Activins, inhibins and follistatins in the large domestic species

Domestic Animal Endocrinology 28 (2005) 1–16

Review

Activins, inhibins and follistatins in the large domestic species David J. Phillips∗ Center for Molecular Reproduction & Endocrinology, Monash Institute of Reproduction & Development, Monash University, 27-31 Wright Street, Clayton, Vic. 3168, Australia Received 29 April 2004; accepted 31 May 2004

Abstract The activins and inhibins are members of the transforming growth factor-␤ (TGF-␤) superfamily and, along with follistatin, a high affinity binding protein of activin, form a group of interrelated factors originally isolated for their role in regulating the release of follicle-stimulating hormone (FSH). Knowledge of their function, particularly that of activin, has expanded since being originally isolated, such that they are now regarded as important paracrine regulators in many cellular systems. This review summarizes the biology of these proteins as has been established in the large domestic animals. While the majority of data relate to the pituitary, ovary, uterus/placenta and testis, consideration is also given to emerging roles in inflammatory processes and in non-reproductive tissues or systems. © 2004 Elsevier Inc. All rights reserved. Keywords: Follistatin; Transforming growth factor; Follicle-stimulating hormone; Activin; Inhibin

1. Introduction The activins, inhibins and the high affinity binding protein, follistatin, form a group of interrelated factors crucial to a number of reproductive and non-reproductive functions. The first two sets of proteins are now classified as members of the transforming growth factor-␤ (TGF-␤) superfamily, based largely on their dimeric structure and the number of cysteine residues [1]. While this nomenclature is relatively recent, the concept of ‘inhibin’ dates back to the 1930s, when it was postulated that a non-steroidal substance was produced by the gonad to specifically regulate pituitary activity [2]. The ensuing hiatus continued until the mid 1980s when first inhibin, then activin and follistatin were isolated from biological fluids and ∗

Tel.: +61 3 9594 7126; fax: +61 3 9594 7114. E-mail address: [email protected] (D.J. Phillips). 0739-7240/$ – see front matter © 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.domaniend.2004.05.006

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Fig. 1. The various subgroups of the activins, inhibins and follistatins. See text for relevant reviews.

sequenced [3]. At that time and in the ensuing years, the three proteins were largely classified as reproductive hormones produced in the gonad that regulated follicle-stimulating hormone (FSH). Of the three, only inhibin is likely to fulfil correctly the definition as a reproductive hormone, whereas activin and follistatin are known to be relevant to many other tissue systems. The reader is referred to examples of relevant reviews that outline these roles [4,5]. As mentioned, both the activins and inhibins are structurally related and form a distinct subfamily of the TGF-␤ superfamily. Inhibins are made up of so-called ␣–␤ dimers, whereas the activins are made up of ␤–␤ dimers (Fig. 1). There are two forms of inhibins known which are composed of different ␤-subunits, leading to inhibin A (␣–␤A) and inhibin B (␣–␤B). Similarly, combinations of relevant ␤-subunits lead to activin A (␤A–␤A), activin B (␤B–␤B) and the heterodimer activin AB (␤A–␤B). By far and away, the function of activin A has been the most extensively studied. Similarly, the role of the more recently isolated activin ␤C- and ␤E-subunits is ill defined and has not been studied in domestic animal models. Follistatin is structurally unrelated to the activins and inhibins, but binds with high affinity to the ␤-subunits and so is able to neutralize the activity of inhibin, and more particularly, activin forms [6]. Most recently, a separate gene product, variously named follistatin-like 3 (FSTL3) or follistatin-related protein (FSRP), has been isolated which shares some of the properties of follistatin and is structurally similar to some extent [7]. Its biological role, however, is only now being delineated and its relevance to domestic species is still not known. Before specific immunoassays were developed for the various human activin/inhibin dimers, much of the initial biology of these proteins was elucidated in the bovine and ovine, focusing particularly on the potential FSH feedback effects. Once the newer generation of assays became available, emphasis switched to human samples or the use of rodent models. This is not to say that fundamental areas in the biology of these proteins are not known

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for the large domestic species. Therefore, this review will focus on what is known for the various reproductive organs, and then in non-reproductive systems/processes such as the author’s own work in the inflammatory setting. Where appropriate, specific examples in other species will be highlighted, particularly where there are significant differences in the biology of activins, inhibins or follistatins compared with the human situation, or in commonly utilized models such as in rats and mice. First, however, it is pertinent to outline the protein measurement of these factors, on which much of the biology is based. For simplicity, where the term ‘activin’ is mentioned this invariably refers to activin A unless otherwise stated, and similarly reference to activin mRNA relates to mRNA of the activin ␤A-subunit.

2. Assays Initial investigations into the biology of inhibin were carried out with a heterologous radioimmunoassay, the so-called ‘Monash’ assay. This assay measured the ␣-subunit of inhibin, and with subsequent validation it was shown that it cross-reacted with all forms of inhibin dimer, including incompletely processed forms and the free ␣-subunit of inhibin, which is present in the circulation and a number of tissues [8]. It is also pertinent to point out that this assay also measured ‘inhibin’ in the various domestic species in addition to human samples. It was not until the development of ELISA-based assay formats specific for the human forms of inhibin and activin by Groome and colleagues (e.g.[9]), that the ‘Monash’ assay was no longer used routinely. In general, the findings emerging from these newer generation assays confirmed those from the previous assay formats, with a few notable exceptions as outlined below. The current status of immunoassays used for large domestic species is outlined below. It should be pointed out that while a number of other assays have been used to measure activins, inhibins or follistatins in humans, primates or rodents, data using these particular assays have not been published in some of the other species pertinent to this review. The human inhibin A and B ELISAs do not cross-react with sheep, cow or pig, but specific ELISAs for sheep inhibin A and B have been developed based on the Groome antibodies [10,11] and recently an ELISA based on polyclonal antisera for bovine inhibin A has been developed by another group [12]. Given that activin A is almost 100% conserved at the protein level across all species looked at, the human activin A ELISA has been validated for a number of species, including the sheep, cow, rat and mouse (A.E. O’Connor, personal communication). Although a human follistatin ELISA has been developed [13], this does not cross-react particularly with other species. Measurement of follistatin in the sheep, for which by far most information is available, is based on heterologous radioimmunoassays [14,15].

