Deer Antlers as a Model of Mammalian Regeneration

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Deer Antlers as a Model of Mammalian Regeneration Joanna Price,* Corrine Faucheux,{ and Steve Allen* *Department of Veterinary Basic Sciences, The Royal Veterinary College London NW1 OTU, United Kingdom { INSERM U-44 1, Pessac, France

I. Introduction A. Development of the First Set of Antlers B. Antler Regeneration C. The Early Stages D. Ontogony E. Longitudinal Growth F. Chondrogenesis G. Regenerating Antlers: Ossification and Remodeling II. Regulation of the Antler Development and Regeneration A. External and Systemic Factors B. Local Mechanisms Acknowledgments References

Deer antlers are cranial appendages that develop after birth as extensions of a permanent protuberance (pedicle) on the frontal bone. Pedicles and antlers originate from a specialized region of the frontal bone; the ‘antlerogeneic periosteum’ and the systemic cue which triggers their development in the fawn is an increase in circulating androgen. These primary antlers are then shed and regenerated the following year in a larger, more complex form. Antler growth is extremely rapid—an adult red deer can produce a pair of antlers weighing
30kg in three months, and involves both endochondral and intramembranous ossification. Since antlers are sexual secondary characteristics, their annual cycles of growth have evolved to be closely coordinated to the reproductive cycle which, in temperate species, is linked to the photoperiod. Cessation of antler growth and death of the overlying skin (velvet) coincides with a rise in circulating testosterone as the autumn breeding season approaches. The ‘dead’ antlers remain attached to the pedicle until they are shed (cast) the following spring when circulating testosterone levels fall. In red deer, the species that we study, casting of the old set of antlers is followed immediately by growth of the new set. Current Topics in Developmental Biology, Vol. 67 Copyright 2005, Elsevier Inc. All rights reserved.

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0070-2153/05 $35.00 DOI: 10.1016/S0070-2153(04)67001-3

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Although the anatomy of antler growth and the endocrine changes associated with it have been well documented, the molecular mechanisms involved remain poorly understood. The case for continuing to decipher them remains compelling, despite the obvious limitations of using deer as an experimental model, because this research will help provide insight into why humans and other mammals have lost the ability to regenerate organs. From the information so far available, it would appear that the signaling pathways that control the development of skeletal elements are recapitulated in regenerating antlers. This apparent lack of any specific ‘antlerogenic molecular machinery’ suggests that the secret of deers’ ability to regenerate antlers lies in the particular cues to which multipotential progenitor/ stem cells in an antler’s ‘regeneration territory’ are exposed. This in turn suggests that with appropriate manipulation of the environment, pluripotential cells in other adult mammalian tissues could be stimulated to increase the healing capacity of organs, even if not to regenerate them completely. The need for replacement organs in humans is substantial. The benefits of increasing individuals’ own capacity for regeneration and repair are self evident. C 2005, Elsevier Inc.

I. Introduction Since ancient times deer antlers have held a fascination for humans as beautiful and spectacular works of nature and as symbols of male superiority and strength. Human alpha males hang antlers on walls as a demonstration of their own wealth and power and because they represent male strength and virility. Extracts of deer antler have also been used for centuries as components of oriental medicine. In many parts of the world, deer are now farmed for the production of antler velvet and this industry has undoubtedly helped rejuvenate antler research in recent years. Antlers have also long been a focus of scientific interest because, while the study of antlers is relevant to many areas of biology—bone biology, developmental biology, zoology, evolutionary biology, and endocrinology—it is their ability to regenerate that makes antlers so important. Mammals have a very limited regenerative ability, whereas most other phyla which include some species which can regenerate large sections of their body plan after injury or amputation. The study of antlers can help shed light on why this may be the case. However, the limitation of antlers for investigating the molecular processes of regeneration must not be overlooked. Few deer gene sequences are known, there is no ‘‘deer genome project’’ on the horizon, and transgenic deer are unlikely to exist outside the realm of science fiction. Although genomic and proteomic approaches are currently being used to identify molecules expressed in antlers, establishing function will always be a

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challenge. As well as being genetically intractable, deer are large wild animals that require specialized management. Notwithstanding, despite not being a mainstream laboratory animal, deer are the only mammals that can regenerate complete appendages and so deserve to retain a place in regeneration research. Regenerative medicine is an expanding field, and, as discussed by Brockes and Martin (2004) at a recent Royal Society discussion meeting on tissue repair and regeneration, the continued study of a variety of natural examples of regeneration can only increase the prospects for the restoration of functional tissues and organs in humans. A. Development of the First Set of Antlers Except in the genus Rangifer (reindeer), antlers develop only in male deer and, in most species, this occurs in the spring of the animal’s second year of life. Thus, while parallels are frequently drawn between antlers and other developing appendages such as limbs, antlers are unique since they develop after birth. Antlers therefore provide a unique model for studying the mechanisms that control the development of a complete bony appendage from tissues that have presumably completed a developmental program. Antlers grow from pedicles, secondary sexual characteristics that are outgrowths of the frontal bone (Fig. 1), and it is from the pedicles that antlers are shed and regenerate each year. However, the presence of a pedicle bone is not an absolute requirement for antler formation because pedicle amputation in a number of species has been shown not to prevent subsequent antler

Figure 1 Two-year-old red deer stag with a set of regenerated antlers at 75 days of growth. The pedicle, the permanent extension of the frontal bone, is marked with the arrow. At this stage of the growth cycle, antlers continue to elongate at the distal tip, but growth in more proximal branches (tines) has stopped.

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development, although the pedicles themselves did not always regenerate (Bubenik and Pavalansky, 1965; Goss, 1961; Jaczewski, 1954, 1955). Li et al. (2001b) have described a potentially interesting relationship between pedicle height and antler phylogeny; more highly evolved deer species, which have larger and more complex antlers, have shorter pedicles, although the functional significance of this is not known. One of the first important issues that antler biologists sought to address was the identity of the tissue(s) that are responsible for initiating pedicle and antler development. A series of transplantation experiments, many carried out several decades ago, demonstrated that periosteum is the tissue involved. Excision of skin and subcutaneous tissues from the frontal bone in young fawns was found to have no eVect on antler development whereas resection of periosteum and surrounding bone prevented development (Goss and Powel, 1985; Goss et al., 1964; Hartwig, 1967; Hartwig and Schrudde, 1974). Transplantation of periosteum to another site on the frontal bone led to antler development at this new location, but not always at the original site (Hartwig, 1968). Hartwig and Schrudde (1974) also showed that transplantation of periosteum to the leg resulted in the development of small antlers which display an annual cycle of growth. Kierdorf and Kierdorf (2000) repeated this experiment and found no ectopic growth until nine years after transplantation and they concluded that the pedicle had to attain a minimum size before an antler could form. This tissue was first described as ‘‘antlerogenic periosteum’’ (AP) (Goss, 1983) and it has been extensively studied and reviewed by Kierdorf and Kierdorf (2001) and by Li and Suttie (2001). Li et al. (2001a) showed that structures resembling antlers, or pedicle-antlers, could be generated if AP is transplanted over the calvarial bones of a nude mouse. As will be discussed in more detail in a later section, an adult derivative of this antlerogeneic periosteum is likely to be the source of progenitor cells from which some, if not all, regenerating antler tissues are derived. Even during fetal life, the sites where future antlers are destined to grow are apparent as small bony elevations on the lateral crests of the frontal bone of the skull (Lincoln, 1973). These anlage of pedicle enlarge between 55 and 100 days of gestation but regress in later stages (Li and Suttie, 2001). However, after birth, the periosteum at this site remains thicker than in other locations on the skull. Initially, the bone beneath this periosteum is made up of flattened plates, characteristic of cranial bones. However, as androgen levels increase at the time of puberty, new trabeculae form beneath the periosteum and a visible pedicle forms (Sempere and Boisson, 1983; Suttie et al., 1984, 1991). Histological studies in red deer by Li and Suttie (1994) have shown that pedicle formation is made up of four ossification stages: (1) intramembranous ossification (direct formation of bone by osteoblasts insignaling cellular periosteum), (2) transitional ossification (formation of osseocartilaginous tissue), (3) pedicle endochondral ossification (when only

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chondrogenesis takes place in the pedicle), and (4) antler endochondral ossification (continued chondrogenesis and the appearance of antler velvet). In vitro studies have shown that insulin-like growth factor I (IGF-I) may be an important systemic regulator of pedicle formation as it stimulates proliferation of antlerogeneic cells from all four ossification stages. Interestingly, testosterone alone has no mitogenic eVect on these cells, even though there are specific binding sites for testosterone in this periosteum; however, in combination with IGF-I, it stimulates proliferation during stages 1 and 2 of pedicle ossification but reduces proliferation of cells from the fourth stage (Li et al., 1999). Initially, pedicles are covered by skin that is identical to that covering the rest of the skull. However, the transition between pedicle growth and antler development (stage 4) is marked by the appearance of characteristic antler skin described as ‘‘velvet’’ (Fig. 1). Velvet contains fewer hair follicles than normal skin does, but each of these has a sebaceous gland associated with it which gives it a ‘‘shiny’’ appearance and it contains no erector pili muscles. These first antlers then continue to elongate, generally as single unbranched spikes, by a modified endochondral process. However, a rise in circulating testosterone levels in autumn leads to cessation of growth, mineralization of the antler, and the consequent shedding of the velvet skin. This leaves a single unbranched antler (Fig. 2) attached to the pedicle until it is shed (cast) the following spring.

Figure 2 The first set of unbranched antlers grown by a red deer stag. These have completed their development and the velvet skin covering them has been shed.

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While the development of the pedicle and first antler has been described at a histological and ultrastructural level (Li and Suttie, 1994, 1998, 2000), the molecular mechanisms that regulate these processes have been little studied, not least because such studies present practical and ethical challenges. For example, a molecular characterization of antlerogeneic periosteum at diVerent developmental stages (before and after birth) needs to be undertaken to help understand the biology of this fascinating tissue. Comparing the molecular pathways which control development of the first antler and regeneration of subsequent antlers may also shed light on the extent to which developmental and regenerative pathways diverge. Furthermore, regenerating antlers in most deer species have branches whereas the first antlers to develop are single spikes and therefore such comparative studies could help to identify molecules that specifically control antler patterning, as distinct from those that control increases in antler size.

