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Nervous system injuries are a major cause of impairment in the human society. Up to now, clinical approaches have failed to adequately restore function following nervous system damage. The regenerative cycle of deer antlers may provide basic information on mechanisms underlying nervous system regeneration. The present contribution reviews the actual knowledge on the antler innervation and the factors responsible for its regeneration and fast growth. Growing antlers are profusely innervated by sensory fibers from the trigeminal nerve, which regenerate every year reaching elongation rates up to 2 cm a day. Antler nerves grow through the velvet in close association to blood vessels. This environment is rich in growth promoting molecules capable of inducing and guiding neurite outgrowth of rat sensory neurons in vitro. Conversely, endocrine regulation failed to show effects on neurite outgrowth in vitro, in spite of including hormones of known promoting effects on axon growth. Additional studies are needed to analyze unexplored factors promoting on growth in antlers such as electric potentials or mechanical stretch, as well as on the survival of antler innervating neurons.
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[Frontiers in Bioscience 17, 1389-1401, January 1, 2012]
Deer antler innervation and regeneration
Manuel Nieto-Diaz1, Daniel Wolfgang Pita-Thomas2, Teresa Munoz-Galdeano1, Cayetana Martinez-Maza3, Rosa
Navarro-Ruiz1, David Reigada1, Monica Yunta1, Marcos Javier Caballero-Lopez1, Manuel Nieto-Sampedro4, Rodrigo
1Molecular Neuroprotection Group, Hospital Nacional de Paraplejicos (SESCAM), Finca la Peraleda s/n, 45071 Toledo (Spain),
2Bascom Palmer Eye Institute, University of Miami Miller School of Medicine, Miami, FL 33136 (USA), 3Museo Nacional de
Ciencias Naturales (CSIC), C/ Jose Gutierrez Abascal 2, 28006 Madrid, Spain, 4Neural Plasticity Group, Instituto Cajal de
Neurobiologia (CSIC), Avda. Doctor Arce 37, 28002 Madrid, Spain
1. Abstract
2. Introduction
2.1. Nervous system injuries
2.2. Therapeutic potential of the antler’s nerve regeneration
3. Antler innervation
3.1. Antler nerve supply
3.2. Nerve fiber distribution in the antler
3.3. Types of antler nerve fibers
4. Antler regulators of axon regeneration
4.1. Paracrine regulation
4.2 Endocrine regulation and other promoting mechanisms
5. Summary and perspectives
6. Acknowledgments
7. References
Nervous system injuries are a major cause of
impairment in the human society. Up to now, clinical
approaches have failed to adequately restore function
following nervous system damage. The regenerative cycle
of deer antlers may provide basic information on
mechanisms underlying nervous system regeneration. The
present contribution reviews the actual knowledge on the
antler innervation and the factors responsible for its
regeneration and fast growth. Growing antlers are profusely
innervated by sensory fibers from the trigeminal nerve,
which regenerate every year reaching elongation rates up to
2 cm a day. Antler nerves grow through the velvet in close
association to blood vessels. This environment is rich in
growth promoting molecules capable of inducing and
guiding neurite outgrowth of rat sensory neurons in vitro.
Conversely, endocrine regulation failed to show effects on
neurite outgrowth in vitro, in spite of including hormones
of known promoting effects on axon growth. Additional
studies are needed to analyze unexplored factors promoting
on growth in antlers such as electric potentials or
mechanical stretch, as well as on the survival of antler
innervating neurons.
2.1. Nervous system injuries
Accidents causing spinal cord injury, traumatic
brain injury, stroke, or peripheral nerve injuries prevent the
correct functioning of the nervous system, causing total or
partial loss of sensory, motor, autonomic or cognitive
functions. Nervous system deficits are a major cause of
disability in developed countries. It has been estimated that
every year in the United States, 1.4 million people sustain a
traumatic brain injury (1), and 12,000 spinal cord injury (2)
while the incidence of nerve injury affects each year 13.9
per 100,000 persons in Sweden (3). In most cases, resulting
deficits are permanent. Less than 40 percent of the patients
suffering spinal cord injury (2) or 60 percent of those with
different peripheral nerve lesions (4) return to work after
injury. The social consequences are enormous since,
contrary to other pathologies like cancer or cardiovascular
diseases, neurological injuries frequently affect people
under 45 years of age, and the medical and social care is
required for decades.
Functional deficits following nervous system
damage are the direct consequence of the interruption of
Deer antler innervation
Figure 1. Illustration of the processes, regenerative capacities and functional outcomes that experience innervation following
peripheral nervous system (PNS) and central nervous system (CNS) injuries compared to those taking place during deer antler
regeneration. For each system, direct effects of damage are listed on top, below are the subsequent processes followed by the
regenerative capabilities of the system and the resulting functional outcome.
neural connections, largely due to the interruption of axons
(axotomy) and the death of neural cells (5) but also due to
inflammation, ischemia, and other processes that result in
extended cell death and disconnection (6-7, Figure 1).
Moreover, the reaction to damage of the nervous system may
also alter the neural circuitry and cause undesirable side effects
like neuropathic pain (5). Prognostic of nervous system injuries
depends on many parameters, including type, location or
extension of the lesion, age, etc. Functional deficits resulting
from severe injuries become permanent due to the limited
regeneration capacity of the nervous system. The high
differentiation degree of some neural cells, particularly neurons
or oligodendrocytes, prevents their proliferation and
replacement (8). The neurons surviving axotomy assume a
regenerative phenotype (9-11) but effective axon growth
depends on local environmental factors (see reviews in 12-13).
Mature central nervous system (CNS) environment is
inhibitory for axon growth (11,13), with few exceptions like
the olfactory or hippocampal tracts (see, for example, 14).
Adult CNS neurons also seem to present intrinsic properties
that reduce their regeneration capabilities (15). On the other
hand, peripheral nervous system (PNS) will regenerate within
the permissive growth environment of the Schwann cells (12),
although it progressively fails to sustain regenerative
response with time after injury (12). Thus, prolonged
denervations and axotomies result in poor functional
recoveries in most cases (16-17). Functional recovery
following injury is also made difficult by the high
specificity that neurons exhibit, which, in practice, means
that any new connections may not result in the recovery of
original circuits but the formation of new aberrant ones
2.2. Therapeutic potential of the antler’s nerve
Despite more than a century of neurological
research and surgical innovation, clinical approaches have
failed to adequately restore function following central or
even peripheral nervous system damage (7). The scientific
and clinical communities have realized that the
development of efficient therapeutic tools depends on our
understanding of the damaged nervous system and our
capacity to manipulate regeneration (19). Much work has
dealt with developmental mechanisms leading to the
formation of the nervous system assuming that their
reactivation may overcome the regrowth limitations after
injury. Less attention has been paid to systems that can
regenerate spontaneously even though these systems may
provide basic information on the mechanisms that rule
nervous system regeneration (20).
Deer antler innervation
The capability to regenerate large sections of the body plan
is typical of some invertebrates and urodele amphibians
(21-23). Among adult mammals, full organ regeneration is
exceptional, being restricted to deer antlers (24). Every
year, male deers shed (cast) their antlers and fulfill a
complete regeneration process that leads to the formation of
a new set of antlers. Antler regeneration cycle has been
recognized as a valuable model to study the mechanisms
underlying organ regeneration and rapid tissue growth in
mammals (25). The whole growth period takes place in about
3 months, reaching growth rates above 2 cm a day to build up
structures of more than 3 Kg (25), up to a 20 percent of the
whole skeleton weight (26). The growing antler is an extension
of the antler pedicle periostium (27) that proliferates and
differentiates into cartilage and bone tissue to form the bone
core of the new antlers. Growing antlers are enveloped in a
hair-covered skin known as velvet that presents several
peculiarities, including lack of sweat glands and arrector pili
muscles and the presence of abundant multilobullated
sebaceous glands (28). At the end of the summer, antlers
become calcified and velvet sheds, leaving the dead bony core
used in agonistic encounters during the rut season (29). Every
year, antler innervation regenerates to provide the antler with
nerve supply (30-33). This growth supposes an extraordinary
enlargement of the peripheral field that likely arises as a local
extension of the nerve fibers that supply the forehead and the
pedicle. Once antlers stop growing and become mineralized,
nerve fibers at the velvet die back to the pedicle where they
apparently remain encapsulated (34). By the end of the winter,
when dry antlers are cast and new ones begin to grow, nerves
reenter into a regenerative state to supply the velvet antler.
Deer antlers are a very interesting source of
information on the mechanisms underlying the nerve
regeneration and functional recovery following injury
(20,23,35). This spontaneous regeneration model is
particularly interesting for therapeutic studies because it
occurs in an adult mammal and involves cells, mechanisms
and/or biochemical pathways which are more likely to be
similar to those in humans than other regenerating models
as the non-mammalian vertebrates (newts or fishes) or
invertebrates. Moreover, antler regeneration takes place in
adult individuals and affects adult neural cells like those
typically involved in nerve injuries and different from those
of embryos with different growth capabilities (see for
example, 15, 36). The extraordinary extension and growth
rate that the nerve fibers achieve during antler regeneration
is particularly outstanding from both the clinical and
biological points of view (Figure 1). Gray and colleagues
(32) demonstrated that nerve supply grows together with all
antler tissues to cover all antler (see also 33,37,38),
reaching elongation rates over 2 to 3 cm a day in the largest
species like the moose (Alces alces; 24) or the wapiti
(Cervus canadenisis; 39). From the biological point of
view, it is particularly remarkable the transport rate and the
amounts of cell material needed to enlarge axons more than
one meter at rates above 2 cm per day, especially if we
consider that maximum axon growth by growth-cone
extension only reaches 1 mm per day (40-41). In fact, such
an extremely fast growth is not consistent with current
understanding of the transport of essential structural
elements such as neurofilament proteins, for which the
average transport speed is limited to 1 mm per day (42-45).