3. Potential feedback regulation and local effects at the pituitary The classical functions of inhibin and follistatin as feedback regulators of FSH release have been conclusively demonstrated in the sheep by injections of recombinant, highly

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purified human preparations [16–19]. While formal proof for activin as an activator of FSH has not been demonstrated in vivo in the domestic species, it would be very unlikely that this was not the case as has been shown in rodent species [20–22] or in the monkey [23,24]. The effects of inhibin on FSH regulation appear to occur entirely at the level of the pituitary, as shown in hypothalamic–pituitary disconnected wethers treated with exogenous inhibin and gonadotropin releasing hormone (GnRH) [16] and considering there was no change in the inhibitory effect when exogenous GnRH inputs were modified [25]. For follistatin, exogenous injections suppress FSH without affecting GnRH basal levels, pulse frequency or amplitude [19]. While the major effect of these proteins is on the regulation of FSH, there may be minor effects of inhibin on LH secretion when GnRH input from the hypothalamus is relatively low [25]. This could occur through an increase in GnRH receptor binding and GnRH-stimulated LH secretion [26,27]. On the other hand, activin dose dependently decreased LH secretion in cultured ovine pituitary cells without affecting LH␤ mRNA [28]. The intrapituitary effects of activin and inhibin have been elucidated in ovine and bovine pituitary cell cultures. Specific inhibin binding sites have been demonstrated in these species [29,30], but it is not clear if these are as yet uncharacterized receptor proteins, betaglycan or the so-called inhibin binding protein/p120 [31]. While the specific pathways, therefore, await elucidation, inhibin has been shown to block FSH production in a proportion of gonadotropes, to lower GnRH receptor number in some FSH-producing gonadotropes and to induce LH GnRH receptors in other gonadotropes [32]. Pituitary cells also have the requisite activin receptors and Smad signaling proteins, and this leads to a rapid upregulation of signaling pathways important for FSH␤ mRNA regulation [28]. Of interest is that unlike many other tissues, the activin of relevance in intrapituitary regulation may well be activin B. This has been demonstrated particularly in the rat [33,34], but is likely to be true at least in the pig [35,36]. The case for the sheep is inferred from a study where an activin B neutralizing antibody decreased FSH secretion in pituitary cultures by around 50% [37], but the expression of the ␤B-subunit versus the ␤A-subunit in the sheep pituitary appears to be relatively low (K.M. Wilson, personal communication).

4. Effects in the ovary The physiology of the activins, inhibins and follistatins in the ovary is one of the few comparatively well-studied organ systems, and so, somewhat ironically, I refer the reader to some recent reviews that cover this area [38–41]. That these proteins are important paracrine regulators of follicle growth and development is unquestioned. How this occurs and at what stages of follicle maturity is the subject of ongoing study. Recent focus has been on the final stages of growth leading to the selection of the dominant follicle [39]. Interestingly, a very recent report using subtractive hybridization of genes from the bovine ovary identified a major upregulation of the activin ␤A-subunit gene near the time when the dominant follicle was being selected [42]. Apart from folliculogenesis, the other major area studied in terms of ovarian paracrine regulation by these proteins is their effects on oocyte maturation. Despite this, some studies showed little or no effect [43–45], while in others, embryo maturation was stimulated at

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least to the morula and blastocyst stage in vitro [46–48]. The difference in the findings of these various studies could be due to the dose and source of activin or inhibin used and/or the type of medium that the oocytes were grown in. Not surprisingly, follistatin has been shown to block activin effects in in vitro-matured oocytes [48], but one cannot rule out the effects on oocyte function that relate to follistatin blocking bone morphogenetic proteins (BMPs). These other members of the TGF-␤ superfamily are potentially bound by follistatin and neutralized (e.g. [49]), and this is particularly relevant given that the BMPs are also known to be important for oocyte development [50]. In terms of specific effects, inhibin and activin may be important in the acquisition of developmental competence during the final stages of oogenesis, with inhibin needed in the early-mid phases and activin in the late phases [47]. Activin does not affect post-fertilization cleavage rate per se, but increases the proportion of cleaved oocytes reaching the blastocyst stage [51]. It also increases the number of cells at the onset of the developmental arrest phase and actually shortens the duration of this quiescent period [52]. A paracrine feedback regulatory network between the oocyte and other follicular cell populations is likely given that the oocyte produces a factor that causes mural granulosa cells to increase their production of activin, inhibin and follistatin, that is, to undergo further development [53]. Furthermore, such a system can most likely be translated to other species, as oocyte maturation involving these factors seems to be operative in the pig [54] and sheep [55].

5. Effects during pregnancy and in the uterus By far and away, most of the research into these proteins in the female reproductive system has focused on the ovary, but that is not to say that this is the only relevant tissue. The inhibin ␣-subunit does not appear to be expressed in the uterus of the sheep [56], but both the ␤A- and ␤B-activin subunits and activin receptors are present in the sheep and pig [56,57], suggestive of a functional paracrine regulatory network involving the activins. Interestingly, follistatin appears to be present in the sheep [56] and horse [58] but not in the pig uterus [57], but this needs to be confirmed in other studies. Feedback effects of the ovary on uterine expression of these proteins appears relevant, as ovariectomy in the ewe led to decreased expression of activin ␤A, follistatin and activin receptor mRNAs, but increased expression of activin ␤B-subunit mRNA [59]. This is particularly interesting, given that ovariectomy does not appear to change circulating levels of activin or follistatin [15,60], due to other organs making the substantial contribution to the circulating pool. During pregnancy, activin, inhibin and follistatin concentrations in the maternal circulation alter. This has generated particular interest, in that in human pregnancies, these proteins appear to rise progressively and notably in the third trimester [61]. This does not appear to be the case for the large domestic animals, where inhibin [10,62,63] and follistatin [14,64] show little change or decrease across gestation in the maternal circulation. The case for circulating activin dynamics during ruminant pregnancy has not been established, but it is known that both the allantoic and amniotic fluid compartments are potentially significant sources [65–67]. Furthermore, a non-human activin binding protein, uterine milk protein, has been demonstrated to be present and possibly be a low affinity, high capacity binding protein in allantoic fluid [68]. The importance of an amniotic fluid source of activin during

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pregnancy is likely given the work of Jenkin and co-workers who have used a sheep model to demonstrate that the amnion is exquisitely sensitive to oxygen status. Generation of hypoxemia by a vascular occluder results in very rapid increases in activin A and follistatin in the maternal circulation [67,69]. As these effects, and their similarly rapid decline following restoration of normoxia, occur in a similar paradigm to prostaglandin E2 , the source of the activin is likely to be the placenta rather than the fetus. Such a concept is consistent with the recently recognized potential of using activin A as a marker of pre-eclampsia in human pregnancies [70]. Given the emerging interest in the physiology of activin, inhibin and follistatin in uterine function and during pregnancy, there is a need to pursue the use of the large domestic species in this regard, and how they differ or how they are compatible with information emerging in other species. For instance, in the sheep and horse, no inhibin ␣-subunit is detectable in the uterus [56,71], whereas in the mouse and human ␣-subunit is expressed [72,73]. This may indicate that placentas of different etiological types have different regulatory systems, particularly in terms of local expression and actions of the activins and inhibins.