B. Antler Regeneration At this stage, it is worth considering why antlers regenerate. A number of explanations have been put forward and the interested reader should refer to Richard Goss’s review of the subject (Goss, 1983). One theory, originally proposed by John Hunter in the eighteenth century, was that stags shed their antlers before calves are born and are thus less likely to cause them harm. Perhaps a more rational explanation may lie in the adage that ‘‘size matters’’; antlers are secondary sexual characteristics and their function is to enable stags to achieve and maintain dominance over a harem of females. Regeneration enables antlers to increase in size each year as stags become more mature; it has been calculated that in red deer, antler size increases one and a half times as much as body size during the course of maturation (Huxley, 1926, 1931). An ability to regrow antlers damaged during fighting is also likely to have conferred a selective advantage. Interestingly, a study in red deer has shown that antler size is heritable and that stags with bigger antlers are the most successful breeders (Kruuk et al., 2002). However, over the 30-year study period, this selection did not generate an evolutionary response in antler size. This led the authors to conclude that environmental factors, in particular, nutrition, also have an important influence on antler size and success in combat (Kruuk et al., 2002). The annual cycle of antler loss and regrowth may also have evolved when deer moved to inhabit more temperate zones. If antlers were to retain their blood supply and continue to grow throughout the year, freezing weather conditions in winter would inevitably lead to tissue damage and necrosis. This could be one explanation for why antlers mineralize and become ‘‘dead’’ during the winter months. Another physiological challenge facing

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deer living in temperate habitats is the significant loss of body mass that takes place during winter. For deer stags (and other short-day breeders such as rams), this presents a particular problem as they have already lost a significant amount of body fat during the few frenzied weeks of sexual activity in autumn. Perhaps this explains why nature has developed a system that releases deer stags from the ‘‘tyranny of testosterone’’ for a few months during the summer, thus enabling them to increase their body mass and, at the same time, grow another set of antlers in time for the next mating season.

C. The Early Stages As described previously, a stag’s first antlers are normally shed in the spring of his second year of life and the process of antler casting has been shrouded in mystery for centuries because lost antlers are rarely found in the wild. In the Middle Ages, folklore had it that stags would deliberately hide their antlers in dark, concealed places in a forest whereas a more likely explanation is that they are tempting food for wild carnivores. Casting is a spontaneous event that appears not to be anticipated by the animal and involves osteoclasts resorbing bone at the interface between the solid antler and the pedicle. In most species, casting is coincident with regeneration; in fact, the velvet skin of the new antler is visible as a swollen ring around the base of the old antler (Fig. 3A). The local mechanisms controlling the process are far from clear although the process is regulated by a decline in circulating testosterone; a number of experimental studies have shown that deer maintained on artificially high levels of testosterone or estrogen do not cast their antlers and castration will result in casting at any time of year (Fletcher, 1978; Goss, 1968; Waldo and Wislocki, 1951). Thus, the trigger for casting appears to be increased osteoclast activity as a consequence of sex hormone withdrawal. As will be discussed in a later section, there is increasing evidence that the testosterone’s eVects on bone cells may be indirect following conversion by aromatase to estrogen (Meinhardt and Mullis, 2002) and we have some evidence that this conversion may also take place in antler tissues. It is known from studies in man and other animals that a decline in estrogen will increase bone resorption and that this is mediated by various cytokines that regulate osteoclast formation and activity (Riggs et al., 2002). Studies in man have shown that estrogen withdrawal is associated with increased synthesis of receptor activator of NF-kB ligand (RANKL), a potent activator of osteoclast diVerentiation (Eghbali-Fatourechi et al., 2003). We have found that RANKL is localized in cells in the blastema (Price and Allen, unpublished observations), although how its synthesis is systemically regulated needs investigation. The surface of the shed antler is concave and has no apparent blood supply (Fig. 3B), whereas the exposed surface of the pedicle bleeds

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Figure 3 Early stages of antler regeneration. (A) Pedicle (arrowhead) with the first antler still attached. The first signs of regeneration are evident as a ‘‘ring’’ of swollen tissue (arrow) at the base of the old antler. (B) Ventral convex surface of a cast antler that has been resorbed by osteoclasts. (C) The regenerating antler a few hours after casting showing the velvet skin (arrows) migrating over the surface of the exposed pedicle. (D) Antler bud 14 days after casting. The epidermis now covers most of the antler surface. The position of the future branches are visible as raised areas (arrows). (E) Section through the central region of a 4-day ‘‘blastema.’’ The wound epithelium (WE) can be seen under the scab migrating over a mass of ‘‘granulationlike’’ tissue and undiVerentiated mesenchymal cells. (F) Higher-power view of boxed region in (E), showing this cellular tissue which contains little extracellular matrix. (G) Section through the outer region of a 4-day blastema showing the transition from wound epithelium (WE) to mature epidermis containing hair follicles (H) and sebaceous glands (S). Scale bars: E,G, 200
m; F, 50
m.

and, within hours, a large scab forms and covers it (Fig. 3C). As in other situations where appendages regenerate, healing of an epidermal wound is required for the process to be initiated. A migratory epithelial layer rapidly covers the exposed pedicle (Fig. 3E) and within 7 to 9 days, epithelialization is complete and after approximately 10 to 14 days, future branches become visible as swollen raised areas at the periphery of this antler ‘‘bud’’ (Fig. 3D). Identifying the source of cells below the zone of amputation that develop into the early antler has been a subject of debate for many years. The tissues that make up the pedicle are skin, bone, periosteum, blood vessels, and nerves. Unlike the situation in the regenerating urodele limb, there is no muscle. Early experimental work involved amputation of the pedicle and/or surrounding regions of the skull (Goss, 1961; Jaczewski, 1955) and this showed that the ‘‘regeneration territory’’ is extensive; however, these surgical procedures involved significant trauma, which makes their interpretation diYcult. There have been no reports that transplantation of pedicle periosteum from a regenerating antler can induce antler growth at another site. However, what became clear from these transplantation studies was that skin of the pedicle cannot give rise to antler tissue; velvet skin transplanted to other sites will survive for several years but does not give rise to antler

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outgrowths (Goss, 1972). In one rather ingenious experiment, ear skin was transplanted onto the distal pedicle bone, which prevented pedicle skin from contributing to formation of the new antler (Goss, 1964). A normal antler formed and it was covered with antler velvet skin, which showed that epidermal diVerentiation is induced by underlying mesenchymal tissues. Notwithstanding, despite the lack of direct evidence, many antler biologists are of the view that the pedicle periosteum is likely to be a source of cells that form the regenerating antler (Kierdorf and Kierdorf, 1992, 2001; Li et al., 2004a,b). What is the evidence? Kierdorf et al. (1994) have shown that double head antlers (these form when the old antler fails to be cast) develop from the pedicle. More recently, Kierdorf et al. (2003) described thickening of the periosteum of the distal pedicle soon after antler casting, which would also suggest expansion of cell populations in this tissue. Our finding that parathyroid hormone-related peptide (PTHrP) is immunolocalized in both periosteum and in mesenchymal cells in the blastema is further evidence that the latter may be derived from the former (Faucheux et al., 1994). Li et al. (2004, 2005) have recently presented convincing morphological evidence that the pedicle bone appears not to contribute to the formation of the early antler and that ‘‘stem’’ cells from the pedicle periosteum proliferate and migrate onto the exposed pedicle surface following antler casting. This has led them to challenge the validity of using the term ‘‘blastema’’ in the context of antlers because they have concluded that cells which form the regenerating antler are stem cell-derived and do not appear to arise by a process of dediVerentiation (Li et al., 2004b, 2005). In contrast, a series of elegant experiments have shown that in newts, cells in the blastema arise by a process of dediVerentiation of mature cell types, including neural ependymal cells and multinucleated muscle fibers (Echeverri and Tanaka, 2002; Echeverri et al., 2001; Kumar et al., 2000; Lo et al., 1993). Thus, if a blastema is defined as a structure that arises exclusively by a process of dediVerentiation, then Li and colleagues are correct and the term should not be used to describe the early stage of antler regeneration. However, Richard Goss, the eminent regeneration biologist, in his excellent book on antler biology (1983), described antlers as ‘‘histologically complex structures derived from a mass of undiVerentiated cells that fits the definition of a blastema’’ and we have also recently described the early antler (<14 days) as a blastema. Goss’s definition of a blastema is ‘‘a rounded mass of cells endowed with the capacity to develop into structure replacing that which was lost’’ (Goss, 1983) and the Collins English Dictionary defines a blastema as ‘‘a mass of undiVerentiated animal cells that will develop into an organ or tissue: present at the site of regeneration of a lost part.’’ Thus, in our opinion, it remains valid to describe specific regions of the early antler as a blastema, or blastema-like, since they contain undiVerentiated cells; the fact that these cells may not arise by a process of dediVerentiation is not relevant. It is worth noting that there is no evidence

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Figure 4 Histology of a 14-day blastema and PCNA staining to show proliferating cells. (A) Section through a blastema at one of the future branches, showing that tissues at diVerent stages of diVerentiation can be distinguished: perichondrium (arrow), undiVerentiated mesenchymal cells (m), chondroprogenitors (cp), cartilage (ca), and bone (bo). Histology (B, D, F, H) and PCNA staining (dark brown nuclei in C, E, G, I) in the diVerent regions of the blastema. (B, C) Mesenchyme: In this very cellular region, a large number of undiVerentiated mesenchymal cells are proliferating; this is the equivalent of the ‘‘growth zone’’ in the growing tip of larger antlers. (D, E) Chondroprogenitors (cp): These cells are no longer dividing and become aligned in columns between which are vascular channels (v); however, there is cell