The rapid and sustained nerve growth observed in the
antlers indicates that the physiological capacity of axons to
expand rapidly and continuously is not limited by protein
synthesis, transport rates, or the availability of structural
constituents (43-44,46-48). The clinical consequences of
understanding this phenomenon would be enormous,
considering that in many injuries, growing axons have to
reach far away targets, up to one meter in the case of some
human nerve fibers. Reconnecting these targets would take
years for axons growing at elongation rates around 1 mm
per day, resulting in functional recovery failure (17).
However, at the rate observed in deer antlers, it would take
a few months of growth.
3.1. Antler nerve supply
The first description on the anatomy of the deer
antler innervation dates back to the 19th century (49). Later
studies on Virginia deer (Odocoileus virginianus; 30), red
deer (Cervus elaphus, 50), wapiti (cervus canadensis, 51),
and fallow deer (Dama dama, 51) showed that the nerves
supplying the antlers come from the supraoptic
(infratrochlear) branch of the ophtalmic division and the
zygomaticotemporal branch of the maxillary division, both
from the trigeminal (5th cranial) nerve (Figure 2). The
supraoptic branch is a single bundle that emerges from the
skull beneath the upper edge of the orbit (1.5-2 cm from the
medial canthus of the eye in red deer according to Adams,
50) and courses medially over the dorsal rim, near the
medial angle of the eye (51). It then courses caudally
through the orbicularis oculi muscle and sends several
nerves toward the base of the antlers where 6 or more small
branches in Virginia deer (30) or a full web in wapiti (51)
are given off to the anterior and medial surfaces of the
pedicle and the antler. The zygomaticotemporal branch of
the maxillary nerve emerges onto the scalp as a large nerve
near the zygomatic arch, at the caudal margin of the
zygomatic process of the frontal bone, and divides
immediately, producing a number of branches coursing
toward the antler base and the ear. The branches that supply
the antler pass caudodorsally through the retrorbital
(periorbital) fat and beneath the frontalis muscle. Midway
from the orbit to the pedicle they branch into several nerves
(6 or more in Virginia deer according to 30), which
disperse on the lateral and posterior surfaces of the antler
pedicle and then onto the antler. Both trigeminal nerve
supplies follow very closely the respective distribution of
the lateral and medial coronary branches of the superficial
temporal artery (the last branch of the external carotid) that
provides the antler blood supply (30,51). Besides the
trigeminal innervation, fibers from the zygomatic branch of
the auriculopalpebral nerve (a branch of the facial or 7th
cranial nerve) have been observed to reach the medial
surface of the pedicle (50-51) although there is no evidence
of its extension into the antler. According to Woodbory and
Haigh (51), no other nerves could be traced to the pedicle.
3.2. Nerve fiber distribution in the antler
Several authors have studied the location of the
nerve supply in the growing antler (30-32,34,37-38).
Deer antler innervation
Figure 2. Illustration detailing the antler nerve supply.
Growing antlers are innervated by the zygomaticotemporal
and supraoptic branches of the trigeminal nerve (solid
black lines) that reach the antler accompanying the
superficial temporal artery (in gray dashed lines).
Growing antlers consist on tips of densely packed
mesenchymal cells which differentiate proximally into
cartilage and bone tissue (Figure 3). Overlaying cartilage
and bone, there is a layer of perichondrium/periostium that
ultimately shows continuity with the mesenchyme
primordia. The entire antler is covered by velvet, a
modified skin with abundant hair follicles and sebaceous
glands but without arrector pili muscles and sweat glands
(28). Histologically, velvet consists on a thick epidermis
without invaginations that covers a thick dermis. Velvet
dermis can be subdivided into an outer dermis, in contact
with the epidermis, that contains numerous hair follicles
and sebaceous glands, and an inner dermis, located
beneath, that does not present any visible structure. Beneath
the dermis appears a layer of highly-vascularized loose
connective tissue -termed vascular layer by several authors
(30-31,33-34,52)- that separates the tegument from the
avascular mesenchyme. Wislocki and Singer (30) used
Bodian's protargol method to show profuse nerve fibers
occurring in small bundles at the vascular layer in Virginia
deer. Similar results were obtained by Vacek (31) for red
deer, fallow deer and roe deer (Capreolus capreolus), and
Li et al. (34) for red deer, specifying that nerves passed
through and above the vascular stratum of the velvet from
where they projected to more superficial layers.
Immunohistochemical analyses in red deers by Gray and
colleagues (30) showed that innervation concentrate on
deep connective layers of the velvet, which anatomically
correspond to the vascular layer. They also observed nerve
fibers in the dermis, the epidermis the
periostium/perichondrium and even in cartilage tissues
within the central core of the antler. In 2007, Li et al. (37)
also observed nerve fibers immunoreactive to the 200 Kda
neurofilament (NF200), traveling through the mesenchyme
primordium. In order to examine antler nerve distribution
in detail, we used antibodies against different
neurofilaments to stain antler sections confirming that, in
agreement with previous studies, most fibers are located
deep in the velvet, mainly between the dermis and the
vascular layer (33,38,53, Figure 3). Most fibers appear
isolated at the antler tip while they usually form bundles at
the base. Nerve branches project from the vascular layer
towards more superficial layers of the velvet but, contrary
to Li et al. (37) and Gray et al. (32), we have not observed
them in the mesenchyme or derived tissues (33,38,53).
Antler nerve fibers are both myelinated and not-myelinated
(30-31) as confirmed by the presence of glial fibrillary
acidic protein (GFAP) immunoreactive Schwann cells
accompanying or enseathing axons in the velvet tissues
from the base to the tip of the antler (Figure 3).
3.3. Types of antler nerve fibers
Antler nerve fibers show specific
immunoreactivity to markers of distinct sensory neuron
populations (32-33,37-38,53). Primary sensory fibers can
be characterized by their neurochemical signature, in
particular their immunoreactivity to CGRP (calcitonin gene
related peptide), Substance P, and NF200 peptides and
isolectin B4 (IB4) from Griffonia simplicifolia (54-56).
According to this classification, myelinated Abeta fibers
are only immunoreactive to NF200, the thin myelinated
Adelta fibers are immunoreactive to both NF200 and
CGRP; while the un-myelinated peptidergic C fibers are
immunoreactive to the nociceptive peptides CGRP and
substance P. Finally, non-peptidergic C fibers are marked by
IB4. Gray and colleagues (32) observed peptidergic C fibers
immunoreactive to substance P and CGRP surrounding blood
vessels and as free fibers. Li and colleagues (37) showed the
presence of nerve fibers immunoreactive to the NF200 in the
antler. Recently, we have combined immunohistochemistry for
NF200 and CGRP to carry out a detailed analysis of the antler
innervation, from the trigeminal or semilunar ganglion to the
antler tip (38). Stained sections of the trigeminal nerve showed
the presence of bundles of NF200 positive, CGRP negative
fibers, others with fibers positive for both markers and some
fibers exclusively marked against CGRP (Figure 4). The same
combinations of markers were observed at the vascular layer of
the antler, mostly as bundles in the base and as free fibers at
the tip, although bundles were occasionally seen at the tip and
free fibers at the base (38,53). However, although we could
establish the presence of fibrous structures IB4 positive, the
lack of colocalization with axon markers and the fact that IB4
also stains vessels and even Schwann cells precludes the
unequivocal confirmation of the presence of these fibers in the
antler. Antler nerve fibers seem to end freely in the tissue,
lacking specialized sensory nerve receptors (30-31). Free fiber
endings are characteristic of mechanoreceptors of Abeta fibers,
and the majority of Adelta and C fibers. Abeta fibers convey
touch information from skin receptors to mechanical
stimuli of low intensity (cutaneous mechanoreceptors),
while Adelta fibers that function as receptors for pressure,
touch and cold temperature as well as convey fast pain
information from acute noxious stimuli, and peptidergic C
fibers, which are warmth and slow pain receptors (also
responsible for itch sensation). In the antler, both Abeta and
Adelta fibers would provide precise localization of the
stimulus or damage location, being responsible for the
withdrawal reflex and the extreme sensitivity to touch
stimuli of the growing antlers (37). On the contrary, C
fibers would cause slow “burning” pain and release CGRP
and Substance P which are involved in inflammation and
wound healing.
Besides sensory innervation, antlers do not seem
to present other nerve components. Motor fibers have not
been observed in agreement with the lack of muscles in the
Deer antler innervation
Figure 3. Anatomy of the antler innervation. A) Histology of the antler tip (hematoxilin eosin) specifying its constituting tissues.
B) Detail of the antler tip (hematoxilin eosin). E=epidermis, OD=outer dermis, ID=inner dermis, VL=vascular layer,
M=mesenchyme, PC=precartilague. C) Corresponding serial section showing nerve fibers (green) immunostained with antibody
against Neurofilament and cell nuclei stained with Hoechst (blue). Axons are located over the vascular layer (arrow), but also at
the dermis (star) or within the vascular layer (asterisk). D) Schwann cells (GFAP, red) enseathing an axon (Neurofilament
panclonal, green) at the antler tip.
antler. Sympathetic innervation of the blood vessels was
suggested by Vacek (31), who described nerve fibers in
the adventitia and the media of the antler arteries.