6. Effects in the testis Not surprisingly, it was assumed for a number of years that activin, inhibin and follistatin would act as gonadal feedback regulators in both sexes. It is only fairly recently that it has been established beyond doubt that, as predicted, inhibin acts as a feedback regulator of FSH in the male, but it and the other proteins may also be important paracrine and autocrine modulators of testicular function. That the testis is the major source of inhibin in the male was demonstrated by measuring inhibin across the testicular compartments, where, for instance, the concentrations of inhibin in the testicular lymph were high as compared with those in the spermatic vein, which was higher than levels in the testicular or jugular veins [74]. In fact, concentrations of inhibin in the rete testis fluid are approximately 25 times that in testicular lymph and over 500 times that in peripheral plasma [75]. A pivotal finding was using the Groome assays specific for inhibin A and B to show that while inhibin A is detectable in the human male circulation, the predominant form is actually inhibin B [76]. This has led to the concept that inhibin B is the ‘male’ inhibin and such a role has been confirmed in rodents and non-human primates, and of relevance to the current review, is consistent with data in the bull [77], boar [78] and stallion [79]. Despite this, the ram differs from all other species checked so far, in that the ‘male’ inhibin in the sheep appears to be inhibin A [11]. Importantly, the study by McNeilly and co-workers confirmed that in all aspects inhibin A in the ram subserved the functions that inhibin B fulfils in males of other species. Consistent with its role as a feedback regulator of FSH and testicular activity, inhibin ␣-subunit protein and/or mRNA have been localized to Sertoli cells in male sheep, stallions, boars and bulls [11,77,78,80–85]. This is not to say that inhibin A in most species (and inhibin B in the sheep) is not present in the testis; this has been shown in the bovine, for instance [85]. Other testicular cell sources have also been suggested, such as Leydig cells, germ cells and the residual bodies of spermatocytes [11,78,80–83], but these cell types are only likely to be involved in paracrine or autocrine regulation of testicular function.

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The case for activin and follistatin as paracrine regulators in the testis is presumed but has not been conclusively demonstrated in the domestic species, although inhibitory effects of activin on porcine Leydig cell steroidogenesis have been reported [86,87]. In the ovine and bovine, the production of inhibin from the testis reaches a peak that occurs at or before puberty, possibly when Sertoli cell numbers are being established, and then declines to reach adult levels [77,84,88–90]. The pituitary also becomes maximally sensitive to feedback by inhibin by the time of puberty [91], suggesting that the gonadal feedback system involving inhibin develops in the prepubertal animal. There has been a particular focus in the livestock industries on whether immunization against inhibin or the ␣-subunit of inhibin can increase male fertility. While immunization often results in increases in circulation FSH levels (e.g. [92–95]) significant effects on other parameters, such as scrotal circumference, sperm output and sperm reserves and density appear to depend on the form of inhibin which is immunized against, the immunization protocol and the species involved [92–94,96,97]. Thus, it is not clear from published studies whether inhibin immunization will ever be routinely used to increase the fertility of valuable stud males. On the other hand, nature’s own selection for fertility appears to use inhibin as one of its parameters, as males from pig breeds with greater prolificacy have higher FSH and lower inhibin levels [98]. Further, seasonality in rams, and therefore their testicular activity, appears to be linked with the secretion of inhibin across a broad spectrum, from native species such as the mouflon through to various domestic breeds [99]. As discussed above, activin and follistatin could well be paracrine regulators of testicular function (e.g. [86,87]). For inhibin, its role is largely to suppress FSH, but as a testicular feedback factor it is likely to be in reaction with FSH inputs, as changes in FSH levels in rams during seasonal variation occur several weeks prior to changes in inhibin [100]. Some minor effects of inhibin on reducing LH have been shown in rams [93,101,102], but it is not clear if this is representative of all species or peculiar to the sheep.

7. Inflammatory processes While it has been established for a number of years that activin has effects on erythropoiesis and in fact was isolated independently for those properties [103], its role and that of follistatin in inflammatory processes has emerged fairly recently. It was demonstrated in the sheep that circulating follistatin actually increased in the first 24 h following gonadectomy [15,104]. As an aside, this was the formal proof that, unlike inhibin, removal of the ovary or testis did not result in almost undetectable circulating levels of follistatin. Surprisingly, in both studies the sham-operated group had a similar surge of follistatin following surgery, suggestive of surgical trauma. This response could be replicated by exposure of sheep to the bacterial cell wall component, lipopolysaccharide (LPS) or endotoxin, or a pro-inflammatory cytokine, such as interleukin-1␤ [104,105]. While profound, the follistatin response is transitory, as shown in growing lambs given an injection of live yeast, and where the inflammatory response (but not that of follistatin) lasts for some days [106]. How activin might fit into acute inflammatory challenge was ill defined until it was established that because of the high homology between species, the human activin A assay [107] could be adapted for sheep studies. When the LPS-stimulated sheep model was uti-

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Follistatin Interleukin-6

TNF- α

LPS

Activin A Temperature

0

1

2

3

4

5

6

7

8

24

Time from LPS injection (h) Fig. 2. Cytokine and temperature profiles in adult ewes treated with LPS. Each bar represents the period when the particular response is elevated, with lighter shading within the bar indicating a biphasic pattern. Data adapted from [109].

lized, it was found that the release of activin A into the circulation occurred within 1 h of exposure and but was back to basal levels within a few hours [108]. This was in stark contrast to the follistatin response, which occurred later (beginning at around 5 h after LPS) and continued for long. This has the important corollary that the activin release in the face of unchanging follistatin levels is likely to be bioactive and not immediately bound and neutralized, and therefore, having as yet undescribed downstream consequences. More detailed analysis of the activin response by us [109] indicated that it occurs within 50 min of LPS and is actually biphasic, with peaks at around 1.3 and 3.5 h. In relation to other cytokines, the response occurs before that of interleukin-6 and is coincident or slightly earlier than the monophasic peak of tumor necrosis factor-␣ (TNF-␣, Fig. 2). Furthermore, blockade of various components of the inflammatory response, such as fever and the key inflammatory cytokines, interleukin-1 and TNF-␣, had little effect on the activin release. While the cellular source(s) of such a rapid and profound release of activin is yet to be established, a direct interaction with the LPS signaling pathways is required, as the release of activin occurs systemically but not in the central nervous system which is protected by the blood–brain barrier [109]. One potential cellular source responsible for the release of activin during inflammation is the vascular endothelium, where aortic porcine endothelial cells were shown to synthesize and release follistatin in response to LPS treatment [110]. The clinical relevance of these studies in the sheep model have been highlighted by the recent confirmation that septic human patients have elevated serum activin and follistatin levels [111]. Whether such an activin/follistatin response also occurs in response to surgical trauma in humans, like in the sheep, has yet to be established. Often major cardiovascular procedures are compromised by the use of heparin, which causes the release of both activin A and follistatin [112], whereas relatively minor procedures, such as the insertion of a pacemaker or a coronary angiography, do not perturb circulating levels.