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for dediVerentiation and metaplasia in the regenerating tail of xenopus, where regeneration resembles normal tissue renewal, and yet it is described as containing a blastema of undiVerentiated tissue (Slack et al., 2004). However, we agree that it is incorrect to describe the entire early antler as a blastema; the term ‘‘bud’’ is more appropriate. It has proved diYcult to establish which cells in the pedicle give rise to regenerating antler tissues because studies of cell lineage, such as those applied in other models of regeneration, are practically diYcult to perform in antlers in vivo. For example, transgenic technology was used to establish cell lineage determination in the xenopus tail (Gargioli and Slack, 2004). In a review paper, Li and Suttie (2001) briefly describe an experiment in which they labeled antlerogeneic periosteum (AP) cells in vivo with LacZ and concluded from this that all cell types in the pedicle and primary antler appear to be the progeny of AP cells. What is not known, because the appropriate studies have not yet been done, is whether cells in the regeneration are derived from true stem cells, as suggested by Li et al. (2004), a strategy used by planaria (Agata et al., 2003; Sanchez Alvarado, 2003, 2004), from multipotent mesenchymal cells in the periosteum (Nakase et al., 1993), or from dediVerentiation of periosteal fibroblasts, osteoblasts, dermal fibroblasts, or nerve cells in the pedicle. Further transplantation studies could help to resolve this issue and these could involve not only whole tissue explants but populations of purified cell types isolated from the pedicle. Genomic and proteomic analyses of cells from antlerogenic petriosteum and its adult derivative would also help to identify the properties of these tissues. The anatomy and histology of the early antler ‘bud’ has recently been described by Li et al. (2004b, 2005) and clearly this is not a morphologically uniform structure like the blastema of regenerating newt limb (Brockes, 1997). For the first few days after casting, the center surface of the pedicle is composed of a granulation-like tissue and interspersed mesenchymal cells below a scab and regenerating wound epithelium (Fig. 3E, F). In contrast, at the periphery of the ‘‘doughnut’’, the pedicle skin is full thickness and contains appendages including hair follicles and sweat glands (Fig. 3G). There is little cell division at this stage and many cells express PTHrP. However, between days 3 and 4 and days 10–14 after casting, there is a significant increase in the number of mesenchymal cells, epithelialization is completed, and diVerent tissue regions are visible in tissue sections (Fig. 4A). There are distinct, relatively avascular ‘‘growth zones’’ of mesenchymal cells beneath the epidermis at sites where the main and subsidiary branches will division associated with perivascular tissue. (F, G) Cartilage: The mature chondrocytes (ca) are surrounded by extracellular matrix and do not divide. (H, I) Bone: In the proximal region where bone (b) is being formed, a proportion of osteoblasts (arrow) are proliferating. Scale bars: A, 1 cm; B, 12.5
m; C, E–I, 25
m; D, 50
m.

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Figure 5 Immunolocalization of parathyroid hormone-related peptide (PTHrP) in the early regenerating antler. Section through a blastema at day 14. (A) Skin: PTHrP is localized (brown stain) in epidermis (e), dermis (d), and around hair follicles (h). (B) Mesenchyme (m)/ chondroprogenitors (cp): The majority of cells stain positive. (C) Cartilage (ca): PTHrP is not present in mature chondrocytes but is localized in perivascular cells. (D) Cellular periosteum: As in the mesenchymal zone, a significant proportion of cells stain positive. Scale bars: A, 100
m; B, 50
m; C, D, 25
m. (See Color Insert.)

develop (Fig. 4B). These cells are proliferating (Fig. 4C) and are not highly diVerentiated as they do not express alkaline phosphatase (ALP), a marker of diVerentiated chondrocytes and osteoblasts. They also express PTHrP (Fig. 5A), which we consider to be a phenotypic marker for antler progenitor cells (Faucheux et al., 2004). Proximally, these mesenchymal cells diVerentiate into chondroblasts that align themselves into columns (Fig. 4D) interspersed with vascular channels. PCNA staining shows that these chondroblasts are not proliferating, although a population of PCNA-positive cells are seen lining vascular spaces. These may be proliferating endothelial cells associated with de novo angiogenesis (Clark et al., 2004) or could be derived from perichondrium/periosteum since RAR mRNA is expressed in the perivascular region of the blastema (Price and Allen, 2004) and in perichondrium (Allen et al., 2002). Lower down the antler bud, these chondroblasts diVerentiate into chondrocytes synthesizing cartilage matrix and this is associated with an increase in ALP activity and decreased synthesis of PTHrP (Fig. 5C). Coincident with the onset of chondrogenesis, increased remodeling activity by osteoclasts and osteoblasts leads to restoration of the integrity of resorbed bony trabeculae on the pedicle surface (Kierdorf et al., 2003). If blastema formation is a defining feature of epimorphic regeneration, so is the presence of a healing wound (Goss, 1983) and antlers will not regenerate if skin is grafted over the pedicle (Goss, 1972). This distinguishes epimorphic regeneration from the process of physiological regeneration

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(e.g., the replacement of red blood cells or skin cells) or tissue repair. For example, fracture repair involves regeneration of bone that is structurally and functionally normal but does not involve wound healing, and thus cannot be defined as epimorphic regeneration. Identification of the local factors that trigger regeneration in response to antler casting (the equivalent of amputation or injury in a newt) is clearly a priority. Pointers can be taken from work in other models of regeneration on the link between regeneration and tissue injury (Brockes, 1997; Brockes et al., 2001). For example, Jeremy Brockes’s group have identified activated thrombin, a critical regulator of the injury response, as being able to regulate cell cycle re-entry in newt myotubes (Tanaka et al., 1999) and in pigment epithelial cells (Simon and Brockes, 2002).

D. Ontogony Antlers elongate with a typical S-shape growth curve (Fig. 6). In the first month to 6 weeks, the branches of a regenerating antler form (Fig. 7) and, at this stage, the growth rate is relatively slow; however, during the summer, growth accelerates rapidly, then slows as autumn approaches. In larger species of deer, this rate of growth is spectacular; for example, in a moose,

Figure 6 The relationship between circulating testosterone (solid line) and antler growth (dotted line). Antlers are cast in the spring when circulating testosterone levels are low. In red deer, the early stages of antler growth do not appear to be dependent on sex steroids. However, as testosterone levels rise in late summer, longitudinal growth of the antlers slows, the bone becomes completely mineralized, and the velvet skin is shed. Polished hard antlers are then used for fighting during the autumn mating season.

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Figure 7 Two-year-old red deer stag with a set of regenerated antlers at approximately 30 days of growth. The major branches (tines) are now visible. The main branch (mt) is most caudal and the brow tine (arrow) most cranial.

antlers can grow up to 1.25 meters long and have a span that exceeds 2 meters. The shape and size is species dependent; small brocket deer which inhabit forests in South America, have simple unbranched antlers, species like red deer and wopiti, have complex branched ‘‘racks’’ (Fig. 1), whereas fallow deer and moose have palmate antlers. The main branch (beam) continues to elongate until the antler’s final size is achieved, whereas proximal branches (tines) of an antler will complete their development and become fully mineralized before the distal branches. Like the developing limb, antlers have a proximodistal axis, an anterioposterior axis, and a dorsoventral axis (Li and Suttie, 2001). Richard Goss demonstrated (1991) that cells in the antlerogenic periosteum control formation of the anteriorposterior axis, as he was able to show that rotation of the periosteum through 180 degrees resulted in antlers forming in a reversed orientation. The signaling molecules that control antler patterning are not known. One possible candidate is retinoic acid (RA), which controls formation of the zone of polarizing activity in the developing limb (Lu et al., 1997) and in the regenerating newt limb induces digit duplication (Scadding and Maden, 1986; Sessions et al., 1999) and conveys positional information (Maden et al., 1982; Pecorino et al., 1996). We have identified retinoic acids and retinoic acid receptors in the early antler bud (Price and Allen, 2004) and application of RA to the pedicle changes antler shape (Kierdorf and Kierdorf, 1998). PTHrP is another candidate, as it is expressed by a large proportion of mesenchymal cells in the antler blastema (Faucheux et al., 2004) and in the mammary gland regulates branching morphogenesis (Wysolmerski et al., 1995). Sonic hedgehog (Shh) is a key regulator of anterioposterior patterning in the limb, however, although we have detected

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the expression of indion hedgehog at sites of chondrogenesis in the antler (Faucheux et al., 2004), we have been unable to detect the expression Shh by RT-PCR. However, we suspect this may be a technical issue; antlers are large structures and if the site of Shh expression is very restricted and/or if its levels of expression are low, it would be easy to miss the critical tissue in any analysis. Alternatively, the lack of expression of Shh may reflect the diVerent embryological origins of the antler from the limb; as frontal bone derivatives antlers are likely to be neural crest-derived (although this has yet to be demonstrated) and Shh’s role in regulating patterning of head structures that are neural crest-derived is limited (Ahlgren and Bronner-Fraser, 1999; Hu and Helms, 1999). As far as the dorsoventral axis is concerned, Wnt-7a is an important regulator in the developing limb and although its expression has yet to be identified in the antler, we have evidence that other components of the Wnt signaling pathway are expressed in antlers (James Mount, unpublished observations).