However, neither previous anatomical studies by
Wislocki and Singer (30) nor later studies by Rayner
and Ewen (57) and Gray et al. (32) have confirmed the
presence adrenergic nerve fibers in the blood vessels of
any part of the antler, although they were evident at the
pedicle (57). In fact, description, antler arteries seem to
be constructed to close themselves by constriction in the
event of being severed, being independent from innervation
4.1. Paracrine regulation
The factors underlying nerve fiber growth in the
antler remain largely unknown. However, a paracrine
regulation can be expected, due to the presence in the antler
Deer antler innervation
Table 1. Axon growth promoters identified in the growing
Trophic factors
Nerve Growth Factor
Brain Derived Growth Factor
Epitelial growth factors
Epidermal Growth Factor
Fibroblast growth factors
basic Fibroblast Growth Factor
Insulin-like growth factors
Insulin-like Growth Factor-1
Insulin-like Growth Factor-2
Transforming growth factor family
Bone Morphogenetic Protein 2
Bone Morphogenetic Protein 3B
Bone Morphogenetic Protein 4
Transforming Growth Factor beta
Vascular endotelial growth factor
Neurite growth promoting factors
Pigment Epitelium Derived Factor
Other factors
Glucose-6-Phosphate Isomerase
Retinoic acid
Extracellular matrix molecules
Collagen I
Heparan sulfate
Cell adhesion molecules
Neuronal Cell Adhesion Molecule
For references see 57
of Schwann cells and endothelial cells which constitute a
normal source of neural growth promoters. In agreement,
antlers express nerve growth factor (NGF, 37) and
neurotrophin 3 (NT3, 56), two well-known axon growth
factors of the neurotrophin family. Recently, we combined
different molecular techniques to determine the gene
expression of axon growth promoters in the antler velvet
(59). Microarray analysis allowed us to hypothesize the
expression or change in expression of 90 promoters or
regulators of the axonal growth. 15 of them were sequenced
and analyzed by semiquantitative RT-PCR, establishing the
expression in the antler velvet of brain derived neurotrophic
factor (BDNF), glucose phosphate isomerase (GPI),
meteorin (MTRN), midkine (MDK), and neuronal cell
adhesion molecule (NRCAM), not previously observed in
deer. Combining these data with previous analyses, a list of
more than 20 axon growth promoters can be obtained
drawing a picture of the growth promoting environment of
the antler innervation during its annual regeneration. The
list (Table 1) comprises several neurotrophins and growth
factors, such as basic fibroblast growth factor (FGFb),
epidermal growth factor (EGF), pleiotrophin, or pigment
epithelium derived factor (PEDF), morphogens of the
transforming growth factor (TGFbeta) family, members of
the insulin-like growth factor (IGF) family, together with
retinoic acid, and several substrate molecules like collagen,
laminin and heparan sulphate. All these molecules have
demonstrated direct or indirect axon growth promoting
properties for different neuronal types, including sensory
neurons like those innervating the antler, either in culture or
in vivo. Most of them also promote axon regeneration
following nervous system damage (60-67) or have been
identified in models of epimorphic regeneration, like in the
newt limbs, or in the fish fins. Classic among them are
retinoic acid (68-69), bone morphogenetic proteins (BMPs,
70-71) or FGFb (69-70,72-73), but also collagen (70,74),
heparan sulphates (75), laminin (76), IGF (77) or TGFbeta
(78). In most cases, these molecules are necessary for the
organ regeneration to be completed or even initiated, but
little is known on their roles in the axon growth during the
epimorphic regeneration process. Table 1 also shows that
most factors are present or expressed in the velvet skin,
where nerves are located, except for the BMPs. In fact,
immunohistochemical and in situ hybridization studies
indicate the presence or expression of most promoters in
the inner vascular layer of the velvet dermis where most
nerve fibers are observed. This is the case of NGF (37),
pleiotrophin (79), EGF (52), FGFb (80), and vascular
endothelial growth factor (VEGF, 80). Such spatial co-
localization indicates a direct exposure of the axons to
these molecules and would confirm a relevant role for them
in antler nerve growth, either promoting axon regrowth or
meeting the trophic requirements of the axons. However,
comparisons with published information on individual
molecules or from human gene expression profiles stored in
the GeneNote database (81) reveals that all identified
promoters are also expressed in normal or wounded skin.
Moreover, comparison of the gene expression between the
antler velvet and the skin overlaying the antler pedicle or
the frontal bone confirms this similarity (59). The only
exception is Midkine, which is significantly overexpressed
in the velvet respect to frontal skin samples, though not
respect to pedicle skin. Somehow, Midkine expression
profile fits with what can be expected from a molecule
involved in the process of rapid axonal growth in the antler.
The capabilities of the antler molecules to
promote neurite growth were first evaluated by Huo and
colleagues (82), who observed neurite outgrowth from
PC12 cells cultured with growing antler tissue extract. We
further analyzed the axon growth promoting properties of
the different tissues of the growing antler (33). The analysis
of the effects of antler extracellular matrix and cell
adhesion molecules on neurite outgrowth from dorsal root
ganglia (DRG) demonstrated that the antler vascular layer
has the capability to guide but not to promote axon growth.
These results suggest the action of axonal guidance
systems, in agreement with a strong orientation of the
collagen fibers in deep velvet layers (83) or the
predominant expression of NGF in the vascular layer of the
dermis (37). On the other hand, soluble molecules secreted
by the antler tip velvet promote a significant increase in
neuritogenesis and neurite outgrowth in DRG neurons from
rat embryos (33), trigeminal neurons from adult rats (53)
and PC12 cells (in preparation), as illustrated by Figure 5.
Biochemical treatments using enzymatic digestion, heat
denaturation, and size filtering treatments proved that the
neurite outgrowth promoters are most likely proteins.
Experiments using blocking antibodies (33,53) showed that
NGF blockage caused a significant reduction of neurite
Deer antler innervation
Figure 4. Fiber types of the antler innervation according to their neurochemical signature. A to D: control staining of the
ophthalmic nerve at the exit of the trigeminal ganglion. A and B show fibers immunostained against NF200 (green) and CGRP
(red) with all possible combinations of the markers (NF200+/CGRP-, NF200+/CGRP+, and NF200-/CGRP+). C and D show the
staining of isolectin B4 (green) and the axon marker panclonal neurofilament. No NFp+ fibers appear marked by IB4. Some IB4+
cells surrounding NFp+ axons (arrows in D) could correspond to Schwann cells. E and F show different combinations of CGRP
(red) and NF200 (green) immunoreactivity in nerve fibers at the base (E) and the tip (F) of the antler. All possible combinations
are onserved, corresponding to Abeta (NF200+/CGRP-), Adelta (NF200+/CGRP+) and peptidergic C fibers (NF200-/CGRP+).
Nerve bunches are observed at both the base (E) and the tip (F) but they are clearly larger at the base.
growth, proving a clear contribution of NGF to the effect of
the velvet in agreement with its expression in vivo (37).
However, velvet medium still kept significant promoting
effects, indicating that other proteins also contributed to
these growth properties.
4.2. Endocrine regulation and other promoting
Antler regeneration cycle is strongly regulated by
hormones (84). Some hormones with axon growth
promoting properties like IGF-1 (85-86) or triiodothyronine
(T3; 87) show high blood levels during the antler growth
period (88-90). However, in vitro analyses indicate that the
neurite growth promoting properties of serum obtained
during antler maximal growth do not differ from those of
serum from other antler cycle stages (33). It is intriguing
why strong differences in the serum concentration of IGF-1
during the antler regeneration cycle (84) are not reflected
by differences in neurite growth in these assays. It is
possible that the increased number of Schwann cells caused
by the deer sera would mask any direct effect of serum
molecules on neurite outgrowth. However, that would point
to a very modest promoting effect of the endocrine
regulators and therefore, they could not be leading
participants of the high growth rates registered in antler
axons. Nevertheless, it is also possible that molecules with
higher levels in serum during antler regeneration could
influence indirectly axon growth, for example, by acting on
the expression of neurotrophic factors or even acting in
synergy with other factors.
Deer antler innervation
Figure 5. Neurite outgrowth induced by soluble molecules secreted by the antler velvet. Illustrative images showing the
differences in neurite outgrowth of adult rat trigeminal neurons (upper row), rat embryo DRG neurons (middle row), and PC12
line cells (lower row) cultured with basal medium (negative control, left column), medium supplemented with NGF (center
column), and conditioned (supplemented) by antler velvet tissue (right column).
In addition to paracrine and endocrine regulation,
other less-classical factors may contribute to cause the very
fast axonal growth observed in the deer antlers, including
mechanical stretch or electric fields. Growing antlers present
negative electrical potentials between their tip and base, which
vary in marked correlation with the growth rate of the antler
(91). Similar electric potentials have been observed to promote
and orientate axonal growth during development and
regeneration (see 92 for review). The influence of electric
fields on antler nerve growth was already proposed by Bubenik
(23,93), who also proposed that electric stimulation of the
growth of the nerve branches innervating the antler would also
induce a general growth of the antler, both in size and
complexity. Parallel evidences also support the contribution of
mechanical stretch to the antler nerve growth. Antler velvet
experience strong mechanical stretch due to the fast growth
of the underlying mesenchyme (83). According to Li and
Suttie (28), this stretch may be responsible for the growth
and properties of the antler velvet. As shown in different in
vitro studies, such tensions can induce higher axon growth
rates than any other promoter (94-95). In fact, mechanical
stretch seems to be responsible for the axon elongation
during postnatal development, when axons have already
connected to their targets but still have to grow (95).
We have just begun to analyze the axon
regeneration in the deer antler (33,37,53,58,59,95). The
analyses have shown that the antler velvet provides the
growing axons with a environment rich in promoting
factors, with NGF having a basic role (33,37,82,96),
although other paracrine regulators should also contribute
(33). In this respect, our group is using proteomic
techniques to isolate and identify axon growth promoters
secreted by the velvet. Preliminary results have identified
more than 80 secreted or membrane proteins with
documented relationships to axon growth, including cell
adhesion, extracellular matrix and soluble molecules.
However, it is also necessary to evaluate the effect of
unexplored factors like mechanical stretch, or electric
fields, which may also contribute or even determine the
growing characteristics of the antler innervation.
Despite the potential of these approaches,
understanding the nerve regeneration process and its
regulators would greatly benefit from the direct study of the
antler neurons. The study of the antler’s transcriptome,
particularly the analysis of the trigeminal neurons
Deer antler innervation
innervating the antler, would be also very helpful to
identify and characterize the processes taking place during
the antler regeneration. This approach, followed by other
cell and molecular analyses, would allow the
characterization of these neurons and the pathways
activated during the antler regeneration. The recent
sequencing and annotation of the red deer genome by
researchers from New Zealand and the development of the
High Throughput Sequencing methodologies capable of
sequencing hundreds of millions RNA molecules have opened
the opportunity to carry out these analyses. The resulting
dataset would help many research groups working on this
model as well as researchers from related fields to access
detailed information on all tissues involved in the process at
different times.