8. Other tissues or systems Apart from reproductive tissues and during inflammatory challenge, the large domestic species have not been studied extensively in terms of activin, inhibin or follistatin

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physiology. The adrenal gland is one of the few organs besides the gonad where inhibin is synthesized, and indeed the sheep adrenal is known to produce mRNAs for the ␣-, ␤Aand ␤B-subunits [113]. Adrenocorticotropin is able to stimulate inhibin ␣-subunit mRNA [113], while activin has a minor suppressive effect on adrenocorticotropin actions in the bovine adrenal [114]. The role of inhibin per se in the adrenal is not well understood, but the inhibin ␣-subunit may act as a tumor suppressor gene [115]. In the porcine thyroid, activin is known to affect proliferation of thyroid cells, but there are conflicting reports on whether it is stimulatory [116] or inhibitory [117]. Most of the signaling machinery is present in thyroid cells and activin is localized to thyroid epithelial but not stromal or endothelial cells [117]. Like the adrenal, however, the particular role of these proteins is not well understood. In bone tissues, activin is produced in bovine bone and stimulates erythroid colony formation [118]. It also has a modest effect on proteoglycan and collagen synthesis in bovine chondrocyte cultures, whereas inhibin has a suppressive effect on collagen formation and on cellular proliferation [119]. An important but restricted body of work carried out in bovine, ovine and porcine models relates to vascular endothelial cells. Its applicability to other species and organs is likely to stand up, although endothelial cell populations are by their nature heterogeneous [120]. Activin is known to be inhibitory to endothelial cell proliferation [121] and may form part of an autocrine/paracrine regulatory loop controlling angiogenesis. This is highlighted by elegant studies in bovine endothelial cells where follistatin, but not activin mRNA, is upregulated in subconfluent, migrating cells compared with confluent, quiescent cells [122]. Furthermore, the known angiogenic factor, basic fibroblast growth factor, has a powerful stimulatory effect on follistatin mRNA. It is also known that follistatin has a heparin binding motif and as such one of the two major isoforms of follistatin, follistatin-288, has an affinity for cell surface proteoglycans [6]. This accounts for the observation that unfractionated heparin caused release of follistatin into the circulation of ewes [123], presumably from the vascular endothelium. While activin A does not have a heparin binding motif itself, it is released by heparin, presumably as a complex between activin and follistatin [112,124]. Recent work by us on the sheep has extended these findings to determine that the release of activin and follistatin by heparin is dose-dependent and saturable [124]. The release is specific in terms of being able to be blocked by the heparin neutralizing agent, protamine, which is routinely used in the clinical setting. Surprisingly, protamine treatment alone caused release of follistatin but not activin, raising the possibility that protamine was interacting with unbound follistatin on the surface of the endothelium. Overall, the above studies highlight that heparin and its analogs are unsuitable as anticoagulants in experimental studies due to artifactual release of activin and follistatin into the circulation. Other coagulants such as EDTA have been found to not cause release of these factors and are now routinely used by us and others.

9. Concluding remarks Research into the activins, inhibins and follistatins, and more particularly in the large domestic species, definitely appears to have followed a ‘boom and bust’ series of cycles. A large amount of effort came out of the 1980s, somewhat of a nadir occurred in the 1990s

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and then certain aspects of research into these proteins has picked up again in the most recent three or four years. This is not to impugn the ever growing body of literature that has consistently emerged since this triumvirate of proteins was discovered and has provided some very illuminating findings and which hopefully has been covered herein. Although each tissue/cellular system is likely to be subtly different in terms of its regulatory pathways, in general where activin is known to have a particular property, inhibin, if present, will have an antagonistic function to that of activin. Similarly, follistatin should be regarded as a blocking protein for activin signaling and, therefore, should neutralize any of activin’s actions in the particular organ in question. In some cases, information on particular tissues has not been elucidated for all of the domestic species, but it can likely be inferred from the human situation or in other animal models what is the probable scenario. The high degree of sequence homology between humans and the domestic animal species for the activin ␤A-subunit, activin ␤B-subunit and follistatin (all <97%) also suggest there will be a large degree of structural and functional overlap. Some caution should be ascribed to the above, however where, for instance, it was assumed that in the male sheep inhibin B would be the dominant inhibin, whereas the opposite has recently been shown to be true. Thus, there is a critical need to ascertain the function of the activins, inhibins and follistatin in each of the large domestic animals, even if this is only confirmatory of the general biology pertinent to all species. A further challenge to be met is to determine if there is any relevance of these proteins to commercial livestock production. Additionally, it remains to be seen whether unraveling of their biology in the domestic animals leads to the development of models of human disease that mimic more closely, moreso than in rodent alternatives, appropriate clinical events for that syndrome. Acknowledgements The author’s research is supported in part by a Program Grant (1143786) from the NHMRC of Australia.

References [1] Chang H, Brown CW, Matzuk MM. Genetic analysis of the mammalian transforming growth factor-␤ superfamily. Endocr Rev 2002;23:787–823. [2] McCullagh DR. Dual endocrine activity of the testis. Science 1932;76:19–20. [3] de Kretser DM, Robertson DM. The isolation and physiology of inhibin and related proteins. Biol Reprod 1989;40:33–47. [4] Phillips DJ. New developments in the biology of inhibins, activins and follistatins. Trends Endocrinol Metab 2001;12:94–6. [5] Phillips DJ. The activin/inhibin family. In: Thomson A, Lotze MT, editors. The Cytokine Handbook, 4th ed., vol. 2. Academic Press; 2003, p. 1153–77. [6] Phillips DJ, de Kretser DM. Follistatin: a multifunctional regulatory protein. Front Neuroendocrinol 1998;19:287–322. [7] Schneyer A, Sidis Y, Xia Y, Saito S, del Re E, Lin HY, et al. Differential actions of follistatin and follistatin-like 3. Mol Cell Endocrinol, in press. [8] Robertson DM, Giacometti M, Foulds LM, Lahnstein J, Goss NH, Hearn MTW, et al. Isolation of inhibin ␣-subunit precursor proteins from bovine follicular fluid. Endocrinology 1989;125:2141–9.