E. Longitudinal Growth Longitudinal growth of antlers takes place at the distal tip of each branch (Fig. 8, Fig. 9), where mesenchymal cells proliferate and subsequently diVerentiate into chondrocytes. The cartilage zone is very extensive and consists of chondrocytes arranged in trabeculae which proximally become mineralized and then form a scaVold for new bone formation. At the same time, new bone is laid down circumferentially along the antler shaft by intramembranous ossification. Thus, the whole temporal and spatial sequence of cellular diVerentiation that takes place during endochondral and intramembranous bone growth is represented in one antler branch. Inevitably, the anatomy of the growing antler tip has been compared to that of the epiphyseal growth plate but there are a number of important diVerences. What is immediately apparent, even with the naked eye, is that the antler tip is a highly vascularized structure (Fig. 8), whereas epiphyseal cartilage is not. The main artery supplying the antler is a branch of the temporal artery whose branches and anastomoses remain superficial within the dermis and direct blood to the distal tip. Blood is then channeled vertically downward into cartilage and bone. Blood in the large vascular spaces in the center of the antler then drains into the pedicle or into large veins in the velvet skin. Another important diVerence between the antler and a growth plate is that the separation of cells into defined zones is far less distinct. In the antler tip, there is a region where cells proliferate (Faucheux et al., 2004; Matich et al., 2003); however, these cells are not diVerentiated whereas cells in the proliferating zone of the growth plate are chondrocytes (Farnum and Wilsman, 1993; Loveridge and Farquharson, 1993). This ‘‘growth zone’’ is

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Figure 8 Histology of the growing tip of a regenerating antler. (A) Longitudinal section through the distal end of the main branch of an antler at approximately 35 days of growth. vs: velvet skin, m: mesenchyme, cp: chondroprogenitors, ca: cartilage, min ca: mineralized cartilage, ps: primary spongiosa, # sites of intramembranous bone formation below the periosteum. (B) Section through skin: epidermis (e), dermis (d), hair follicles (h), and sebaceous glands (s). (C) The junction between fibrous perichondrium (fpc) and mesenchyme (m). (D) Junction between fibrous periosteum (fpo) and cellular periosteum (cpo). (E) Shows intramembranous ossification at the periphery of the antler shaft; new bone (b) is formed by osteoblasts (arrows). (F) Region of chondroprogenitors (cp), vascular channel (v). (G) cartilage (ca). (H) Junction between mineralized cartilage and primary spongiosa. Cartilage trabeculae are resorbed by osteoclasts (shown by arrows at higher power in inset box) and new bone is being formed by osteoblasts (asterisk). (I) Von Kossa-stained section (brown/black) to show mineralization of cartilage trabeculae. (J) Spongy bone (b) in the midshaft of the antler. Scale bars: A, 2 cm; B, 400
m; C-J, 50
m; insets H, J, 10
m.

Figure 9 Characteristics of mesenchymal cells in a rapidly growing regenerating antler. (A) Cryostat section through an antler tip showing low levels of staining for alkaline phosphatase (ALP (red stain)) in skin (s), perichondrium (p), and mesenchyme (m). ALP activity increases in cartilage (c). (B) Cells cultured from this region show low levels of staining for ALP. (C) In the presence of betaglycerol phosphate (BGP) and ascorbic acid, long-term cultures of mesenchymal cells will form mineralized nodules as detected by Von Kossa staining (black, indicated by asterisk). Scale bars: A, 1 cm; B, 10
m; C, 50
m.

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a relatively avascular region composed of tightly packed small cells (Fig. 8C) below a fibrous perichondrium which is continuous with the fibrous periosteum covering the lower antler shaft. Interestingly, whereas type I collagen mRNA (a phenotypic marker of cells of the osteoblast and fibroblast lineages) is highly expressed in the fibrous perichondrium, the hybridization signal is much weaker in these mesenchymal cells, consistent with there being little matrix synthesis (no metachromatic staining). The nomenclature used to describe this region can be somewhat confusing. It has been variously described as ‘‘hyperplastic perichondrium’’ and ‘‘reserve mesenchyme’’ (Banks and Newbrey, 1983), and we also describe this as the mesenchymal zone. However, we have also, perhaps rather confusingly, described cells cultured from the site as being ‘‘perichondrium-derived’’ because they are dissected from below the fibrous perichondrium. PCNA staining has shown that in both the regenerating and developing antlers these mesenchymal cells are dividing rapidly (Faucheux et al., 1994; Matich et al., 2003) and, in culture, these cells (Fig. 9) have a doubling time of 24 to 48 hours and will even continue to proliferate in low concentrations of serum (Price and Faucheux, 2001). We have also shown that these cells have an extended life span in culture (they can be cultured for over 80 passages), a characteristic which may underpin the ability of antlers to grow at such phenomenal rates (Price and Faucheux, 2001). In vivo and in vitro (Fig. 9A), these cells express only low levels of alkaline phosphatase (Price and Faucheux, 2001; Price et al., 1994) and do not express type II collagen (Price et al., 1996) or aggrecan protein, which reflects their lack of diVerentiation. In fact, like stromal cells derived from bone marrow (Huttmann et al., 2003), these cells appear to have the capacity to diVerentiate along more than one lineage. For example, dexamethasone, a factor that is known to induce osteoblast diVerentiation in marrow stromal cells (Beresford et al., 1994), will induce alkaline phosphatase in antler mesenchymal cells (Price and Faucheux, 2001). In contrast, when cultured in the presence of rabbit serum, adipocyte-like cells will diVerentiate. When cultured for longer periods in the presence of betaglycerol phosphate and ascorbate, these cells will also form mineralized collagenous nodules (Fig. 9C). Factors that regulate the growth of these cells will be discussed in a later section.

F. Chondrogenesis For many years, there was controversy among antler biologists as to the type of ossification taking place in growing antlers. For example, only 24 years ago, Beresford (1980) classified unmineralized tissues in antler as chondroid bone and concluded that antler cartilage was neither hyaline, elastic, nor fibrocartilage. In contrast, a series of histological and ultrastructural studies

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concluded that antler development involved a modified form of endochondral ossification and intramembranous ossification (Banks, 1974; Frasier and Banks, 1973; Frasier et al., 1975). The problem was that, until 1996, only types I and III collagen had been reported to be present in antler cartilage (Newbrey et al., 1983; Speer, 1983), and these collagens are not characteristic of normal hyaline cartilage. However, we were able to demonstrate by in situ hybridization and immunocytochemistry that antler chondrocytes express type II collagen (Fig. 10E), a marker of the chondrocyte phenotype (Price et al., 1996) and type II collagen has also been identified

Figure 10 Characteristics of the cartilage region of a rapidly growing regenerating antler. C: cartilage. V: vascular spaces. (A) In situ hybridization for collagen I (black silver grains) shows that expression is maintained in the ‘‘transition zone’’ where cells start to diVerentiate into chondroprogenitors; note that cells start to become aligned in columns. (B) In situ hybridization for collagen I (black silver grains) shows that expression is now restricted to cells in perivascular tissue that are likely to be of the osteoblast lineage. (C) Alcian blue staining to show proteoglycans in cartilage matrix. (D) Staining for alkaline phosphatase in cryostat sections (red) shows many positive cells associated with the vascular channels. (E, F, G, H) Immunolocalization of collagen types II (E), VI (F), X (G), and aggrecan in cartilage (red stain).

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biochemically, albeit at low levels (Rucklidge et al., 1997). Histologically the boundary between the zone of dividing mesenchymal cells and the onset of chondrogenetic diVerentiation is not distinct; there is a gradual increase in cell size, decreased proliferation, increased synthesis of extracellular matrix, and cells begin to assume a columnar arrangement (Figs. 8 and 10A). We define the onset of chondrogenic diVerentiation as where cells start to express type IIA collagen mRNA, which transiently precedes the onset of type IIB collagen expression (Price et al., 1996). Type IIA collagen is a splice form of type II procollagen synthesized primarily by chondroprogenitor cells in developing cartilage but is not expressed by mature growth plate chondrocytes (Sandell et al., 1994, 2001). However, chondroprogenitors in this region continue to express type I collagen mRNA (Fig. 10A) and thus this is a ‘‘transition zone’’ with poorly defined boundaries. Interestingly, the synthesis of type I and type II collagen has been demonstrated in the cartilage blastema of the developing chick limb between stages 24 and 26 and the predominant form of type II procollagen is the type IIA isoform (Nah et al., 1992). However, whereas type IIB collagen mRNA expression is maintained in chondrocytes throughout antler cartilage, the expression of type IIA is transient; it only persists into the upper regions of longitudinal trabeculae (Price et al., 1996). Intensity of type II collagen immunoreactivity also increases as chondrocytes hypertrophy in more proximal trabeculae. Antler cartilage is a very abundant source of cells for in vitro studies; in a rapidly growing antler, up to 100 million cells can be harvested from nonmineralized cartilage. Cells cultured from antler cartilage have a slower rate of growth than cells from the growth zone (doubling time of 48–96 hours) and undergo replicative senescence after approximately 20 passages. They also express higher levels of alkaline phosphatase than cells cultured from the growth zone, reflecting their more diVerentiated state (Price et al., 1994) (Fig. 11A). However, antler chondrocytes cultured as monolayers synthesize high levels of type I collagen and lose ALP expression with serial passaging, reflecting their tendency to dediVerentiate rapidly. They maintain their phenotype far better when cultured as high-density micromasses (Fig. 11E, F) but continue to synthesize type I as well as type II collagen. Interestingly, antler chondrocytes synthesize very little type X collagen in vitro (unpublished observations), despite the abundance of this collagen in cartilage in vivo (Gibson et al., 1996). Type X collagen is present throughout nonmineralized and mineralized cartilage (Fig. 10F) and thus is distributed far more widely in antler cartilage than in growth plate (Faucheux et al., 2004; Price et al., 1996). The problem with the antler tip, unlike the growth plate, is that it is very diYcult to define the onset of chondrocyte hypertrophy. If it is taken to be the site where type X collagen is first expressed, then essentially most chondrocytes in antler can be considered to be hypertrophic, since chondrocytes which express type IIB collagen also express type X

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Figure 11 In vitro characteristics of cells cultured from antler cartilage. (A) Primary cultures of cells cultured from antler cartilage have high levels of alkaline phosphatase activity but lose this with serial passaging. (B) High-density monolayer of chondrocytes showing proteoglycan synthesis (alcian blue stain). (C) Cells cultured from cartilage synthesize aggrecan (red stain). (D) Mineralized nodules (*) form in cartilage-derived cells cultured with BGP and ascorbate. (E, F) Antler cartilage-derived cells cultured as micromasses showing high levels of alkaline phosphase activity (red stain) and proteoglycan synthesis (alcian blue stain). Scale bars: A, B, 50
m; C, D, 50
m; E, F, 1 mm. (See Color Insert.)