Besides nerve regeneration, functional repair of
injured nervous system also depends on protecting nerve cells
from cell death. Apoptotic cell death following CNS injury is
responsible for a significant loss of functional capabilities (97-
98). Even in the peripheral nervous system, 20 to 50 percent
of sensory neurons die through apoptosis or similar
processes during the weeks following injury (6,7,99-100),
and some specific populations almost completely disappear
(100). The total number of dying cells varies depending on
factors like the age, the location of the injury or its extent
(7). Apoptosis of axotomized neurons may be triggered by
antidromic electrical activity or neurotoxic inflammatory
agents, or most likely by the subsequent loss of target-
derived neurotrophic support (101). Conversely, the
sensory neurons supplying the growing antler do not seem
to experience relevant cell death processes even 8 months
after axotomy. Although direct evidence of neuron survival
or death after antler mineralization is still lacking, the
presence of encapsulated axon ends at the pedicle (34)
suggests that neurons innervating the antler survive
axotomy and regenerate as collaterals from these
encapsulated fibers to supply the new antler one year later.
The present contribution was supported by
funding from the Health Department of the Castilla-La
Mancha government (projects ICS-06025 and PI2008-38).
We are deeply in debt to Andres García, Tomas Landete,
Laureano Gallego from IREC and UCLM, and Javier
Martin from VenissonDeer for providing the deer samples
used in our studies. We also thank an anonimous reviewer
for his/her helful comments on the manuscript.
1. Centers for Disease Control (CDC). Traumatic Brain
Injury in the United States: A Report to Congress. (2001)
2. National Spinal Cord Injury Statistical Center (NSCISC).
Facts and figures at a glance-February 2010. (2010).
3. Maria Asplunda, Mats Nilssona, Anders Jacobsson, Hans
von Holst: Incidence of Traumatic Peripheral Nerve
Injuries and Amputations in Sweden between 1998 and
2006. Neuroepidemiol 32, 217-228 (2009)
4. Coen NP Bruyns, Jean-Bart Jaquet, Ton AR Schreuders,
Sandra Kalmijn, Paul DL Kuypers, Steven ER Hovius:
Predictors for return to work in patients with median and
ulnar nerve injuries. J Hand Surg Am 28 (1), 28-34 (2003)
5. Xavier Navarro: Neural plasticity after nerve injury and
regeneration. Int J Neurobiol 87, 483-505 (2009)
6. Mike J Groves, T Christopherson, Bruno Giometto,
Francesco Scaravilli: Axotomy-induced apoptosis in adult
rat primary sensory neurons. J Neurocytol 26, 615-624
7. Andrew M Hart, Giorgio Terenghi, Mikael Wiberg:
Neuronal death after peripheral nerve injury and
experimental strategies for neuroprotection. Neurol Res 30
(10), 999-1011 (2008)
8. Ferdinando Rossi, Elena Cattaneo: Neural stem cell
therapy for neurological diseases: dreams and reality. Nat
Rev Neurosci 3, 401-409 (2002)
9. Frank Bosse, Kerstin Hasenpusch-Theil, Patrick Kury,
Hans Werner Müller: Gene expression profiling reveals
that peripheral nerve regeneration is a consequence of
both novel injury-dependent and reactivated
developmental processes. J Neurochem 96 (5), 1441-
1457 (2006)
10. Yi Yang, Yuanyuen Xie, Hong Chai, Ming Fan,
Shuhong Liu, Hong Liu, Iain Bruce and Wutian Wu:
Microarray analysis of gene expression patterns in adult
spinal motoneurons after different types of axonal
injuries. Bran Res 1075, 1-12 (2006)
11. Santiago Ramon y Cajal: Degeneration and
regeneration of the nervous system. Eds: DeFelipe J,
Jones, EG, Hafner Press, NY (1928)
12. Keith Fenrich, Tessa Gordon: Axonal regeneration
in the peripheral and central nervous systems – current
issues and advances. Can J Neurol Sci 31, 142-156
13. Michael E Selzer: Promotion of axonal regeneration
in the injured CNS. Lancet Neurol 2, 157-166 (2003)
14. Pasquale PC Graziadei, Ariella G Monti Graziadei:
Regeneration in the olfactory system of vertebrates. Am
J Otolaryngol. 4 (4), 228-33 (1983)
15. Jeffrey L Goldberg, Matthew P Klassen, Ying Hua,
Ben A Barres: Amacrine-signaled loss of intrinsic axon
growth ability by retinal ganglion cells. Science 296,
1860-1864 (2002)
16. Susan Y Fu, Tessa Gordon: Contributing factors to
poor functional recovery after delayed nerve repair:
prolonged axotomy. J Neurosci 15, 3876-3885 (1995)
Deer antler innervation
17. Susan Y Fu, Tessa Gordon: Contributing factors to poor
functional recovery after delayed nerve repair: prolonged
denervation. J Neurosci 15, 3886-3895 (1995)
18. Antoni Valero-Cabre, Konstantin Tsironis, Emmanouil
Skouras, Xavier Navarro, Wolfram F. Neiss: Peripheral
and spinal motor reorganization after nerve injury and
repair. J Neurotrauma 21, 95-108 (2004)
19. Simon PJ Kay, Mikael Wiberg, Daniel JA Thornton:
Nerves are living structures whose injury requires urgent
repair. J Plast Reconstr Aesthet Surg 63, 139-1940 (2010)
20. Patrizia Ferretti, Fang Zhang, Paul O'Neill: Changes in
spinal cord regenerative ability through phylogenies and
development: lessons to be learnt. Dev Dyn 226, 245-256
21. Richard J Goss: Principles of regeneration. Academic
Press, NY (1969)
22. Miranda Mladinic, Kenneth J Muller, John G Nicholls:
Central nervous system regeneration: from leech to
opossum. J Physiol 587, 2775-2782 (2009)
23. Ioannis V Yannas: Tissue and Organ Regeneration in
Adults. New York, Springer (2001)
24. Richard J Goss: Problems of antlerogenesis. Clin
Orthop Relat Res 69, 227-238 (1970)
25. Joanna S Price, Steve Allen, Corrine Faucheux,
Thnaian Althnaian, James G Mount: Deer antlers: a
zoological curiosity or the key to understand organ
regeneration in mammals? J Anat 207, 603-618 (2005)
26. Tomas Landete-Castillejos, Jose A Estevez, Fernando
Ceacero, Andres J Garcia, Laureano Gallego: A review of
factors affecting antler composition and mechanics. Front
Biosci (this volume)
27. Chunyi Li, James M Suttie: Deer antlerogenic
periostium: a piece of postnatally retained embryonic
tissue? Anat Embryol 204, 375-388 (2001)
28. Chunyi Li, James M Suttie: Histological studies of
pedicle skin formation and its transformation to antler
velvet in red deer (Cervus elaphus). Anat Rec 260, 62-71
29. John D Currey, Tomas Landete-Castillejos, Jose A
Estevez, Fernando Ceacero, Augusto Olguin, Andres J
Garcia, Laureano Gallego. The mechanical properties of
red deer antler bone when used in fighting. J Exp Biol 212,
3895-3993 (2009)
30. George B. Wislocki, Marcus Singer: The occurrence
and function of nerves in the growing antlers of deer. J
Comp Neurol 85, 1-19 (1946)
31. Zdenek Vacek: Innervace lyci rostoucia parohu u
cervidu. Cslkgl Morf 3, 249–264 (1955)
32. Collin Gray, Mika Hukkanen, Yrjo T Konttinen,
Giorgio Terenghi, Timothy R Arnett, Sheila J Jones,
Geoffrey Burnstock, Julia M Polak: Rapid neural growth:
calcitonin gene-related peptide and substance P-containing
nerves attain exceptional growth rates in regenerating deer
antler. Neurosci 50, 953-963 (1992)
33. Daniel Wolfgang Pita-Thomas, Manuel Nieto-
Sampedro, Rodrigo M Maza, Manuel Nieto-Diaz:
Factors promoting neurite outgrowth during deer antler
regeneration. J Neurosci Res 88, 3034-3047 (2010)
34. Chunyi Li, James M Suttie, Dawn E Clark:
Histological examination of antler regeneration in red
deer (Cervus elaphus). Anat Rec A Discov Mol Cell Evol
Biol 282, 163-174 (2005)
35. Brian W Payton. History of medicinal leeching and
early medical references. In: Neurobiology of the Leech.
Eds: Muller K, Nicholls J, Stent G, Cold Spring Harbor,
NY: Cold Spring Harbor Laboratory, pp. 27–34. (1981)
36. Phillip L Lamoureux, Matthew R O'Toole, Steven R
Heidemann, Kyle E Mille: Slowing of axonal
regeneration is correlated with increased axonal
viscosity during aging. BMC Neurosci 11, 140 (2010)
37. Chunyi Li, Jo-Ann L Stanton, Tracy M Robertson,
James M Suttie, Philip W Sheard, A John Harris, Dawn
E Clark: Nerve Growth Factor mRNA Expression in the
Regenerating Antler Tip of Red Deer (Cervus elaphus).
PLoS ONE 2 (1), e148 (2007)
38. Daniel Wolfgang Pita-Thomas, Rodrigo Martinez-
Maza, Monica Yunta, David Reigada, Rosa Navarro-
Ruiz, Marcos Javier Lopez Rodriguez, Manuel Nieto
Sampedro, Manuel Nieto-Diaz: Caracterización
inmunohistoquímica de la inervación de las astas de
ciervo en crecimiento. Resumenes del Primer Encuentro
Internacional Virtual de Educación e Investigación en
Ciencias Morfológicas, Argentina (2009)
39. Enrique Gaspar-Lopez, Tomas Landete-Castillejos,
Laureano Gallego, Andres Garcia: Antler growth rate in
yearling iberian red deer (Cervus elaphus hispanicus).