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[9] Evans LW, Groome NP. Development of immunoassays for inhibin, activin and follistatin. In: Muttukrisha S, Ledger W, editors. Inhibin, activin and follistatin in human reproductive physiology. Imperial College Press; 2001, p. 11–60. [10] Knight PG, Feist SA, Tannetta DS, Bleach ECL, Fowler PA, O’Brien M, et al. Measurement of inhibin-A (␣␤A dimer) during the oestrous cycle, after manipulation of ovarian activity and during pregnancy in ewes. J Reprod Fertil 1998;113:159–66. [11] McNeilly AS, Souza CJH, Baird DT, Swanston IA, McVerry J, Crawford J, et al. Production of inhibin A not B in rams: changes in plasma inhibin A during testis growth, and expression of inhibin/activin subunit mRNA and protein in adult testis. Reproduction 2002;123:827–35. [12] Kaneko H, Noguchi J, Kikuchi K, Todoroki J, Hasegawa Y. Alterations in peripheral concentrations of inhibin A in cattle studied using a time-resolved immunofluorometric assay: relationship with estradiol and follicle-stimulating hormone in various reproductive conditions. Biol Reprod 2002;67:38–45. [13] Evans LW, Muttukrishna S, Groome NP. Development, validation and application of an ultra-sensitive two-site enzyme immunoassay for human follistatin. J Endocrinol 1998;156:275–82. [14] Xia Y, O’Shea T, Murison R, McFarlane JR. Concentrations of progesterone, follistatin, and folliclestimulating hormone in peripheral plasma across the estrous cycle and pregnancy in Merino ewes that are homozygous or noncarriers of the Booroola gene. Biol Reprod 2003;69:1079–84. [15] Klein R, Findlay JK, Clarke IJ, de Kretser DM, Robertson DM. Radioimmunoassay of FSH-suppressing protein in the ewe: concentrations during the oestrous cycle and following ovariectomy. J Endocrinol 1993;137:433–43. [16] Tilbrook AJ, de Kretser DM, Clarke IJ. Human recombinant inhibin A and testosterone act directly at the pituitary to suppress plasma concentrations of FSH in castrated rams. J Endocrinol 1993;138:181–9. [17] Tilbrook AJ, de Kretser DM, Clarke IJ. Human recombinant inhibin A suppresses plasma follicle-stimulating hormone to intact levels but has no effect on luteinizing hormone in castrated rams. Biol Reprod 1993;49:779–88. [18] Tilbrook AJ, Clarke IJ, de Kretser DM. Human recombinant follistatin-288 suppresses plasma concentrations of follicle-stimulating hormone but is not a significant regulator of luteinizing hormone in castrated rams. Biol Reprod 1995;53:1353–8. [19] Padmanabhan V, Battaglia D, Brown MB, Karsch FJ, Lee JS, Pang WQ, et al. Neuroendocrine control of follicle-stimulating hormone (FSH) secretion: II. Is follistatin-induced suppression of FSH secretion mediated via changes in activin availability and does it involve changes in gonadotropin-releasing hormone secretion? Biol Reprod 2002;66:1395–402. [20] Schwall R, Schmelzer CH, Matsuyama E, Mason AJ. Multiple actions of recombinant activin-A in vivo. Endocrinology 1989;125:1420–3. [21] Rivier C, Vale W. Effect of recombinant activin-A on gonadotropin secretion in the female rat. Endocrinology 1991;129:2463–5. [22] Doi M, Igarashi M, Hasegawa Y, Eto Y, Shibai H, Miura T, et al. In vivo action of activin-A on pituitary–gonadal system. Endocrinology 1992;130:139–44. [23] McLachlan RI, Dahl KD, Bremner WJ, Schwall R, Schmelzer CH, Mason AJ, et al. Recombinant human activin-A stimulates basal FSH and GnRH-stimulated FSH and LH release in the adult male macaque, Macaca fascicularis. Endocrinology 1989;125:2787–9. [24] Stouffer RL, Woodruff TK, Dahl KD, Hess DL, Mather JP, Molskness TA. Human recombinant activin-A alters pituitary luteinizing hormone and follicle-stimulating hormone secretion, follicular development, and steroidogenesis during the menstrual cycle in rhesus monkeys. J Clin Endocrinol Metab 1993;77:241–8. [25] Tilbrook AJ, Clarke IJ. Negative feedback regulation of the secretion and actions of gonadotropin-releasing hormone in males. Biol Reprod 2001;64:735–42. [26] Laws SC, Beggs MJ, Webster JC, Miller WL. Inhibin increases and progesterone decreases receptors for gonadotropin-releasing hormone in ovine pituitary culture. Endocrinology 1990;127:373–80. [27] Muttukrishna S, Knight PG. Inverse effects of activin and inhibin on the synthesis and secretion of FSH and LH by ovine pituitary cells in vitro. J Mol Endocrinol 1991;6:171–8. [28] Dupont J, McNeilly J, Vaiman A, Canepa S, Combarnous Y, Taragnat C. Activin signaling pathways in ovine pituitary and L␤T2 gonadotrope cells. Biol Reprod 2003;68:1877–87. [29] Castillo RJ, Olivieri E, Vega I. Inhibin binding sites in bovine pituitary membranes. Biol Res 1996;29:183–8.