collagen. In contrast, in mammalian growth plate, type X collagen is only expressed in the lower hypertrophic zone (Reichenberger et al., 1991; Sandell et al., 1994). In fact, the pattern of type X synthesis in antler is similar to that in the developing avian limb, where endochondral ossification occurs in an irregular pattern along the entire diaphysis and the zone of hypertrophic cartilage is far more extensive than in mammalian cartilage (Kwan et al., 1989; Oshima et al., 1989). Because type X collagen is first expressed in antler cartilage at least 1 cm distal to where mineralization of cartilage takes place, this suggests that type X does not play a role in initiating the process. Since type X collagen mRNA is first expressed where there is coincident enlargement of vascular channels, this suggests that it may play some role in promoting formation of the extensive network of these channels in the antler tip. Another usual feature of antler cartilage is that it contains type VI collagen (unpublished observations) (Fig. 10E); however, the significance of this is not known. Another very important matrix component of antler cartilage are glycosaminoglycans (GAGs) (Sunwoo and Sim, 2001) that can be detected by alcian blue staining, which increases in intensity as chondrocytes diVerentiate

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(Fig. 10C). The major GAG in antlers is chondroitin sulfate (Sunwoo et al., 1998a,b) which is an important component of the large aggregating proteoglycan aggrecan, a marker of the chondrocyte phenotype (Heinegard et al., 1985). We have found the spatial expression of aggrecan core protein mRNA expression to be similar to that of type IIB collagen mRNA, although the level of expression is lower (unpublished observations). Aggrecan immunoreactivity is also strong in diVerentiated chondrocytes (Fig. 10H), whereas only faint immunoreactivity is detected in chondroprogenitors in the transition zone. Cultured chondrocytes also synthesize aggrecan (Fig. 11C), whereas undiVerentiated cells from the mesenchyme do not. Another important diVerence between antler and growth plate is that there is no type I expression in growth plate (Fukunaga et al., 2003; Sandell et al., 1994), whereas type I collagen mRNA is expressed throughout antler cartilage (Price et al., 1996) in a population of small cells in perivascular tissue adjacent to the columns of chondrocytes (Fig. 10B). This evidence, and the high level of ALP activity in this region, has led us to conclude that these cells are of the osteoblast lineage, the progenitors of diVerentiated osteoblasts found in primary spongiosa. These perivascular cells also express the retinoic acid receptor RAR and the retinoic acid synthesizing enzyme RALDH2 (Allen et al., 2002), which suggests that retinoic acid regulates the diVerentiation of osteoblasts in antlers. The perivascular tissue is also a site of osteoclast diVerentiation and this will be discussed in more detail in a later section.

G. Regenerating Antlers: Ossification and Remodeling As has been mentioned, the zone of nonmineralized cartilage in antlers is extensive; in red deer, it can extend for than more 70 cell layers proximodistally. Calcification of cartilage, as detected by alizarin red staining or von Kossa staining (Molello et al., 1963), occurs initially as discrete foci in the middle of trabeculae, at a depth of
1.5 cm from the top of the cartilage trabeculae. These foci of mineralization then coalesce to completely surround chondrocytes and eventually the entire cartilage spicule contains mineral (Fig. 8I). Matrix vesicles, another important component of epiphyseal growth plate cartilage that contain high levels of the enzyme ALP, are present in antler cartilage and act as an initial focus for mineralization (Newbrey and Banks, 1975). ALP activity can be detected in sections of antler cartilage (Fig. 9A) and bone (Kuhlman et al., 1963; Molello et al., 1963; Price et al., 1994) and cells cultured from cartilage have significant amounts of ALP activity and will form mineralized nodules in vitro (Fig. 11A, D). Interestingly, the ALP activity of cells from antler cartilage is high (up to 3.6 mmol/mgprotein/minute) compared to levels recorded for other skeletal cell types, including rabbit growth plate chondrocytes

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(0.065 mmol/mg protein/min) (Boyan et al., 1988) and is likely to reflect a high rate of cellular diVerentiation in antlers. In more proximal regions calcified cartilage is resorbed by osteoclasts and this leads to trabeculae losing their typical columnar arrangement to become irregularly shaped. Banks and Newbrey (1983) describe this region as the ‘‘primary spongiosa.’’ Eventually, cartilage trabeculae are replaced by woven bone that is remodeled into lamellar bone. At the same time, woven and lamellar bone are being formed around the antler shaft by a process of intramembranous ossification. Compact lamellar bone is found only in the periphery of the antler shaft whereas cancellous lamellar bone occupies the midshaft (Fig. 8J). Osteoblasts at sites of both endochondral and intramembranous ossification synthesize osteocalcin, a marker of diVerentiated osteoblast phenotype (Allen et al., 2002). Antler bone contains both all-trans and 9-cis-retinoic acid and the RA synthesizing enzyme RALDH2, which suggests that RA may control the activity of mature osteoblasts as well as the diVerentiation of osteoblast progenitors. Antlers are similar to other developing bones in being dependent on osteoclasts to remodel mineralized matrix and facilitate the penetration of large vascular channels. However, the unparalleled rate of endochondral ossification in antlers requires very large numbers of osteoclasts to diVerentiate locally. Perivascular tissue in nonmineralized cartilage is the site where osteoclastic diVerentiation is initiated since it contains cells that express tartrate-resistant acid phosphatase (TRAP) and vitronectin receptors (VNRs) (Fig. 12), phenotypic markers of osteoclasts (Faucheux et al., 2001; Price et al., 1994, 1996). As expected, the number of TRAP-positive cells increases as cartilage mineralizes and is replaced by bone (Szuwart et al., 2002). Interestingly, TRAP-positive cells are also present in mesenchymal tissue of the early antler bud. This regulation of osteoclast formation by chondrocytes/mesenchymal cells has also been shown to occur in fetal bones (Thesingh and Burger, 1983; Van De Wijngaert et al., 1989). Molecules responsible for regulating osteoclast diVerentiation are PTHrP and RANKL; both are expressed in perivascular tissues and in antler cartilage (Faucheux et al., 2001, 2002). Macrophage colony stimulating factor (M-CSF) (Faucheux et al., 2001) and transforming growth factor beta (TGF-) (Faucheux et al., 2004) are also present in antler cartilage in vivo and are also likely to support osteoclastogenesis. When chondrocytes from nonmineralized cartilage are cultured at high density in micromasses (200,000 cells per 10
l), they support the formation of multinucleated osteoclast-like cells that have the phenotypic characteristics of mammalian bone-derived osteoclasts (Fig. 12). They express TRAP, VNRs, calcitonin receptors (the ‘‘gold standard’’ of osteoclast phenotypic markers) and, when cultured on a mineralized substrate, form F-actin rings and large resorption pits (Faucheux et al., 2001). We have found this

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Figure 12 Micromass cultures of cartilage-derived cells support the diVerentiation of osteoclasts. (A) Cryostat section of antler cartilage stained for TRAP showing numerous positive cells (red stain) adjacent to the vascular channels (V). (B) TRAP-stained osteoclasts in micromass culture after
14 days. (C) Confocal image of a diVerentiated osteoclast cultured on a dentine substrate. The vitronectin receptor, a specific osteoclast marker, and an actin ring is present, indicating that the osteoclast is actively resorbing. (D) Resorption pits formed by antler osteoclasts in a micromass culture. Scale bars: A, 100
m; B, 25
m; C, 10
m; D, 1 mm.

model to be very reproducible and now use it routinely for investigating the molecular mechanisms that regulate osteoclast diVerentiation. What is particularly useful about this system is that chondrocytes cryopreserved immediately after extraction from an antler can support osteoclastogenesis. However, the numbers of cells that form ‘‘spontaneously’’ is reduced if cells have been previously frozen, and then it is normally necessary to ‘‘drive’’ osteoclastogenesis with exogenous factors such as PTHrP or RANKL. In contrast, in ‘‘fresh’’ micromass cultures, osteoclasts will form spontaneously in the absence of factors normally required to stimulate osteoclasts in vitro (1,25(OH)2D3, PTH, M-CSF, RANKL, etc.). This may reflect the high concentration of PTHrP synthesized into conditioned medium (Faucheux and Price, 1999), a very potent stimulator of antler osteoclast formation, and/or the expression of RANKL and M-CSF in these micromass cultures (Faucheux et al., 2001, 2002). PTHrP’s eVect on osteoclastogenesis is partially mediated via RANKL as OPG, the soluble decoy receptor for RANKL (Simonet et al., 1997; Tsuda et al., 1997), and can partially inhibit PTHrP-induced osteoclast diVerentiation (Faucheux et al., 2002). However, unlike most mammalian osteoclasts, antler osteoclasts express PPRs, evidence that there may be direct eVects of PTHrP on antler osteoclasts (Faucheux et al., 2002). Retinoic acid is another factor that can regulate osteoclast formation in these micromass cultures (unpublished observations) and, as will be described in more detail in a later section, retinoic acids are present and are synthesized in perivascular tissues of antler cartilage.