Eur J Wildl Res 54, 753-755 (2008)
40. Phillip Lamoureux, Jing Zheng, Robert E. Buxbaum,
Steven R. Heidemann: A cytomechanical investigation
of neurite growth on different culture surfaces. J Cell
Biol 118, 655-661 (1992)
41. Hellen M Buettner, Randall N Pittman, Jonathan K
Ivins: A model of neurite extension across regions of
nonpermissive substrate: simulations based on
experimental measurement of growth cone motility and
filopodial dynamics. Dev Biol 163, 407-422 (1994)
42. Ralph A Nixon: The slow axonal transport of
cytoskeletal proteins. Curr Opin Cell Biol 10, 87-92
Deer antler innervation
43. Anthony Brown: Slow axonal transport: stop and go
traffic in the axon. Nat Rev Mol Cell Biol 1, 153-156
44. Subhojit Roy, Pilar Coffee, George Smith, Ronald KH
Liem, Scott T Brady, Mark M Black: Neurofilaments are
transported rapidly but intermittently in axons: implications
for slow axonal transport. J Neurosci 20, 6849-6861 (2000)
45. Jagesh V Shah, Don W Cleveland: Slow axonal
transport: fast motors in the slow lane. Curr Opin Cell Biol
14, 58-62 (2002)
46. Phillip Lamoureux, Robert E Buxbaum, Steven R
Heidemann: Axonal outgrowth of cultured neurons is not
limited by growth cone competition. J Cell Sci 111, 3245-
3252 (1998)
47. Jaime Alvarez, Antonio Giuditta, Edward Koenig: Protein
synthesis in axons and terminals: significance for maintenance,
plasticity and regulation of phenotype with a critique of slow
transport theory. Prog Neurobiol 62, 1-62 (2000)
48. Perry A Brittis, Qiang Lu, John G Flanagan: Axonal
protein synthesis provides a mechanism for localized
regulation at an intermediate target. Cell 110, 223-235 (2002)
49. Arnold Adolph Berthold: Uber das Wachstum, den abfall
und die wiedererzogung der Hirchgeweihe. Beitrag z Anat
Zool u Physiol 5, 39-96 (1831)
50. John Lewis Adams: Innervation and blood supply of the
antler pedicle of the red deer. New Zealand Vet J 27, 200-201
51. Murray R Woodbury, Jerry C Haigh: Innervation and
anesthesia of the antler pedicle in wapiti and fallow deer. Can
Vet J 37, 486-489 (1996)
52. Peter M Barling, Angela KW Lai, Louise FB Nicholson:
Distribution of EGF and its receptor in growing red deer antler.
Cell Biol Int 29, 229-36 (2005)
53. Daniel Wolfgang Pita-Thomas: Estudio de los factores
responsables de la regeneracion axonal en astas de ciervo
adulto. PhD Thesis, University Complutense of Madrid (2009)
54. Sally N Lawson, Mark J Perry, Elizabeth Prabhakar, Peter
W McCarthy: Primary sensory neurones: neurofilament,
neuropeptides, and conduction velocity. Brain Res Bull 30,
239-243 (1993)
55. Theodore J Price, Christopher M Flores: Critical evaluation
of the colocalization between calcitonin gene-related peptide,
substance P, transient receptor potential vanilloid subfamily
type 1 immunoreactivities and isolectin B4 binding in primary
afferent neurons of the rat and mouse. J Pain 8, 263-272
56. Jason J Ivanusic: Size, neurochemistry, and segmental
distribution of sensory neurons innervating rat tibia. J
Comp Neurol 517, 276-283 (2009)
57. Vernon Rayner, Stanley W Ewen: Do the blood vessels
of the antler velvet of the red deer have an adrenergic
innervation? Q J Exp Physiol 66, 81-6 (1981)
58. R L Garcia, Mehri Sadighi, Susan M Francis, James M
Suttie, Jean S Fleming: Expression of neurotrophin-3 in the
growing velvet antler of the red deer Cervus elaphus. J Mol
Endocrinol 19 (2), 173-182 (1997)
59. Daniel Wolfgang Pita-Thomas, Carmen Fernandez-
Martos, Monica Yunta, Rodrigo M. Maza, Rosa Navarro-
Ruiz, Marcos Javier Lopez-Rodriguez, David Reigada,
Manuel Nieto-Sampedro, Manuel Nieto-Diaz: Gene
expression of axon growth promoting factors in the deer
antler. PLoS One 5 (12), e15706 (2010)
60. Hoke A, Redett R, Hameed H, Jari R, Zhou C, Li ZB,
Griffin JW, Brushart TM: Schwann cells express motor and
sensory phenotypes that regulate axon regeneration. J
Neurosci 26 (38), 9646-9655 (2006)
61. Zarife Sahenk, Janet Oblinger, Chris Edwards:
Neurotrophin-3 deficient Schwann cells impair nerve
regeneration Exp Neurol, 212 (2), 552-556. Epub (2008)
62. Claudia Grothe, Guido Nikkhah: The role of basic
fibroblast growth factor in peripheral nerve regeneration.
Anat Embryol (Berl) 204, 171-177 (2001)
63. Jorg Mey: New therapeutic target for CNS injury? The
role of retinoic acid signaling after nerve lesions. J
Neurobiol 66, 757-779 (2006)
64. Chongyang Fu, Guangxiang Hong, Fabin Wang:
Favorable effect of local VEGF gene injection on axonal
regeneration in the rat sciatic nerve. J Huazhong Univ Sci
Technolog Med Sci 27, 186-189 (2007)
66. Ruifa Mi, Weiran Chen, Ahmet Hoke: Pleiotrophin is a
neurotrophic factor for spinal motor neurons. Proc Natl
Acad Sci USA 104, 4664-4669 (2007)
66. Charles Q. Yu, Min Zhang, Krisztina I. Matis, Charles
Kim, Mark I. Rosenblatt:Vascular endothelial growth factor
mediates corneal nerve repair. Invest Ophthalmol Vis Sci
49, 3870-3878 (2008)
67. Sawako Unezaki, Satoru Yoshii, Tamaki Mabuchi,
Akira Saito, Seiji Ito: Effects of neurotrophic factors on
nerve regeneration monitored by in vivo imaging in thy1-
YFP transgenic mice. J Neurosci Methods 178, 308-315
68. Lijoy K Mathew, Sumitra Sengupta, Jill A Franzosa,
Jessica Perry, Jane La Du, Eric A Andreasen, Robert L
Tanguay: Comparative expression profiling reveals an
essential role for raldh2 in epimorphic regeneration. J Biol
Chem 284, 33642-33653 (2009)
69. Tamara L Tal, Jill A Franzosa, Robert L Tanguay:
Molecular signaling networks that choreograph epimorphic
Deer antler innervation
fin regeneration in zebrafish - a mini-review. Gerontology
56, 231-240 (2010)
70. Rei Katogi, Yuki Nakatani, Tadasu Shin-i, Yuji Kohara,
Keiji Inohaya, Akira Kudo: Large-scale analysis of the
genes involved in fin regeneration and blastema formation
in the medaka, Oryzias latipes. Mech Dev 121, 861-872
71. Esther J Pearl, Donna Barker, Robert C Day, Caroline
W Beck: Identification of genes associated with
regenerative success of Xenopus laevis hindlimbs. BMC
Dev Biol 8, 66 (2008)
72. Kenneth D Poss, Jiaxiang Shen, Alex Nechiporuk,
Gerald McMahon, Bernard Thisse, Christine Thisse, Mark
T Keating: Roles for Fgf signaling during zebrafish fin
regeneration. Dev Biol 222, 347-358 (2000)
73. Mohamed Bouzaffour, Pascale Dufourcq, Virginie
Lecaudey, Petra Haas, Sophie Vriz: Fgf and Sdf-1
pathways interact during zebrafish fin regeneration. PLoS
One 4, e5824 (2009)
74. Jesus A Santamaria, Jose Becerra: Tail fin regeneration
in teleosts: cell-extracellular matrix interaction in blastemal
differentiation. J Anat 176, 9-21 (1991)
75. Henri E Young, Bernell K Dalley, Roger R Markwald:
Glycoconjugates in normal wound tissue matrices during
the initiation phase of limb regeneration in adult
Ambystoma. Anat Rec 223, 231-241 (1989)
76. Liria M Masuda-Nakagawa, Kenneth J Muller, John G
Nicholls: Accumulation of laminin and microglial cells at
sites of injury and regeneration in the central nervous
system of the leech. Proc Biol Sci 241, 201-206 (1990)
77. Fabian Chablais, Anna Jazwinska: IGF signaling between
blastema and wound epidermis is required for fin regeneration.