12

D.J. Phillips / Domestic Animal Endocrinology 28 (2005) 1–16

[30] Hertan R, Farnworth PG, Fitzsimmons KL, Robertson DM. Identification of high affinity binding sites for inhibin on ovine pituitary cells in culture. Endocrinology 1999;140:6–12. [31] Phillips DJ, Woodruff TK. Inhibin: actions and signalling. Growth Factors 2004;22:13–8. [32] Ghosh BR, Wu JC, Strahl BD, Childs GV, Miller WL. Inhibin and estradiol alter gonadotropes differentially in ovine pituitary cultures: changing gonadotrope numbers and calcium responses to gonadotropin-releasing hormone. Endocrinology 1996;137:5144–54. [33] Corrigan AZ, Bilezikjian LM, Carroll RS, Bald LN, Schmelzer CH, Fendly BM, et al. Evidence for an autocrine role of activin-B within rat anterior pituitary cultures. Endocrinology 1991;128:1682–4. [34] Bilezikjian LM, Vaughan JM, Vale WW. Characterization and the regulation of inhibin/activin subunit proteins of cultured rat anterior pituitary cells. Endocrinology 1993;133:2545–53. [35] Li MD, MacDonald GJ, Ford JJ. Breed differences in expression of inhibin/activin subunits in porcine anterior pituitary glands. Endocrinology 1997;138:712–8. [36] Schneider O, Nau R, Michel U. Comparative analysis of follistatin-, activin beta A- and activin beta B-mRNA steady-state levels in diverse porcine tissues by multiplex S1 nuclease analysis. Eur J Endocrinol 2000;142:537–44. [37] Baratta M, West LA, Turzillo AM, Nett TM. Activin modulates differential effects of estradiol on synthesis and secretion of follicle-stimulating hormone in ovine pituitary cells. Biol Reprod 2001;64:714–9. [38] Knight PG, Glister C. Local roles of TGF-␤ superfamily members in the control of ovarian follicle development. Anim Reprod Sci 2003;78:165–83. [39] Mihm M, Austin EJ. The final stages of dominant follicle selection in cattle. Domest Anim Endocrinol 2002;23:155–66. [40] McNatty KP, Fidler AE, Juengel JL, Quirke LD, Smith PR, Heath DA, et al. Growth and paracrine factors regulating follicular formation and cellular function. Mol Cell Endocrinol 2000;163:11–20. [41] Braw-Tal R. The initiation of follicle growth: the oocyte or the somatic cells? Mol Cell Endocrinol 2002;187:11–8. [42] Sisco B, Hagemann LJ, Shelling AN, Pfeffer PL. Isolation of genes differentially expressed in dominant and subordinate bovine follicles. Endocrinology 2003;144:3904–13. [43] Van Tol HTA, de Loos FAM, Vanderstichele HMJ, Bevers MM. Bovine activin A does not affect the in vitro maturation of bovine oocytes. Theriogenology 1994;41:673–9. [44] Smith LC, Olivera-Angel M, Groome NP, Bhatia B, Price CA. Oocyte quality in small antral follicles in the presence or absence of a large dominant follicle in cattle. J Reprod Fertil 1996;106:193–9. [45] Izadyar F, Zeinstra E, Colenbrander B, Vanderstichele HM, Bevers MM. In vitro maturation of bovine oocytes in the presence of bovine activin A does not affect the number of embryos. Anim Reprod Sci 1996;45:37–45. [46] Yoshioka K, Kamomae H. Recombinant human activin A stimulates development of bovine one-cell embryos matured and fertilized in vitro. Mol Reprod Dev 1996;45:151–6. [47] Stock AE, Woodruff TK, Smith LC. Effects of inhibin A and activin A during in vitro maturation of bovine oocytes in hormone- and serum-free medium. Biol Reprod 1997;56:1559–64. [48] Yoshioka K, Suzuki C, Iwamura S. Activin A and follistatin regulate developmental competence of in vitro-produced bovine embryos. Biol Reprod 1998;59:1017–22. [49] Otsuka F, Moore RK, Iemuro S-I, Ueno N, Shimasaki S. Follistatin inhibits the function of the oocyte-derived factor BMP-15. Biochem Biophys Res Commun 2001;289:961–6. [50] Shimasaki S, Moore RK, Erickson GF, Otsuka F. The role of bone morphogenetic proteins in ovarian function. Reproduction 2003;Suppl. 60:323–37. [51] Silva CC, Knight PG. Modulatory actions of activin-A and follistatin on the developmental competence of in vitro-matured bovine oocytes. Biol Reprod 1998;58:558–65. [52] Yoshioka K, Suzuki C, Iwamura S. Effects of activin A and follistatin on developmental kinetics of bovine embryos: cinematographic analysis in a chemically defined medium. J Reprod Fertil 2000;118:119–25. [53] Glister C, Groome NP, Knight PG. Oocyte-mediated suppression of follicle-stimulating hormone- and insulin-like growth factor-induced secretion of steroids and inhibin-related proteins by bovine granulosa cells in vitro: possible role of transforming growth factor ␣. Biol Reprod 2003;68:758–65. [54] Kaneko H, Kikuchi K, Noguchi J, Hosoe M, Akita T. Maturation and fertilization of porcine oocytes from primordial follicles by a combination of xenografting and in vitro culture. Biol Reprod 2003;69:1488–93.

D.J. Phillips / Domestic Animal Endocrinology 28 (2005) 1–16

13

[55] Thomas FH, Armstrong DG, Telfer EE. Activin promotes oocyte development in ovine preantral follicles in vitro. Reprod Biol Endocrinol 2003;1:76–82. [56] Hayashi K, Carpenter KD, Gray CA, Spencer TE. The activin–follistatin system in the neonatal ovine uterus. Biol Reprod 2003;69:843–50. [57] van de Pavert SA, Boerjan ML, Stroband HWJ, Taverne MAM, van den Hurk R. Uterine–embryonic interaction in pig: activin, follistatin, and activin receptor II in uterus and embryo during early gestation. Mol Reprod Dev 2001;59:390–9. [58] Sugawara Y, Yamanouchi K, Naito K, Tachi C, Tojo H, Sawasaki T. Molecular cloning of cDNA for equine follistatin and its gene expression in the reproductive tissues of the mare. J Vet Med Sci 1999;61:201–7. [59] Carpenter KD, Hayashi K, Spencer TE. Ovarian regulation of endometrial gland morphogenesis and activin–follistatin system in the neonatal ovine uterus. Biol Reprod 2003;69:851–60. [60] Sakai R, Shiozaki M, Tabuchi M, Eto Y. The measurement of activin/EDF in mouse serum: evidence for extragonadal production. Biochem Biophys Res Commun 1992;188:921–6. [61] Muttukrishna S. Inhibin, activin and follistatin in human pregnancy. In: Muttukrishna S, Ledger W, editors. Inhibin, activin and follistatin in human reproductive physiology. Imperial College Press; 2001, p. 119–40. [62] Findlay JK, Doughton BD, Russell DL. Peripheral concentrations of immunoreactive inhibin during pregnancy and parturition in the ewe. Reprod Fertil Dev 1991;3:543–9. [63] Miller S, Wongprasartsuk S, Young IR, Wlodek ME, McFarlane JR, de Kretser DM, et al. Source of inhibin in ovine fetal plasma and amniotic fluid during late gestation: half-life of fetal inhibin. Biol Reprod 1997;57:347–53. [64] McFarlane JR, Xia Y, O’Shea T, Hayward S, O’Connor AE, de Kretser DM. Follistatin concentrations in maternal and fetal fluids during the oestrous cycle, gestation and parturition in Merino sheep. Reproduction 2002;124:259–65. [65] de Kretser DM, Foulds LM, Hancock M, McFarlane J, Goss N, Jenkin G. The isolation of activin from ovine amniotic fluid. Endocrinology 1994;134:1231–7. [66] Foulds LM, de Kretser DM, Farnworth P, Buttress D, Jenkin G, Groome NP, et al. Ovine allantoic fluid contains high concentrations of activin A: partial dissociation of immunoactivity and bioactivity. Biol Reprod 1998;59:233–40. [67] Jenkin G, Ward J, Loose J, Schneider-Kolsky M, Young R, Canny B, et al. Physiological and regulatory roles of activin A in late pregnancy. Mol Cell Endocrinol 2001;180:131–8. [68] McFarlane JR, Foulds LM, O’Connor AE, Phillips DJ, Jenkin G, Hearn MTW, et al. Uterine milk protein, a novel activin-binding protein, is present in ovine allantoic fluid. Endocrinology 1999;140:4745–52. [69] Jenkin G, Ward J, Hooper SB, O’Connor AE, de Kretser DM, Wallace EM. Feto-placental hypoxemia regulates the release of fetal activin A and prostaglandin E2. Endocrinology 2001;142:963–6. [70] Bersinger NA, Smárason AK, Muttukrishna S, Groome NP, Redman CW. Women with preeclampsia have increased serum levels of pregnancy-assiucatd plasma protein A (PAPP-A), inhibin A, activin A and soluble E-selectin. Hypertens Pregnancy 2003;22:65–75. [71] Yamanouchi K, Hirasawa K, Hasegawa T, Ikeda A, Chang K-T, Matsuyama S, et al. Equine inhibin/activin ␤A-subunit mRNA is expressed in the endometrial gland, but not in the trophoblast, during pregnancy. Mol Reprod Dev 1997;47:363–9. [72] Yamaguchi M, Fujii M, Miyake A. Cyclic AMP regulates expression of activin and inhibin subunit mRNAs in the mouse placenta and decidua: a short communication. Eur J Endocrinol 1996;135:379–81. [73] Jones RL, Salamonsen LA, Critchley HOD, Rogers PAW, Affandi B, Findlay JK. Inhibin and activin subunits are differentially expressed in endometrial cells and leukocytes during the menstrual cycle, in early pregnancy and in women using progestin-only contraception. Mol Hum Reprod 2000;6:1107–17. [74] Tilbrook AJ, de Kretser DM, Clarke IJ. Studies on the testicular source of inhibin and its route of secretion in rams: failure of the Leydig cell to secrete inhibin in response to a human chorionic gonadotrophin/LH stimulus. J Endocrinol 1991;130:107–14. [75] Voglmayr JK, Jolley D, Vale W, Willoughby D, Moser A, So CK, et al. Effects of follicle-stimulating hormone on inhibin release by different testicular compartments in the adult ram. Biol Reprod 1992;47:573–81. [76] Illingworth PJ, Groome NP, Byrd W, Rainey WE, McNeilly AS, Mather JP, et al. Inhibin-B: a likely candidate for the physiologically important form of inhibin in men. J Clin Endocrinol Metab 1996;81:1321–5. [77] Kaneko H, Noguchi J, Kikuchi K, Akagi S, Shimada A, Taya K, et al. Production and endocrine role of inhibin during the early development of bull calves. Biol Reprod 2001;65:209–15.