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II. Regulation of the Antler Development and Regeneration A. External and Systemic Factors Since antlers are secondary sexual characteristics, their annual cycle of regeneration has evolved to be closely coordinated to the reproductive cycle (Lincoln, 1971) and there is a fixed relationship between the stage of the antler cycle, plasma testosterone, testis diameter, and body weight (Fig. 6). The regulation of the annual cycle of antler growth was studied extensively in the 1970s and 1980s, with a number of groups worldwide contributing to advancement of the field. In temperate species such as red deer, which have defined mating seasons, the environmental cue which triggers changes in reproductive activity is the photoperiod (Lincoln and Short, 1980). In these species, decreasing day length triggers pituitary gonadotrophin activity for 2 months before the autumn mating season (Lincoln and Kay, 1979). There have also been studies to indicate an activation of reproductive hormones in the spring (Bubenik et al., 1982; West and Nordan, 1976). However, during the spring increasing day length does not activate reproduction; instead, it is responsible for initiating antler growth. In fact, it may be the change in photoperiod that is the critical trigger because, under laboratory conditions, the antler cycle can be initiated by either increasing or decreasing day length (Goss, 1976). Notwithstanding, in seasonally breeding deer, the photoperiod would appear to modify rather than act as a ‘‘main driver’’ of antler growth cycle since there is an endogenous rhythm to antler growth. For example, animals maintained on less than 24 hours of continuous lighting show cycles of antler growth and reproductive activity (West and Nordan, 1976). In a study which maintained stags under 8 hours of light and 12 hours of dark, antlers were cast 5 months before those in a control group kept under natural lighting conditions. This occurred because a second luteinizing hormone (LH) peak in the spring did not take place and the consequence of this was a fall in plasma testosterone and premature antler casting (Suttie et al., 1984). The mechanism whereby the photoperiod is linked to antler growth and reproductive activity is not clear but may involve changes in melatonin secretion from the pineal gland, which then acts on the hypothalamopituitary axis (Bubenik et al., 1986). In young animals, the pineal gland may play a particularly important role as pinealectomy leads to changes in the antler growth cycle (Plotka et al., 1984) as does the hormone melatonin (Lincoln et al., 1984). In red deer, exogenous melatonin has similar eVects to that of a short photoperiod, namely, advanced growth of the testis and premature cessation of growth (Webster et al., 1991). In contrast, this photoperiodic-pineal link to the reproductive system is absent, or of only

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minor significance, in species of deer from subtropical or tropical zones (Loudon and Curlewis, 1988). 1. Sex Steroids As discussed earlier, testosterone is required for pedicle and first antler development but, in the first year of life, hormone secretion is not regulated by photoperiodic cycles. Sex hormones are also generally considered to be the most important internal regulators of the timing of the annual cycles of regeneration, although they clearly have to function in a complex endocrine environment and in the context of other internal influences such as genotype, which plays a major role in shaping antler size and shape. As illustrated in Fig. 6, the antlers are cast in the spring, when testosterone levels are low and day length is increasing (Lincoln and Kay, 1979). The growth of the new antlers starts slowly in the spring but accelerates exponentially in the summer when testosterone levels remain low; in fact, studies in red deer have shown that this phase of antler growth appears not to require sex hormones (Fennessy et al., 1988; Suttie et al., 1989). However, this is an area where some controversy exists, as Bubenik and colleagues have argued that testosterone, possibly from nongonadal sources, may be required for initiating regeneration and for regulating longitudinal growth (Bartos et al., 2000). What is not in dispute is that as testis size increases and testosterone levels rise above 1 ng/ml, antler growth slows and mineralization of antler bone increases (Muir et al., 1988). The velvet skin is then lost (Fig. 13) and antlers are ‘‘polished’’ in preparation for the mating (rutting) season, with testosterone levels peaking in October and November in most species of deer (Lincoln, 1971). The long-held assumption has been that antlers lose their blood supply after they have completed velvet shedding and are thus ‘‘dead.’’ However, it has been shown that a functioning vascular system and living cells remain in the core of the hard antlers of fallow deer (Rolf and Enderle, 1999). After the mating period (rut) is over, testosterone levels decline and, in species such as reindeer, moose, and caribou, this is associated with antler casting in early winter; however, in most species, this does not occur until the following spring when testosterone levels fall below 1 ng/ml (Muir et al., 1988). Deer maintained on artificially high levels of sex steroids do not cast their antlers (Fletcher, 1978; Wislocki et al, 1947), whereas orchidectomy (castration) when a stag has hard antlers leads to their premature casting (Jaczewski et al., 1976), as will administration of cyproterone acetate, a specific blocker of androgen receptors (Bubenik et al., 1984). The eVects of castrating a stag when his antlers are growing provide further evidence that sex steroids are key regulators of the start and end of

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Figure 13 Shedding of velvet skin in late summer in a 2-year-old red deer stag.

the cycle of antler growth. However, the eVects of androgen withdrawal are species specific; in reindeer, which are considered to be a phylogenetically primitive genus, castration has little eVect (Bubenik, 1983), whereas in phylogenetically intermediate red deer, the first set of antlers grown by a castrated stag often appear normal but they do not complete their mineralization nor shed their velvet. This means that temperature changes in winter can lead to tissue damage which can have an important eVect on antler structure; castrate antlers have been reported to develop multiple branches and/or get bigger. In contrast, in species such as roe deer which have the longest phylogenetic development, castrate antlers can develop into large benign tumors and thus greatly incapacitate the animal bearing them (Bubenik, 1983; Goss, 1983). Why androgen withdrawal should have such species-specific eVects is not known, but it raises intriguing questions relating to the phylogeny of the function of sex steroids. The castrate antlers of roe deer also raise questions about the relationship between regeneration and cancer and, as such, merit further study. Despite the obvious importance of testosterone as an ‘‘internal driver’’ of antler regeneration, its mechanism(s) of action are not well understood. Androgen receptors have been immunolocalized in antlers (J. Price, unpublished observations); however, Li et al. (1999) failed to demonstrate any eVect of testosterone on the proliferation of antler cells, nor does testosterone sensitize antler cells to the proliferative eVects of IGF-I (Sadighi et al., 2001). However, studies that we have undertaken (unpublished) have shown that testosterone will induce the proliferation of cells cultured from antlers and 17- estradiol has a similar eVect, although the proliferative response

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appears to depend on the stage of antler growth and the stage of cell diVerentiation. Estrogen receptors have been localized in antler tissues (Barrell et al., 1999; Lewis and Barrell, 1994) and in vivo studies undertaken many years ago showed that when administered to castrated stags, animals exhibit rutting behavior and fail to cast their antlers (Fletcher and Short, 1974). Richard Goss demonstrated over 30 years ago that estrogen would induce premature mineralization of antlers in male Sikka deer and was actually more eVective than testosterone (Goss, 1968). Bubenik and Bubenik (1978) reported that administration of an estrogen receptor antagonist to white tail deer stags during growth leads to decreased thickness of compact bone of the antler shaft. These diVerent lines of evidence suggest that testosterone may have indirect eVects on antler cells in vivo, via its conversion to estrogen by aromatase. We have undertaken an experiment similar to that undertaken by Richard Goss in red deer stags and administered 17- estradiol on a single occasion during the phase of rapid growth; longitudinal growth stopped and within 2 weeks the antler was solid bone (J. Price, unpublished observations). These observations suggest that estrogen’s role is to prevent continued antler growth from the distal antler tip; this would be consistent with ERs being present at this site. Thus estrogen’s role in the antler would be similar to its role in the growing male human skeleton; a man lacking ER was found to be profoundly osteopenic and did not stop growing in height after the end of puberty. A similar phenotype has been described in patients lacking a functional aromatase gene (Carani et al., 1997; Morishima et al., 1995). That the function of ER should be conserved is perhaps not surprising as that the ER has been shown to be the most ancient steroid receptor (Thornton, 2001). 2. Insulin-like Growth Factor If sex steroids have a minor role to play early in the cycle when antlers are growing rapidly, this begs the question as to the identity of the ‘‘antler growth stimulus’’ (AGS) that was proposed by Wislocki (1943). Over the years, growth hormone, thyroxine, and prolactin and testosterone have been proposed as likely candidates. However, important work by the group of Suttie et al. (1985) led to the conclusion that IGF-I is likely to be AGS. Several studies have now shown a positive correlation between antler growth and circulating IGF-I (Suttie et al., 1985). IGFs I and II have also been detected at the mRNA level, by PCR, in antler tissues (Francis and Suttie, 1998). Furthermore, ligand binding studies have shown that receptors for both IGF I and IGF II are present in the antler tip (Elliott et al., 1992, 1993) and both IGF I and IGF II promote proliferation of cultured cells of the progenitor and cartilage regions of antler in vitro (Fig. 14; Price et al., 1994; Sadighi et al., 1994). There is evidence for an association between

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Figure 14 The eVect of FGF-2 and IGF-I on antler cell proliferation. Cells from mesenchyme and cartilage were cultured as monolayers in 48 well plates. When subconfluent, cells were serum deprived for 12 hours, then treated for 48 hours with FGF-2 (A) or IGF-I (B) in BGJb medium containing 2% fetal bovine serum. Cultures were pulsed for the final 12 hours with 3H thymidine and incorporated into the cells assayed as described previously (Price et al., 1994). IGF-I stimulates proliferation in both cell types whereas FGF-2 only stimulates growth of the less diVerentiated progenitor cells. *p < 0.05, **p < 0.02, ***p < 0.001, ****p < 0.0001.

IGF-I concentrations and serum testosterone (DitchkoV et al., 2001; Li et al., 2003), although whether previously elevated plasma testosterone levels are directly responsible for the subsequent IGF-I peak remains unclear. There is also evidence that IGF-I concentrations can be influenced by the photoperiod (Suttie et al., 1991). The source of IGF-I that regulates antler growth is likely to be the liver, where its secretion is regulated by growth hormone (GH); and in red deer stags pulsatile increases in GH precede by one month increases in circulating IGF-I, weight gain, and antler growth (Suttie et al., 1989). There have been no reports describing the localization of IGFs or IGF-binding proteins in vivo or in vitro, and stags surgically prevented from growing antlers have elevated IGF-I levels, an observation that lead Suttie et al. (1988) to propose that, although the antler is a major target for IGF-I, antler cells are not a

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major source of the growth factor. However, the importance of locally produced, compared with circulating, IGFs remains unclear and deserves further investigation. In fact, the role of IGFs in antler should be revisited in light of recent advances in the biology of IGFs and their receptors, particularly in the context of regeneration. For example, muscle-specific expression of insulin-like growth factor I has been shown to preserve regenerative capacity in senescent skeletal muscle (Musaro et al., 2001) and to preserve muscle function in mice lacking dystrophin (Barton et al., 2002).