Development 137, 871-879 (2010)
78. Marco Patruno, Michael C Thorndyke, Maria Daniela
Candia Carnevali, Francesco Bonasoro, Philip W Beesley:
Growth factors, heat-shock proteins and regeneration in
echinoderms. J Exp Biol 204, 843-848 (2001)
79. Dawn E Clark, Eric A Lord, James M Suttie: Expression of
VEGF and pleiotrophin in deer antler. Anat Rec A Discov Mol
Cell Evol Biol 288, 1281-1293 (2006)
80. Angela K W Lai, Wei Lin Hou, Daniel John Verdon,
Louise F B Nicholson, Peter M. Barling: The distribution of
the growth factors FGF-2 and VEGF, and their receptors, in
growing red deer antler. Tissue Cell 39, 35-46 (2007)
81. GeneNote database Weizmann Institute of Science
82. Yu Huo, Virgil R Schirf, Wendell D Winter: The
differential expression of NGFS-like substance from fresh
pilose antler of Cervus nippon Temminck. Biomed Sci
Instrum 33, 541-543 (1997)
83. Donald Speer: The collagenous architecture of antler
velvet. In: Antler development in cervidae. Eds: Brown
R, Caesar Klever Wildlife Research Institute, TX (1982)
84. George A Bubenik: Neuroendocrine regulation of
the antler cycle. In: Horns, pronghorns and antlers. Eds:
Bubenik GA, Bubenik AB, Springer-Verlag, NY (1990)
85. Erik D Rabinovsky: The multifunctional role of
IGF-1 in peripheral nerve regeneration. Neurol Res 26,
204-210 (2004)
86. Staci D Sanford, Jesse C Gatlin, Tomas Hokfelt,
Karl H Pfenninger: Growth cone responses to growth
and chemotropic factors. Eur J Neurosci 28 (2), 268-278
87. Michel Schenker, Beat Michel Riederer, Thierry
Kuntzer, Ibtissam Barakat-Walter: Thyroid hormones
stimulate expression and modification of cytoskeletal
protein during rat sciatic nerve regeneration. Brain Res
957 (2), 259-270 (2002)
88. James M Suttie, Peter D Gluckman, John H Butler,
Peter F Fennessy, Ian D Corson, Frans J Laas: Insulin-
like growth factor 1 (IGF-1) antler-stimulating
hormone? Endocrinology 116 (2), 846-848 (1985)
89. James M Suttie, Peter F Fennessy, Ian D Corson,
Frans J Laas, Stuart F Crosbie, John H Butler, Peter D
Gluckman: Pulsatile growth hormone, insulin-like
growth factors and antler development in red deer
(Cervus elaphus scoticus) stags. J Endocrinol 121 (2),
351-360 (1989)
90. George A Bubenik, Antoine J Sempere, Joe Hamr:
Developing antler, a model for endocrine regulation of
bone growth. Concentration gradient of T3, T4, and
alkaline phosphatase in the antler, jugular, and the
saphenous veins. Calcif Tissue Int 41 (1), 38-43 (1987)
91. Francis T Lake, Gordon C Solomon, Robert W
Davis, Noel Pace, John R Morgan: Bioelectric potentials
associated with the growing deer antler. Clin Orthop
Relat Res 142, 237-43 (1979)
92. Colin D McCaig, Ann M Rajnicek, Bing Song, Min
Zhao: Has electrical growth cone guidance found its
potential? Trends Neurosci 25 (7), 354-359 (2002)
93. George A Bubenik, Anthony B Bubenik, E Don
Stevens, Allan G Binnington: The effect of neurogenic
stimulation on the development and growth of bony
tissues. J Exp Zool 219 (2), 205-216 (1982)
94. Bryan J Pfister, Akira Iwata, David F Meaney,
Douglas H Smith: Extreme stretch growth of integrated
axons. J Neurosci 24 (36), 7978-7983 (2004)
Deer antler innervation
95. Douglas H Smith: Stretch growth of integrated axon
tracts: extremes and exploitations. Prog Neurobiol 89 (3),
231-239 Epub (2009)
96. Manuel Nieto-Diaz, Daniel Wolfgang Pita-Thomas,
Rodrigo M Maza, Monica Yunta, Marcos Javier Lopez-
Rodriguez, Rosa Navarro-Ruiz, David Reigada, Carmen
Fernandez-Martos, Manuel Nieto-Sampedro: Factors
promoting axon growth in the deer antler. Animal Prod
Science 51, 1-4 (2011)
97. Xiao Z Liu, Xiao M Xu, Rong Hu, Cheng Du, Shu X
Zhang, John W McDonald, Hong X Dong, Ying J Wu,
Guang S Fan, Mark F Jacquin, Chung Y Hsu, Dennis W
Choi: Neuronal and glial apoptosis after traumatic spinal
cord injury. J Neurosci 17, 5395-5406 (1997)
98. Michael S Beattie, Akhlaq A Farooqui, Jaqueline C
Bresnahan: Review of current evidence for apoptosis after
spinal cord injury. J Neurotrauma 17, 915-925 (2000)
99. Per A R Ekstrom: Neurones and glial cells of the mouse
sciatic nerve undergo apoptosis after injury in vivo and in
vitro. Neuroreport 6 (7), 1029-1032 (1995)
100. Ivan S Raginov, Yu A Chelyshev: Post-traumatic
survival of sensory neurons of different subpopulations.
Neurosci Behav Physiol 35 (1), 17-20 (2005)
101. Susan Y Fu, Tessa Gordon: The cellular and molecular
basis of peripheral nerve regeneration. Mol Neurobiol 14
(1-2), 67-116 (1997)
Abbreviations: CNS central nervous system, PNS
peripheral nervous system, NF200 200Kda neurofilament,
CGRP Calcitonin-gene related peptide, IB4 isolecting B4,
GFAP glial fibrillary acidic protein, NGF nerve growth
factor, NT-3 neurotrophin 3, BDNF brain derived
neurotrophic factor, GPI glucose phosphate isomerase,
MTRN meteorin, MDK midkine, NRCAM neuronal cell
adhesion molecule, FGFb basic fibroblast growth factor,
EGF epidermal growth factor, PEDF pigment epithelium
derived factor, TGFbeta transforming growth factor beta,
BMP bone morphogenetic protein, IGF insulin-like growth
factor, VEGF vascular endothelial growth factor, DRG
dorsal root ganglia.
Key Words Deer antler, Velvet, Endocrine, Paracrine
regulation, Fast axon growth, Trigeminal nerve, Nervous
system injury, Spontaneous regeneration, Neuron, Gene
expression, in vitro analysis
Send correspondence to: Manuel Nieto-Diaz, Molecular
Neuroprotection Group, Hospital Nacional de Paraplejicos
(SESCAM), Finca La Peraleda s/n. 45071 Toledo, Spain,
Tel: 34 925396834, Fax: 34 925247745, E-mail:
... Growing antlers are profusely innervated by solely sensory fibers regenerating from the pedicle sensory nerve stumps, which are derived from the zygomaticotemporal and supraorbital branches of the trigeminal nerve [81][82][83][84]. The newly regenerative antler nerves also elongate within the vascular layer peripherally, following very closely the distribution of antler arteries [11,81,83,85]. ...
... Growing antlers are profusely innervated by solely sensory fibers regenerating from the pedicle sensory nerve stumps, which are derived from the zygomaticotemporal and supraorbital branches of the trigeminal nerve [81][82][83][84]. The newly regenerative antler nerves also elongate within the vascular layer peripherally, following very closely the distribution of antler arteries [11,81,83,85]. The profuse nerve fibers occur in small bundles, projecting from the vascular layer towards more superficial layers of the velvet skin [27,83]. ...
... The newly regenerative antler nerves also elongate within the vascular layer peripherally, following very closely the distribution of antler arteries [11,81,83,85]. The profuse nerve fibers occur in small bundles, projecting from the vascular layer towards more superficial layers of the velvet skin [27,83]. In antler tip, the nerve fibers follow the route of the artery branches, abundantly locating at the deep connective tissue above the reserve mesenchyme of the antler [83,86,87]. ...
Periosteum is essential for bone regeneration and damage repair in mammals. Most species of deer family (Cervidae) develop two kinds of special periosteum, antler periosteum and pedicle periosteum, both supporting the complete regeneration of antler. Antler is the bone organ with the fastest growth rate in mammals. Along with the fast growth of antler, its external tissues such as blood vessels, nerves and the covering skin also grow rapidly. Currently, it is still unclear whether antler periosteum contributes to the fast growth of antler and how. It is also unclear why the regenerative capacity of antler periosteum is weaker than that of pedicle periosteum. In this study, the in vitro culture system for antler periosteal cells (AnPC) was constructed for the first time using the mid-beam antler periostea during antler fast-growth period. According to our results, the cultured AnPC expressed classical MSC markers, consistent with the pedicle periosteal stem cells (PPSC). However, the fluorescence intensities of the MSC markers on AnPC were significantly weaker than those on PPSC. In addition, AnPC showed much lower proliferation rates than PPSC. The proliferation rates of the AnPC also gradually decreased after successive passages, while the proliferation rates of the pedicle periosteal stem cells remained unchanged. These findings may partially explain the weaker regenerative capacity of antler periosteum. Further comparative global gene analysis revealed clearly the different gene expressed patterns between AnPC and PPSC. AnPC may mainly function on promoting angiogenesis, nerve growth and intramembrane bone formation during antler regeneration, whereas PPSC may primarily be involved in androgen signaling receptor pathway and PI3K-Akt signaling pathway and function on maintaining stem cell renewal.
... Throughout this growth period, the antlers become enveloped by an integument which supplies the growing antler with blood from the arteries of its vascular layer, and is critical for the metabolic demands of the regenerating tissue [30]. The integument is highly innervated by sensory fibres of the trigeminal nerve, whose exons grow at a pace accompanying that of the antler (2 cm/day) [83];. Rapid antler bone growth occurs through a combination of modified endochondral ossification within each antler distal tip following cartilage formation, and intramembranous ossification within the shaft which increases antler diameter [6,54]. ...
... Rapid growth at the tip of deer antlers is sustained by distinct populations of ASCs. Reserve mesenchymal stem cells (RMCs) within the RM undergo intense and sequential proliferation [83]; [112,114,115], mediated by signals from growth factors, including melatonin and IGF-1 [118,119]. Chondro/osteogenic differentiation of RMCs is regulated by the increased expression of genes which are characteristic of robust mammalian cartilage and bone development, including fibroblast growth factor (FGF), TGF-β, Wnt, and bone morphogenetic proteins (BMPs) [41,92]. ...
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The freshwater crayfish is capable of regenerating limbs, following autotomy, injury and predation. In arthropod species, regeneration and moulting are two processes linked and strongly regulated by ecdysone. The regeneration of crayfish limbs is divided into wound healing, blastema formation, cellular reprogramming and tissue patterning. Limb blastema cells undergo proliferation, dedifferentiation and redifferentiation. A limb bud, containing folded segments of the regenerating limb, is encased within a cuticular sheath. The functional limb regenerates, in proecdysis, in two to three consecutive moults. Rapid tissue growth is regulated by hormones, limb nerves and local cells. The TGF‐β/activin signalling pathway has been determined in the crayfish, P. fallax f. virginalis, and is suggested as a potential regulator of tissue regeneration. In this review article, we discuss current understanding of tissue regeneration in the crayfish and various crustaceans. A thorough understanding of the cellular, genetic and molecular pathways of these biological processes is promising for the development of therapeutic applications for a wide array of diseases in regenerative medicine.
... Throughout this growth period, the antlers become enveloped by an integument which supplies the growing antler with blood from the arteries of its vascular layer, and is critical for the metabolic demands of the regenerating tissue [30]. The integument is highly innervated by sensory fibres of the trigeminal nerve, whose exons grow at a pace accompanying that of the antler (2 cm/day) [83];. Rapid antler bone growth occurs through a combination of modified endochondral ossification within each antler distal tip following cartilage formation, and intramembranous ossification within the shaft which increases antler diameter [6,54]. ...