14

D.J. Phillips / Domestic Animal Endocrinology 28 (2005) 1–16

[78] Jin W, Arai KY, Herath CB, Kondo M, Ishi H, Tanioka Y, et al. Inhibins in the male Göttingen miniature pig: Leydig cells are the predominant source of inhibin B. J Androl 2001;22:953–60. [79] Tanaka Y, Taniyama H, Tsunoda N, Shinbo H, Nagamine N, Nambo Y, et al. The testis as a major source of circulating inhibins in the male equine fetus during the second half of gestation. J Androl 2002;23:229–36. [80] Nagata S, Miyake YI, Nambo Y, Nagamine N, Watanabe G, Tsunoda N, et al. Inhibin secretion in the stallion. Equine Vet J 1998;30:98–103. [81] Nagata S, Tsunoda N, Nagamine N, Tanaka Y, Taniyama H, Nambo Y, et al. Testicular inhibin in the stallion: cellular source and seasonal changes in its secretion. Biol Reprod 1998;59:62–8. [82] Fujimura S, Hondo E, Kobayashi T, Yamanouchi K, Inoue N, Nagata S, et al. Expression of inhibin ␣-subunit in horse testis. J Vet Med Sci 1998;60:937–42. [83] Jarred RA, Cancilla B, Richards M, Groome NP, McNatty KP, Risbridger GP. Differential localization of inhibin subunit proteins in the ovine testis during fetal gonadal development. Endocrinology 1999;140:979– 86. [84] Matsuzaki S, Uenoyama Y, Okuda K, Watanabe G, Kitamura N, Taya K, et al. Prepubertal changes in immunoreactive inhibin concentration in blood serum and testicular tissue in Holstein bull calves. J Vet Med Sci 2001;63:1303–7. [85] Kaneko H, Noguchi J, Kikuchi K, Hasegawa Y. Molecular weight forms of inhibin A and inhibin B in the bovine testis change with age. Biol Reprod 2003;68:1918–25. [86] Mauduit C, Chauvin MA, de Peretti E, Morera AM, Benahmed M. Effect of activin A on dehydroepiandrosterone and testosterone secretion by primary immature porcine Leydig cells. Biol Reprod 1991;45:101–9. [87] Lejeune H, Chuzel F, Sanchez P, Durand P, Mather JP, Saez JM. Stimulating effect of both human recombinant inhibin A and activin A on immature porcine Leydig cell functions in vitro. Endocrinology 1997;138:4783–91. [88] MacDonald RD, Deaver DR, Schanbacher BD. Prepubertal changes in plasma FSH and inhibin in Holstein bull calves: responses to castration and (or) estradiol. J Anim Sci 1991;69:276–82. [89] Isaacs KL, McNatty KP, Condell L, Shaw L, Heath DA, Hudson NL, et al. Plasma FSH, LH and immunoreactive inhibin concentrations in FecBB /FecBB and FecB+ /FecB+ Booroola ewes and rams from birth to 12 months of age. J Reprod Fertil 1995;103:89–97. [90] Sanford LM, Moore C, Voglmayr JK, Fahmy MH. Sexual maturational changes circulatory inhibin concentration in relation to FSH concentration and testicular size in Suffolk and DLS rams. Theriogenology 2000;54:719–30. [91] Tilbrook AJ, de Kretser DM, Clarke IJ. Changes in the suppressive effects of recombinant inhibin A on FSH secretion in ram lambs during sexual maturation: evidence for alterations in the clearance rate of inhibin. J Endocrinol 1999;161:219–29. [92] Martin TL, Williams GL, Lunstra DD, Ireland JJ. Immunoneutralization of inhibin modifies hormone secretion and sperm production in bulls. Biol Reprod 1991;45:73–7. [93] McKeown RM, O’Callaghan D, Roche JF, Boland MP. Effect of immunization of rams against bovine inhibin alpha 1-26 on semen characteristics, scrotal size, FSH, LH and testosterone concentrations. J Reprod Fertil 1997;109:237–45. [94] Bame JH, Dalton JC, Delegos SD, Good TE, Ireland JL, Jimenez-Krassel F, et al. Effect of long-term immunization against inhibin on sperm output in bulls. Biol Reprod 1999;60:1360–6. [95] Araki K, Arai KY, Watanabe G, Taya K. Involvement of inhibin in the regulation of follicle-stimulating hormone secretion in the young adult male Shiba goat. J Androl 2000;21:558–65. [96] Voglmayr JK, Mizumachi M, Washington DW, Chen C-LC, Bardin CW. Immunization of rams against human recombinant inhibin ␣-subunit delays, augments, and extends season-related increase in blood gonadotropin levels. Biol Reprod 1990;42:81–6. [97] Schanbacher BD. Pituitary and testicular responses of beef bulls to active immunization against inhibin alpha. J Anim Sci 1991;69:252–7. [98] Borg KE, Lunstra DD, Christenson RK. Semen characteristics, testicular size, and reproductive hormone concentrations in mature Duroc, Meishan, Fengjing, and Minzhu boars. Biol Reprod 1993;49:515–21. [99] Lincoln GA, Lincoln CE, McNeilly AS. Seasonal cycles in the blood plasma concentration of FSH, inhibin and testosterone, and testicular size in rams of wild, feral and domesticated breeds of sheep. J Reprod Fertil 1990;88:623–33.