B. Local Mechanisms One of the keys to understanding antler development and regeneration is to identify the pathways that control the expansion of progenitor cell populations, cell diVerentiation and survival, and epithelial–mesenchymal interactions. Antlers are also unique among regenerating structures in that the interaction between systemic factors and the local signaling pathways has also to be considered. Evidence has accumulated that developmental programs can be recapitulated in the adult under specific circumstances, and studies in other models of regeneration have shown that many developmental signaling molecules are expressed in regenerating tissues (Brockes, 1997; Holstein et al., 2003; Poss et al., 2003; Slack et al., 2004). It is also necessary to understand how injury (in the case of antlers, the process of antler casting) will ‘‘trigger’’ these pathways. Unfortunately, not a great deal is currently known about the mechanisms that control antler development and regeneration. However, this situation is changing since recent technological advances are applied in antler research. For example, the group in New Zealand has generated a database of genes expressed in regenerating red deer antler tissues (Lord et al., 2001, 2004). They have identified 4516 expressed sequence tags (ESTs) that include 930 gene sequences. The first proteomic analysis of growing red deer antler has recently been completed (Park et al. 2004) and this identified
130 proteins including structural proteins, matrix proteins, metabolic enzymes, and signaling and cell growthrelated proteins. The main driver of this type of research is industry; the successful marketing of velvet antler as a nutraceutical/therapeutic product depends on the components of antler being identified and on studies being undertaken to establish eYcacy. The caveat is that although these ‘‘fishing trip’’ approaches will identify the molecules involved, dissecting out their function is extremely labor intensive and not straightforward in an animal that has limited genetics. Unfortunately, there are probably not enough research scientists working in the field to take full advantage of the huge amount of information that these genomic and proteomic studies will generate. A second caveat is that industry is interested in identifying factors that

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regulate the growth of fully formed, rapidly growing antlers, because these provide the starting product for antler extract production. Therefore, practically nothing is known about the pathways that regulate the very early stages of antler development/regeneration. This area will remain neglected unless more support for basic biological research is forthcoming. Unfortunately, many funding bodies are wary of supporting research that utilizes animal models that are not mainstream and which are ‘‘genetically intractable.’’ In addition to components of extracellular matrix, a number of growth factors, cytokines, hormone receptors, and other signaling molecules have been identified in antlers (the most important are listed in Table I), although there is very little experimental evidence regarding their function. It is also Table I Growth Factors, Hormone Receptors, and other Signaling Molecules Shown to be Present in Antler Tissues Molecule EGF EGF, EGF receptor 1,25(OH)2D3

Deer species

Protein/mRNA

Reference

Red deer antler

protein

Cultured roe deer antler cells Red deer antler

protein

BMP-2, BMP-4 BMP-2, BMP-4, BMP-14, BMPRI, ACTRIII Neurotrophin-3 c-myc, c-fos TGF-1, TGF-II TGF-1 IGF-I, IGF-II IGF I & IGFII receptors

White tail deer Red deer antler

mRNA protein

Red deer antler Red deer antler Red deer antler

mRNA mRNA mRNA protein mRNA protein

RANKL & M-CSF Dermatopontin, glutathione peroxidase all-trans, 9-cis RA RARs& RXRs RALDH2 PTHrP, PTHrP/PTHR, Indian hedgehog (IHH) FGF-2, FGFR1, FGFR2, FGFR3 VEGF, KDR

Red deer antler Red deer antler

mRNA mRNA

Red deer antler

Allen et al. (2002)

Red deer antler

protein mRNA protein protein

Red deer antler

protein

Barling et al. (2004)

Red deer antler

protein

Clark et al. (2004)

Estrogen receptor

Red deer antler

protein

Barling et al. (2004); Ko et al. (1986) Sempere et al. (1989) Barrell et al. (1999); Lewis et al. (1994); Park et al. (2004) Barling et al. (2004); Feng et al. (1995, 1997) Garcia et al. (1997) Francis et al. (1998) Faucheux et al. (2004); Francis et al. (1998) Elliott et al. (1992, 1993); Francis et al. (1998) Faucheux et al. (2000) Lord et al. (2001)

Faucheux et al. (2004)

Note: this does not include the 130 proteins identified in the recent proteome analysis of red deer antler (Park et al. 2004).

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diYcult to draw a consensus from the literature when various species of deer and diVerent stages of antler development/regeneration have been used for such studies. The approach that we have taken has been to focus on molecules that are known to play an important role in development and/or regeneration in lower vertebrates and to use a variety of techniques (cell culture, in situ hybridization, immunolocalization, etc.) to try and explore their role in red deer antlers at diVerent stages of regeneration. A class of molecules that are important regulators of skeletal development and regeneration in urodeles are the retinoic acids (RAs) (Cash et al., 1997; Koyama et al., 1999; Lohnes et al., 1994; Pecorino et al., 1994, 1996; Scadding and Maden, 1994; Schilthuis et al., 1993; Tassava, 1992; Yamaguchi et al., 1998). There is also in vivo evidence that application of RA can influence the development of the antlers; injection of RA into the pedicle anlagen (Kierdorf and Kierdorf, 1998) has been shown to lead to an alteration of pedicle and antler shape. Exogenous RA also increased growth rate of the first antler and this was suggested to occur via an increase in the proliferation of periosteal cells (Kierdorf and Bartos, 1999). We have subsequently undertaken several studies which provide evidence to confirm that RA plays a potentially vital role in the regulation of cellular diVerentiation in regenerating antlers (Allen et al., 2002). HPLC analysis showed that significant amounts of all-trans-RA are present in all antler tissues, with the exception of chondroprogenitors, at levels similar to those found in regenerating amphibian limbs (Scadding and Maden, 1994). 9-cis-RA was also found in perichondrium, mineralized cartilage, and in bone, which suggests that it may have a particular function in regenerating bone since 9-cis-RA has not been detected in the developing limb. The RA synthesizing enzyme RALDH2 is present in skin, perichondrium, mesenchyme, perivascular tissue, and in osteoblasts. In contrast, the only site where RALDH2 is expressed in the chick limb is the perichondrium (Berggren et al., 2001). RAs act through nuclear receptors and retinoic acid receptors (RARs) and retinoid X receptors (RXRs). RAR, RAR, and RXR are all highly expressed in velvet antler skin, although we have not addressed what their role may be at this site. The expression pattern of RAR suggests that it also inhibits the diVerentiation of chondroprogenitors and maintains the perichondrium in a ‘‘prechondrogenic state,’’ consistent with its role in the developing limb (Weston et al., 2000). RAR is also expressed in perivascular tissue and its pattern of expression is so similar to that of type I collagen that it suggests a role for this receptor in regulating osteoblast diVerentiation. RAR is expressed in perivascular cells of the osteoblast (or osteoclast) lineage. RAR is not highly expressed in antler tissues, whereas in the developing limb its expression increases with chondrocyte diVerentiation (Dolle et al., 1989; Koyama et al., 1999; Mendelsohn et al., 1992). Only a

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small number of perivascular cells express RXR and RXR, whereas RXR has a widespread expression in hypertrophic chondrocytes. RA will induce dediVerentiation in micromass cultures of antler chondrocytes, detected by a decrease in GAG synthesis (Allen et al., 2002) and this eVect is dependent on RAR signaling. This is entirely consistent with RA’s eVect on chondrocytes in other species (Ballock et al., 1994; Takigawa et al., 1980). We have also studied PTHrP in some detail in the regenerating antler because, acting through the type I PTH/PTHrP receptor (PPR), it is a ‘‘master’’ regulator of chondrocyte diVerentiation in the developing limb (Kronenberg, 2003). This work has demonstrated how a single factor can have multiple roles in regenerating antler. PTHrP can be localized in epidermis, dermis, hair follicles, and sebaceous glands (Fig. 5A); this suggests that PTHrP may play an important role in controlling the growth of skin in regenerating appendages. These results also suggest that it may be important in mediating epithelial mesenchymal interactions in antlers, as it does in developing structures (Foley et al., 2001; Karperien et al., 1994). PTHrP also has an important role in endochondral and intramembranous ossification; it is present in perichondrium, mesenchyme, and chondroprogenitors. However, unlike the situation in the developing limb, PTHrP is not expressed in diVerentiated chondrocytes (Farquharson et al., 2001; Faucheux et al., 2004; Kartsogiannis et al., 1997) (Fig. 5). In vitro PTHRP inhibits antler chondrocyte diVerentiation and increases proliferation (Faucheux and Price, 1999), as it does in the developing limb. The localization of PTHrP and the PPR in antler osteoblasts indicates that it may also have a role regulating osteoblast diVerentiation. PTHrP has previously been shown to promote (Carpio et al., 2001; Motomura et al., 1996) osteoblast diVerentiation in vitro and adult mice heterozygous for PTHrP are osteopenic (Amizuka et al., 2000). The group of Peter Barling (Barling et al., 2004a) have recently cloned and sequenced PTHrP and the PPR from red deer and have studied their distribution in the developing antler by in situ hybridization and immunocytochemistry. They report the localization of PTHrP and PPR in epidermis, in dermis, and in skin appendages as well as in developing cartilage and bone. The expression of PPR mRNA in velvet antler skin is of particular interest since it has only recently been shown that the receptor is expressed in the skin of newborn rats (Errazahi et al., 2003) and changes in PPR mRNA expression have been shown to take place in mice during the hair cycle (Wang et al., 2002). The similar distribution of PTHrP and PPR in developing and regenerating antlers is further evidence that regeneration recapitulates development. Another molecule that may interact with PTHrP to control chondrogenesis is Indian hedgehog (IHH) that is immunolocalized in early hypertrophic