... Rapid growth at the tip of deer antlers is sustained by distinct populations of ASCs. Reserve mesenchymal stem cells (RMCs) within the RM undergo intense and sequential proliferation [83]; [112,114,115], mediated by signals from growth factors, including melatonin and IGF-1 [118,119]. Chondro/osteogenic differentiation of RMCs is regulated by the increased expression of genes which are characteristic of robust mammalian cartilage and bone development, including fibroblast growth factor (FGF), TGF-β, Wnt, and bone morphogenetic proteins (BMPs) [41,92]. ...
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Generally, mammals are unable to regenerate complex tissues and organs however the deer antler provides a rare anomaly to this rule. This osseous cranial appendage which is located on the frontal bone of male deer is capable of stem cell-based organogenesis, annual casting, and cyclic de novo regeneration. A series of recent studies have classified this form of regeneration as epimorphic stem cell based. Antler renewal is initiated by the activation of neural crest derived pedicle periosteal cells (PPCs) found residing within the pedicle periosteum (PP), these PPCs have the potential to differentiate into multiple lineages. Other antler stem cells (ASCs) are the reserve mesenchymal cells (RMCs) located in the antlers tip, which develop into cartilage tissue. Antlerogenic periosteal cells (APCs) found within the antlerogenic periosteum (AP) form the tissues of both the pedicle and first set of antlers. Antler stem cells (ASCs) further appear to progress through various stages of activation, this coordinated transition is considered imperative for stem cell-based mammalian regeneration. The latest developments have shown that the rapid elongation of the main beam and antler branches are a controlled form of tumour growth, regulated by the tumour suppressing genes TP73 and ADAMTS18. Both osteoclastogenesis, as well as osteogenic and chondrogenic differentiation are also involved. While there remains much to uncover this review both summarises and comprehensively evaluates our existing knowledge of tissue regeneration in the deer antler. This will assist in achieving the goal of in vitro organ regeneration in humans by furthering the field of modern regenerative medicine. The Translational potential of this article As a unique stem cell-based organ regeneration process in mammals, the deer antler represents a prime model system for investigating mechanisms of regeneration in mammalian tissues. Novel ASCs could provide cell-based therapies for regenerative medicine and bone remodelling for clinical application. A greater understanding of this process and a more in-depth defining of ASCs will potentiate improved clinical outcomes.
... If ingestion of a reindeer's antlers with sporadic prion disease was the event that initiated the outbreak of CWD in Norway, prions would have to be present in the antlers. Antlers show an impressive growth rate of up to 2 to 3 cm per day, where growing antler tissue is well innervated with sensory nerves originating from the trigeminal nerve 22 , i.e., cranial nerve V. Infective prions (PrP Sc ) were found in the trigeminal ganglia of patients with Creutzfeldt-Jacobs disease 23 and in other tissues innervated by the trigeminal nerve (cornea and gingiva). Antler bones remain viable and well perfused with blood vessels for several months after velvet shedding 24 . ...
... Transfer of prions to antlers H1 PrP Sc in disintegrating axons integrated in antler bone matrix. Velvet and growth cartilage innervated from the trigeminal nerve 22 . PrP Sc is found in the trigeminal ganglia and tissues innervated therefrom in sCJD patients 23 H2 PrP Sc in antler bone marrow. ...
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Prion diseases constitute a class of invariably fatal and degenerative encephalopathies. Chronic Wasting Disease (CWD) is a contagious prion disease among cervids, which is spreading and causing marked population declines in USA and Canada. The first outbreak of CWD in Europe was discovered in a reindeer population in Norway in 2016. In the worst-case scenario with continental-wide spreading of CWD in Eurasia, an annual harvest of around 4 million cervids is at stake only in Europe, with huge economic and cultural significance. An in situ origin of CWD was suspected, and it appear urgent to identify the likely cause to prevent future emergences. Here, we document the novel phenomenon of extensive antler cannibalism prior to shedding among reindeer in the CWD-infected population. The extent of antler cannibalism increased over the last decades when CWD emerged, and included ingestion of vascularized antlers. Ingestion of tissues from conspecifics is a risk factor for the emergence of prion diseases, where the presence of extensive antler cannibalism opens the intriguing possibility of a ‘Kuru-analogue’ origin of CWD among the reindeer in Europe. Based on general insight on pathology of prion diseases and strain selection processes, we propose an hypothesis for how contagious CWD may emerge from sporadic CWD under the unique epidemiological conditions we document here. More research is required to document the presence of prions in reindeer antlers, and whether antler cannibalism actually led to a strain selection process and the emergence of a contagious form of CWD from a sporadic form of CWD.
... It was observed that when the algometer was applied rapidly after initial touch, the deer reacted quickly in a more predictable manner. This could be explained by the sensory nerve pathways arising from the trigeminal nerve (CNV) in growing antlers and A-delta fibres, which are the receptors for pressure and which transmit pain information from an acute noxious stimulus [39,40]. Alternatively, if the algometer is applied slowly, it is more likely that the reaction would be due to A-beta fibres that transmit information from skin receptors and C fibres, which are slow pain receptors [39,40]. ...
... This could be explained by the sensory nerve pathways arising from the trigeminal nerve (CNV) in growing antlers and A-delta fibres, which are the receptors for pressure and which transmit pain information from an acute noxious stimulus [39,40]. Alternatively, if the algometer is applied slowly, it is more likely that the reaction would be due to A-beta fibres that transmit information from skin receptors and C fibres, which are slow pain receptors [39,40]. ...
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New Zealand deer farming centres on the production of meat and velvet antler. Velvet antler removal is a painful procedure and currently, New Zealand Animal Welfare regulations dictate surgical removal of velvet antlers under lignocaine anaesthesia. To improve our knowledge on the efficacy and duration of other local anaesthetics to mitigate pain after antler removal, it is important to accurately assess and quantify pain arising from antler removal. Therefore, the current study was designed to validate mechanical nociceptive threshold (MNT) testing using a Wagner hand-held algometer, and to apply this methodology to assess the efficacy and duration of action of articaine for antler removal in deer. Baseline force (N) required to elicit the nociceptive response was recorded in 40 yearling male red deer on three alternate days. Ten of the 40 animals were selected for antler removal after administration of 4% articaine hydrochloride as a ring block. The duration of analgesic efficacy of articaine was assessed by algometry across 5 time points. There was a significant difference in MNTs among the three days (day 3 versus day 1 (p < 0.0001), day 2 versus day 1 (p < 0.0001), and day 1 versus day 2 (p < 0.01)). Positive correlations were observed between weight, antler length and thresholds. The MNT values remained above 20N for 6 h after removal of velvet antlers under the articaine ring block. This study provides valuable information about the use of MNT in red deer. These findings lay a foundation for future studies in the topics of peri-operative and postoperative pain management in deer antler removal, and a possible alternative use for articaine.
... In addition to VEGF, the special biological process of antler chondrocytes is also inseparable from the regulation of a large number of growth factors. According to the literature, the apical tissue of antler contains the expression of many growth factors, including TGF-β1 and TGF-β2 [48], among which TGF-β is one of the many growth factors. TGF-β has many biological functions, such as cell proliferation, differentiation, apoptosis, and embryogenesis (A. ...
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Antler is the secondary sexual characteristic of deer, which develops on the forehead at puberty. It is the only organ that can be regenerated entirely in mammals. Therefore, it is often used as a research model in the field of organ regeneration and wound repair. Many growth factors and proteins play an active role throughout the developmental process of antler regeneration. With the rapid development of sequencing technology, more and more noncoding RNAs (ncRNAs) have been discovered, and the relationship between ncRNA and antler regeneration has gradually become clear. This paper focuses on the research progress of several ncRNAs (including miRNA and lncRNA) in deer antler tissues, which are helpful to reveal the molecular mechanism of deer antler regeneration at the molecular level.
... Deer antler has a stable growth cycle, grows with a fixed shape and position, and can be completely regenerated every year, making it an ideal mammalian morphogenesis model [20]. Previous studies have shown that deer antler derives from the proliferation and differentiation of antlerogenic periosteum (AP) stem cells [21]. ...
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Background: Deer antler is the only mammalian organ that can be completely regenerated every year. Its periodic regeneration is regulated by multiple factors, including transforming growth factor β (TGF-β). This widely distributed multi-functional growth factor can control the proliferation and differentiation of many types of cell, and it may play a crucial regulatory role in antler regeneration. This study explored the role of TGF-β1 during the rapid growth of sika deer antler. Methods: Three CRISPR-Cas9 knockout vectors targeting the TGF-β1 gene of sika deer were constructed and packaged with a lentiviral system. The expression level of TGF-β1 protein in the knockout cell line was determined using western blot, the proliferation and migration of cartilage cells in vitro were respectively determined using EdU and the cell scratch test, and the expression levels of TGF-β pathway-related genes were determined using a PCR array. Results: Of the three gRNAs designed, pBOBI-gRNA2 had the best knockout effect. Knockout of TGF-β1 gene inhibits the proliferation of cartilage cells and enhances their migration in vitro. TGF-β signaling pathway-related genes undergo significant changes, so we speculate that when the TGF-β pathway is blocked, the BMP signaling pathway mediated by BMP4 may play a key role. Conclusions: TGF-β1 is a newly identified regulatory factor of rapid growth in sika deer antler.
