenesis is lacking both overall and in specific organs. To date, this has hampered the identification of regulators and effectors of

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activin action during branching. In the near future, the fields of proteomics and genomics, coupled with techniques such as laser capture microdissection, will have a substantial impact on the characterization of factors involved in branching. For example, detailed profiling of gene expression will identify differentially regulated genes in microdissected epithelium versus mesenchyme in a particular organ. Further, these techniques will allow a comparison of analogous cell compartments across different branched organs. Factors responsible for generating characteristics common to all branching organs, as well as those responsible for giving a specific organ its unique morphology and function, may be identified. Thus, the contribution of activins to these processes can be documented. Detailed studies of the localization and expression of the full range of activin subunits and related proteins are required, in the examination of complex cell– cell interactions. In vitro studies should focus on dissecting the direct and indirect effects of activin signaling in branching morphogenesis by identifying downstream effectors of activin action and pathways with which activin signaling cross-talks. A detailed analysis of genes both regulating and regulated by activins is essential to reveal the role of activins in branching morphogenesis. The studies that showed that activin A stimulates branching in the lung posed an interesting anomaly to our general concept that activin A inhibits branching morphogenesis and raised several questions concerning the interaction and/or redundancy between activins and other ligands. It was not possible to identify a common bioactivity for a particular ligand that was the same for all branched organs; for example, a global inhibition of branching by activin A or global stimulation of branching by TGF␤. Nor was it possible to show that the TGF␤ family members had the same action in a particular branched organ; for example, a global stimulation of branching morphogenesis in the lung given that activin A and BMP-4 stimulated branching but TGF␤1, -␤2, and -␤3 were inhibitory (Kaartinen et al., 1995; Zhou et al., 1996; Weaver et al., 1999; Liu et al., 2000; Shi et al., 2001). Irrespective of the different bioactivities of the ligands in the separate organs, any attempts to simplify the data were further complicated by studying the downstream signaling effectors, for example, Smads. In the lung, exogenously added activin A and TGF␤1 had opposite effects but signal through common Smad proteins (Smad-2, -3, -4). When these Smads were blocked by antisense oligonucleotides, branching was stimulated, implying that the composite effect of endogenous TGF␤ family members was inhibitory (Zhao et al., 1998). Several members of the TGF␤ superfamily, including activin A, were expressed in branched organs. However, the relative levels of expression of the individual ligands and their relative bioactivities are unknown. No comparisons about the relative effects of ligands were reported in any given system. For example, BMP-2, -4, and -7, activin A, and TGF␤ are all expressed in the kidney but it is not known which of these ligands has the most potent bioactivity

(Ritvos et al., 1995; Piscione et al., 1997; Miyazaki et al., 2000; reviewed in Davies and Davey, 1999). A further complication to comparisons such as these is the consideration that branching is a dynamic process that occurs over a period of time. At any point in the process, one member of the TGF␤ family may be more important than another but this may not be related to the developmental stage of the organ as a whole. Rather, the presence of several generations of branches and the spatial organization of the organ might result in a particular ligand being important in maintaining duct length between secondary and tertiary branches, whereas other ligands might be responsible for determining the angle of the branchpoints, point of cleft formation, and so on. For example, in vitro activin A and TGF␤ are inhibitory in a number of organs but activin A inhibited ductal branch formation and elongation, whereas TGF␤ inhibited branching and not elongation or tubulogenesis (Ritvos et al., 1995; Sakurai and Nigam, 1997; Chinoy et al., 1998; Liu et al., 2000). Therefore any analysis of the interaction and redundancy between activins and other ligands requires consideration of the temporal, developmental, and spatial organization of the organ that is undergoing branching and the exact role that each ligand has in that process. Overall, the studies presented in this review provide sufficient evidence to conclude that activins have functional roles in branching morphogenesis. Future studies to determine the roles of any of the TGF␤-superfamily ligands will require consideration of the activins. How the activins perform their roles relative to each other and members of the TGF␤ superfamily represents a significant challenge for investigators in this field.

ACKNOWLEDGMENTS We thank Professor John Bertram, Department of Anatomy and Cell Biology, Monash University; Drs. Ghanim Almahbobi, Richard Mollard, and Jacqueline Schmitt, Monash Institute of Reproduction and Development, Monash University, for critically appraising this manuscript prior to submission and Dr. Belinda Cancilla, University of California San Francisco, for helpful discussions on the topic. The authors are funded by a Program Grant from the National Health and Medical Research Council of Australia (NHMRC).

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