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chondrocytes in regenerating antlers (Faucheux et al., 2004). In the developing limb, IHH is expressed in prehypertrophic and hypertrophic chondrocytes and acts in a feedback loop with PTHrP to control chondrocyte diVerentiation (Chung et al., 2001; Kobayashi et al., 2002; Lanske et al., 1996; Vortkamp et al., 1996). In IHH null mice endochondral ossification is also impaired (St-Jacques et al., 1999) and IHH may have a similar role in antlers since it can be immunolocalized in osteoblasts in spongy bone in the midshaft of the antler, but is not found in osteoblasts at sites of intramembranous ossification. Lord et al. (2004) have also localized IHH mRNA in developing antler cartilage/bone by in situ hybridization. In a screen of antler extracts for factors that could potentially stimulate bone formation, a number of molecules were identified, including IGF-I and IGF II and fibroblast growth factors (Mundy, 2001). This study identified BMP-4 (Feng et al., 1995) and BMP-2 (Feng et al., 1997), two members of the transforming growth factor  superfamily that are known to be involved in embryonic skeletal development (HoVman and Gross, 2001). Barling et al. (2004b) have recently reported that BMP-2, BMP-4, BMP-14, and the BMP receptors BMPRI and ACTRII can be localized in the skin, cartilage, and bone of the developing red deer antlers. Experimental data from our laboratory (unpublished) suggests that BMP-2 regulates the diVerentiation of mesenchymal progenitor cells cultured from developing antlers; BMP-2 decreases proliferation and increased alkaline phosphatase activity (Fig. 15). BMP-2 also stimulates osteoblast diVerentiation of rat pluripotent progenitor cells (Hiraki et al., 1991). That BMPs should regulate antler regeneration is consistent with their role in regulating endochondral ossification and bone repair; both BMP-2 and BMP-4 will induce ectopic bone formation and BMP-2 will heal cortical bone defects by an endochondral process (Toriumi et al., 1991; Wang et al., 1990; Yasko et al., 1992). TGF1 and TGF2 have also been found at the mRNA level (Francis and Suttie, 1998) and TGF1 at the protein level (Faucheux et al., 2004) in antlers during their rapid growth phase. The role(s) of TGFs is currently unclear; however, the widespread distribution of TGF1 in mesenchymal cells, perivascular cells, and in osteoblasts suggests that it may have multiple functions, depending on its target cell type and its local concentration. For example, treatment of cells derived from mesenchyme with TGF1 will increase cellular diVerentiation (ALP activity) and this is associated with a decrease in cellular proliferation, but only at lower treatment doses (Fig. 16). TGF1 is also able to stimulate the production of PTHrP by antler cells in vitro (Faucheux et al., 2004), suggesting it may have a role similar to TGF2 in embryonic limb, where it is produced by the perichondrium and regulates chondrogenesis by inducing PTHrP (Alvarez et al., 2002). A molecule that may interact with TGF in antlers is dermatopontin that it was identified by Lord et al. (2001) in their cDNA library screen of regenerating

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Figure 15 The eVect of BMP-2 on proliferation and alkaline phosphatase activity of progenitor cells cultured from mesenchyme. (A) For proliferation assays, cells were cultured in 48 well plates and treated for 48 hours with BMP-2 in BGJb medium containing 2% fetal bovine serum. Cultures were pulsed for the final 12 hours with 3H thymidine and incorporated into the cells assayed. (B) For alkaline phosphatase assays, confluent monolayers of cells were treated for 72 hours with BMP-2 in BGJb medium containing 2% fetal bovine serum. Cell cultures were then lysed and the supernatants assayed for alkaline phosphatase using a colorimetric assay (Price et al., 1994). Results were corrected for protein content of the cultures and expressed as
mol product/
g protein/minute. BMP-2 inhibits growth but at higher doses promotes diVerentiation. *p < 0.05 compared with control, **p < 0.001 compared with control.

red deer antlers. Dermatopontin has been shown to interact with TGFs and enhance their activity (Okamoto et al., 1999) and its expression is decreased in tissues where fibrosis is occurring (Catherino et al., 2004; Kuroda et al., 1999), suggesting a role for this molecule in inhibiting fibrosis/scar formation in antlers. Another family of growth factors that may be important local regulators are Fibroblast Growth Factors (FGFs). Peter Barling (Barling et al., 2004b) presented data to show the localization of FGF2 and the FGF receptors FGFR1, FGFR2, and FGFR3 in skin, cartilage, and bone of developing red deer antlers. FGF-8 has also been detected in antler skin (Ashery, 1999). We have evidence that FGF-2 may be an important growth factor in antlers; addition of FGF-2 to cell cultures derived from both mesenchyme and, to a lesser extent, cartilage regions increased thymidine incorporation (unpublished observations) (Fig. 14). Consistent with the findings of Barling et al. (2004b), we have also detected FGFR3 receptor mRNA transcripts in cartilage of the regenerating antler tip, further evidence that FGFs may control chondrocyte growth and/or diVerentiation, as they do in the developing limb (Ornitz and Marie, 2002). An epidermal growth factor-like molecule has also been found in antler velvet tissue (Ko et al., 1986) and Barling et al. (2004b) have immunolocalized EGF and the EGF receptor in the developing antlers, although staining for the EGF receptor was far less

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35

Figure 16 The eVect of TGF-1 on proliferation (A) and alkaline phosphatase activity (B) of progenitor cells cultured from mesenchyme. The protocol was as described for Figs. 14 and 15. Like BMP-2, TGF-1 inhibits cell proliferation and inhibits diVerentiation. p < 0.05 compared with control, **p < 0.001 compared with control.

intense than that of its ligand. In addition to the growth of skin and bone in growing antlers, there is rapid growth of nerves since antlers are highly innervated structures. Garcia et al. (1997) reported widespread expression of Neurotrophin 3, a factor that controls nerve growth, particularly in the epidermis and dermis. Antler growth also depends on angiogenesis; these structures have to be highly vascularized because rapidly growing tissues have high metabolic demands. The expression of vascular endothelial growth factor (VEGF) and the VEGF receptor KDR have been mapped in the rapidly growing red deer antlers and extracts of antler will induce migration and growth of endothelial cells in culture (Clark et al., 2004). The same study showed that the angiogenic and chondrogenic

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factor pleiotrophin is also highly expressed in antler blood vessels and in chondrocytes (Clark et al., 2004). The studies reviewed here involved either developing antlers or regenerating antlers during their exponential growth phase. Surprisingly little is known about the pathways that control the very early stages of antler regeneration. Questions that need to be answered include: What is the trigger that induces cells in the pedicle to enter the cell cycle and to form a ‘‘blastema’’ or antler bud? What molecules prevent scar formation? What regulates branching? How do hormonal factors interact to regulate these pathways? How is infection prevented before epithelialization is complete? A technique that may prove valuable for establishing the function of signaling pathways in the early antler is biolistic particle delivery. This method has been used to transfect cells in the regenerating newt blastema (Pecorino et al., 1994, 1996) and we have used it to successfully transfect cells on the pedicle surface shortly after the previous set of antlers had been cast (Price and Allen, 2004). To date, molecules reported to be expressed in the early regenerating antler (<14 days of growth) include PTHrP, the PPR, TGF, RA, retinoic acid receptors, RALDH2, and TRAP (Faucheux et al., 2001, 2004; Price and Allen 2004). PTHrP and PPRs are both expressed in the regenerating wound epithelium, in ‘‘normal’’ epidermis, and dermis at the periphery of the antler bud, as well as in a large number of mesenchymal cells (Fig. 5). Since PTHrP is also expressed in periosteum, we have taken this to be evidence, albeit indirect, that these mesenchymal cells may be derived from pedicle periosteum. As mesenchymal cells in the blastema diVerentiate into chondrocytes, PTHrP synthesis is downregulated, which suggests that its role is to maintain mesenchymal cells in an undiVerentiated state. TGF1 is also expressed in the early antler and induces PTHrP synthesis in cultured cells from the blastema (Faucheux et al., 2004). TGF1 may also be involved in controlling fibrosis. As discussed in a previous section, we also have evidence that RA may regulate the early stages of antler regeneration since HPLC analysis has detected several RAs in antlers at fewer than 14 days of development, as well as RALDH2, RAR, RAR, and RXR (unpublished observations). To further explore our hypothesis that embryonic signaling pathways are recapitulated in regenerating antlers, we have also studied the localization of FGF-4 in the early antler (day 4) and found it to be immunolocalized in the regenerating epidermis and cells of the dermis (Fig. 17) (unpublished observations), which suggests that it may control epithelial–mesenchymal interactions. FGF-4 is also associated with vascular channels in the early antler (4-day) bud, although the significance of this observation is not clear. At a slightly later stage of growth (Day 14), FGF-4 is expressed in chondroprogenitors but not in fully diVerentiated chondrocytes, which suggests that it may also play a role in chondrogenesis.

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Figure 17 Immunolocalization of FGF 4 in the early regenerating antler. (A & B) Sections through a 4-day blastema. FGF-4 (brown stain) is localized in the (A) wound epithelium (we), cells of the dermis (d), and (B) in cells lining vascular channels (v). (C & D) Sections through a 4-day blastema. There is positive staining in chondroprogenitors but there is no FGF-4 synthesis by mature chondrocytes (c). Scale bars: 50
m. (See Color Insert.)

Clearly, our understanding of the complex signaling pathways that regulate antler development and regeneration remains fairly rudimentary. However, from the pieces of the jigsaw so far available, it would appear that the pathways that control the development of embryonic skeletal elements and the first set of antlers are conserved, even though antlers are appendages that develop after birth. There is also evidence that developmental signaling pathways are recapitulated during the repeated rounds of antler regeneration. Furthermore, it would appear that the molecules involved can have multiple functions, for example, retinoic acid controls the diVerentiation of progenitor cells, chondrocytes, osteoblasts, and osteoclasts. Identifying the components of these signaling pathways, and determining their function and their local and systemic interactions, will not only contribute to our understanding of basic regenerative processes but could, in the longer term, have applications in regenerative medicine, for example, for enhancing the repair of functionally competent cartilage, bone, skin, and nerves.

Acknowledgments This work has been supported by grants from The Wellcome Trust, The Medical Research Council, and the BBSRC. The authors thank Mr. Brian Nichols, Mr. James Mount, Mr. Thnaian AlThnaian, and Dr. Mariusz Muzylak for their contribution to this work and

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their collaborators Dr. Jeanine Danks, Professor Malcolm Maden, Professor Jeremy Brockes, Professor Mike Horton, and Dr. Monica Colitti.

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