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Deer antlers are male secondary sexual characters that develop to become bone; they are unique appendages that, once lost, can fully regenerate from the permanent bony protuberances or pedicles. Pedicle periosteum (PP) is the tissue that gives rise to the regenerating antlers with three differentiation stages, namely, dormant (DoPP), potentiated (PoPP), and activated (AcPP). Thus far, the transition from the PoPP to the AcPP has not been studied. Our results showed that the AcPP cells maintained their original stem cell features by expressing mesenchymal stem cell (MSC) markers CD73, CD90, and CD105, although they had entered the proliferation mode. The differentially expressed genes (DEGs) in the AcPP compared with those of the PoPP were mainly involved in protein processing, cell cycle, and calcium signaling pathways. Calreticulin (CALR), an androgen response gene, was significantly differentially upregulated in the AcPP cells, and its expression level was negatively regulated by androgens, in contrast to the currently known model systems where all regulation is positive. The downregulation of CALR expression in the AcPP cells in vitro inhibited cell proliferation, induced apoptosis, and inhibited cell cycle progression at G1-S transition. Therefore, CALR is likely a downstream mediator of androgen hormones for triggering initiation of antler regeneration. We believe that the identification of CALR has not only discovered “one critical piece” of the “jigsaw puzzle” in the initiation of antler regeneration but also helps in revealing the mechanism underlying this unique mammalian epimorphic regeneration and has also opened a new avenue for the study of the nature of CALR regulation by androgen (putative binding partners), thus facilitating the identification of potential molecule(s) for investigation as targets for clinical evaluation.
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The vertebrate nervous system exhibits dramatic variability in regenerative capacity across species and neuronal populations. For example, while the mammalian central nervous system (CNS) is limited in its regenerative capacity, the CNS of many other vertebrates readily regenerates after injury, as does the peripheral nervous system (PNS) of mammals. Comparing molecular responses across species and tissues can therefore provide valuable insights into both conserved and distinct mechanisms of successful regeneration. One gene that is emerging as a conserved pro-regenerative factor across vertebrates is activating transcription factor 3 (ATF3), which has long been associated with tissue trauma. A growing number of studies indicate that ATF3 may actively promote neuronal axon regrowth and regeneration in species ranging from lampreys to mammals. Here, we review data on the structural and functional conservation of ATF3 protein across species. Comparing RNA expression data across species that exhibit different abilities to regenerate their nervous system following traumatic nerve injury reveals that ATF3 is consistently induced in neurons within the first few days after injury. Genetic deletion or knockdown of ATF3 expression has been shown in mouse and zebrafish, respectively, to reduce axon regeneration, while inducing ATF3 promotes axon sprouting, regrowth, or regeneration. Thus, we propose that ATF3 may be an evolutionarily conserved regulator of neuronal regeneration. Identifying downstream effectors of ATF3 will be a critical next step in understanding the molecular basis of vertebrate CNS regeneration.
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Long non-coding RNA (lncRNA) is a transcription product of the mammalian genome that regulates the development and growth in the body. The present study aimed to analyze the expression dynamics of lncRNA in sika antler mesenchymal and cartilage tissues by high-throughput sequencing. Bioinformatics was applied to predict differentially expressed lncRNAs and target genes and screen lncRNAs and mRNAs related to osteogenic differentiation, cell proliferation, and migration. Finally, the expression of the lncRNAs and target genes were analyzed by qRT-PCR. The results showed that compared to the cartilage tissue, the transcription levels of lncRNA and mRNA, 1212 lncRNAs and 518 mRNAs, in mesenchymal tissue were altered significantly. Thus, a complex interaction network was constructed, and the lncRNA-mRNA interaction network correlation related to osteogenic differentiation, cell proliferation, and migration was analyzed. Among these, the 26 lncRNAs and potential target genes were verified by qRT-PCR, and the results of qRT-PCR were consistent with high-throughput sequencing results. These data indicated that lncRNA promotes the differentiation of deer antler mesenchymal tissue into cartilage tissue by regulating the related osteogenic factors, cell proliferation, and migration-related genes and accelerating the process of deer antler regeneration and development.
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During their annual regeneration, antlers are innervated by trigeminal sensory axons growing at the highest rate recorded for any adult mammal. Previous analyses established the presence in the antler of nerve growth factor and neurotrophin 3 neurotrophins, which may underlie this rapid nerve growth. We are currently exploring the expression of other molecules that may be involved in such growth (axon growth promoters) combining several gene-expression techniques. Preliminary results indicate the expression of different growth promoters in the antler velvet, five of them not previously described in deer. The expression of these molecules as well as others described in the literature suggests that antler velvet promotes axon growth. However, most promoters expressed in the velvet are also present in unmodified deer skin. Thus, it must be asked why axons grow so fast in the antler? To answer that question, we developed a series of in vitro experiments using sensory neurons from adult and embryo rodents. These studies suggested that soluble proteins secreted by the velvet strongly promote neurite outgrowth. Using specific blocking antibodies, we demonstrated that nerve growth factor is partially responsible for these effects although other yet unidentified proteins seem also to be involved. The studies also showed that neither endocrine serum factors nor antler tissue substrate stimulate neurite outgrowth, although deep velvet layers cause neurite outgrowth orientation.
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Antlers constitute the only mammal model for limb regeneration. A number of factors affect antler regeneration. In this review, we examine such factors and the potential consequences for organ regeneration. As body mineral stores are depleted to grow antlers, physiological exhaustion is shown in the mineral composition, mechanical performance and, according to preliminary studies, porosity of the antler bone material. Nutrition plays an important role in antler characteristics. Thus, antler composition can be used as a diagnostic tool to assess mineral deficiencies in deer. Studies on ecological effects of exceptional weather in plants suggest that minor minerals, particularly Mn, may play disproportionately roles in mechanical performance of bone material. This suggests that Mn (and perhaps other minerals) is essential to incorporate Ca and P from resorbed skeleton material in antlers. Apart from implications for game management, some effects may have applications for medicine.
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The annual regeneration cycle of deer (Cervidae, Artiodactyla) antlers represents a unique model of epimorphic regeneration and rapid growth in adult mammals. Regenerating antlers are innervated by trigeminal sensory axons growing through the velvet, the modified form of skin that envelopes the antler, at elongation velocities that reach one centimetre per day in the common deer (Cervus elaphus). Several axon growth promoters like NT-3, NGF or IGF-1 have been described in the antler. To increase the knowledge on the axon growth environment, we have combined different gene-expression techniques to identify and characterize the expression of promoting molecules not previously described in the antler velvet. Cross-species microarray analyses of deer samples on human arrays allowed us to build up a list of 90 extracellular or membrane molecules involved in axon growth that were potentially being expressed in the antler. Fifteen of these genes were analysed using PCR and sequencing techniques to confirm their expression in the velvet and to compare it with the expression in other antler and skin samples. Expression of 8 axon growth promoters was confirmed in the velvet, 5 of them not previously described in the antler. In conclusion, our work shows that antler velvet provides growing axons with a variety of promoters of axon growth, sharing many of them with deer's normal and pedicle skin.
The antlers are one of the secondary sexual characters of males in all genera of Cervidae except Rangifer (where antlers are found in both sexes). Their development, seasonal renewal, maturation, and casting are closely related to the activity of the reproductive system. This fact was already well known to Aristotle, who described the alterations of antlerogenesis caused by castration (Goss 1968). However, it took another 2,000 years before progress in chemistry and endocrinology made it possible to investigate the detail roles of various humoral factors in the regulation of the antler cycle.
Deer antlers and their antecedent pedicles are made up of two components, interior osseocartilage and exterior integument. In a previous study, we described that histogenesis of the interior osseocartilage proceeds through four ossification stages. These are intramembranous (IMO), transition (OPC), pedicle endochondral (pECO), and antler endochondral (aECO). In the present study, we used histological techniques to examine pedicle skin formation and its transformation to antler velvet. The results showed that pedicle skin initiated from the apex of a frontal lateral crest and was formed through three distinctive stages. These stages are 1) compression of the subcutaneous loose connective tissue at the OPC stage, 2) stretching of the undulated epidermis at the early pECO stage, and 3) neogenesis of the skin and its associated appendages at the mid pECO stage. Transformation into antler velvet, which occurs at the late pECO stage, is mainly associated with alteration in the skin appendages. This alteration includes the loss of arrector pili muscle and sweat glands, and the gain of the large bi- or multi-lobed sebaceous glands. These results suggest that pedicle skin expansion occurs to release the mechanical tension created by underlying forming antlerogenic tissue, initially in response to it by mechanical stretch, and then by neogenesis of skin. In turn, the stretched pedicle skin may exert mechanical pressure on the underlying antlerogenic tissue causing it to change in ossification type. Antler velvet generation may be accomplished by both mechanical stimulation and chemical induction from the underlying pECO stage antlerogenic tissue. If this hypothesis is correct it is likely that mechanical stimulation would drive skin formation and chemical induction then determine skin type. Furthermore, asynchronous transformation of the interior and exterior components during pedicle formation and antler generation may result from the delayed chemical induction and the way antler velvet initially generates. The results from both mitotic cell labelling of the basal layer and ultrastructure of the basement membrane of the apical skin in the study support these hypotheses. Anat Rec 260:62–71, 2000. © 2000 Wiley-Liss, Inc.
The aims of this study were to describe first antler growth rate and the factors affecting it. Growth rate reached a peak in week 14. Mean first antler growth rate during the whole growing period was 1.95 ± 0.05cm/week. Lactation influenced antler growth rate as this was positively affected by total protein yield, while antlers grew at a lower rate the later they started to grow.
Following amputation of a urodele limb or teleost fin, the formation of a blastema is a crucial step in facilitating subsequent regeneration. Using the zebrafish caudal fin regeneration model, we have examined the hypothesis that fibroblast growth factors (Fgfs) initiate blastema formation from fin mesenchyme. We find that fibroblast growth factor receptor 1 (fgfr1) is expressed in mesenchymal cells underlying the wound epidermis during blastema formation and in distal blastemal tissue during regenerative outgrowth. fgfr1 transcripts colocalize with those of msxb and msxc, putative markers for undifferentiated, proliferating cells. A zebrafish Fgf member, designated wfgf, is expressed in the regeneration epidermis during outgrowth. Furthermore, we show that a specific inhibitor of Fgfr1 applied immediately following fin amputation blocks blastema formation, without obvious effects on wound healing. This inhibitor blocks the proliferation of blastemal cells and the onset of msx gene transcription. Inhibition of Fgf signaling during ongoing fin regeneration prevents further outgrowth while downregulating the established expression of blastemal msx genes and epidermal sonic hedgehog. Our findings indicate that zebrafish fin blastema formation and regenerative outgrowth require Fgf signaling.