D.J. Phillips / Domestic Animal Endocrinology 28 (2005) 1–16

15

[100] Sanford LM, Price CA, Leggee DG, Baker SJ, Yarney TA. Role of FSH, numbers of FSH receptors and testosterone in the regulation of inhibin secretion during the seasonal testicular cycle of adult rams. Reproduction 2002;123:269–80. [101] Tilbrook AJ, de Kretser DM, Clarke IJ. Influence of the degree of stimulation of the pituitary by gonadotropin-releasing hormone on the action of inhibin and testosterone to suppress the secretion of the gonadotropins in rams. Biol Reprod 2001;64:473–81. [102] Wheaton JE, Godfrey RW. Plasma LH, FSH, testosterone, and age at puberty in ram lambs actively immunized against an inhibin ␣-subunit peptide. Theriogenology 2003;60:933–41. [103] Eto Y, Tsuji T, Takezawa M, Takano S, Yokogawa Y, Shibai H. Purification and characterization of erythroid differentiation factor (EDF) isolated from human leukemia cell line THP-1. Biochem Biophys Res Commun 1987;142:1095–103. [104] Phillips DJ, Hedger MP, McFarlane JR, Klein R, Clarke IJ, Tilbrook AJ, et al. Follistatin concentrations in male sheep increase following sham castration/castration or injection of interleukin-1␤. J Endocrinol 1996;151:119–24. [105] Klein R, Clarke IJ, Hedger MP, Robertson DM. Plasma follistatin concentrations increase following lipopolysaccharide administration in sheep. Clin Exp Pharmacol Physiol 1996;23:754–5. [106] Phillips DJ, de Kretser DM, Pfeffer A, Ng Chie W, Moore LG. Follistatin has a biphasic response but follicle-stimulating hormone is unchanged during an inflammatory episode in growing lambs. J Endocrinol 1998;156:77–82. [107] Knight PG, Muttukrishna S, Groome NP. Development and application of a two-site enzyme immunoassay for the determination of ‘total’ activin-A concentrations in serum and follicular fluid. J Endocrinol 1996;148:267–79. [108] Jones KL, Brauman JN, Groome NP, de Kretser DM, Phillips DJ. Activin A release into the circulation is an early event in systemic inflammation and precedes the release of follistatin. Endocrinology 2000;141:1905– 8. [109] Jones KL, de Kretser DM, Clarke IJ, Scheerlinck J-PY, Phillips DJ. Characterisation of the rapid release of activin A following acute lipopolysaccharide challenge in the ewe. J Endocrinol 2004;182:69–80. [110] Michel U, Schneider O, Kirchhof C, Meisel S, Smirnov A, Wiltfang J, et al. Production of follistatin in porcine endothelial cells: differential regulation by bacterial compounds and the synthetic glucocorticoid RU 28362. Endocrinology 1996;137:4925–34. [111] Michel U, Ebert S, Phillips D, Nau R. Serum concentrations of activin and follistatin are elevated and run in parallel in patients with septicemia. Eur J Endocrinol 2003;148:559–64. [112] Phillips DJ, Jones KL, McGaw DJ, Groome NP, Smolich JJ, Pärsson H, et al. Release of activin and follistatin during cardiovascular procedures is largely due to heparin administration. J Clin Endocrinol Metab 2000;85:2411–5. [113] Crawford RJ, Hammond VE, Evans BA, Coghlan JP, Haralambidis J, Hudson B, et al. ␣-inhibin gene expression occurs in the ovine adrenal cortex, and is regulated by adrenocorticotropin. Mol Endocrinol 1987;1:699–706. [114] Nishi Y, Haji M, Tanaka S, Yanase T, Takayanagi R, Etoh Y, et al. Human recombinant activin-A modulates the steroidogenesis of cultured bovine adrenocortical cells. J Endocrinol 1992;132:R1–4. [115] Matzuk MM, Finegold MJ, Mather JP, Krummen L, Lu H, Bradley A. Development of cancer cachexia-like syndrome and adrenal tumors in inhibin-deficient mice. Proc Natl Acad Sci USA 1994;91:8817–21. [116] Kotajima A, Miyamoto Y, Tsuruo M, Kosaka M, Saito S. Effects of activin A on deoxyribonucleic acid synthesis, iodine metabolism, and cyclic adenosine monophosphate accumulation porcine thyroid cells. Endocrinology 1995;136:1214–8. [117] Franzén A, Piek E, Westermark B, ten Dijke P, Heldin N-E. Expression of transforming growth factor-␤1, activin A, and their receptors in thyroid follicle cells: negative regulation of thyrocyte growth and function. Endocrinology 1999;140:4300–10. [118] Ogawa Y, Schmidt DK, Nathan RM, Armstrong RM, Miller KL, Sawamura SJ, et al. Bovine bone activin enhances bone morphogenetic protein-induced ectopic bone formation. J Biol Chem 1992;267:14233–7. [119] Luyten FP, Chen P, Paralkar V, Reddi AH. Recombinant bone morphogenetic protein-4, transforming growth factor-␤1, and activin A enhance the cartilage phenotype of articular chondrocytes in vitro. Exp Cell Res 1994;210:224–9.

16

D.J. Phillips / Domestic Animal Endocrinology 28 (2005) 1–16

[120] Ribatti D, Nico B, Vacca A, Roncali L, Dammacco F. Endothelial cell heterogeneity and organ specificity. J Hematother Stem Cell Res 2002;11:81–90. [121] McCarthy SA, Bicknell R. Inhibition of vascular endothelial cell growth by activin-A. J Biol Chem 1993;268:23066–71. [122] Kozian DH, Ziche M, Augustin HG. The activin-binding protein follistatin regulates autocrine endothelial cell activity and induces angiogenesis. Lab Invest 1997;76:267–76. [123] Klein R, Robertson DM, Clarke IJ. Studies in sheep examining plasma follistatin elevations due to frequent blood sampling or surgery. Reprod Fertil Dev 1996;8:273–7. [124] Jones KL, de Kretser DM, Phillips DJ. Effect of heparin administration to sheep on the release profiles of circulating activin A and follistatin. J Endocrinol 2004;181:307–14.