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Advances in Evolutionary Developmental Biology, First Edition. Edited by J. Todd Streelman.© 2014 John Wiley & Sons, Inc. Published 2014 by John Wiley & Sons, Inc.

ECTODERMAL ORGAN STEM CELLS: MORPHOGENESIS, POPULATION

REGENERATIVE BEHAVIOR, AND EVO-DEVO

7

PHYSIOLOGICAL REGENERATION OF ECTODERMAL ORGAN STEM CELLS

Regenerative biology and medicine is emerging as one of the most fascinating fields with potentially high clinical applications. Nature is a rich resource from which to learn how to engineer stem cells for the application of regenerative medicine, particularly for integu-mentary ectodermal organs. Integumentary ectodermal organs include hairs, feathers, scales, horns, nails, claws, teeth, and glands (i.e., sebaceous, sweat, and mammary) (Chuong 1998; Wu et al. 2004) (Figure 7.1A). These organs are at the interface between an organism and its external environment and therefore face constant wear and tear. Animals have evolved successful regenerative mechanisms to accommodate renewal with minimal functional interruption.

Episodic Stem Cell Regeneration

Different ectodermal organs can use diverse regeneration strategies. These can range from continuous renewal of the epidermis to the episodic regeneration of hairs and feathers (Petersson et al. 2011; Wu et al. 2004). Reptile bodies are covered by different types of scales (Chang et al. 2009). Birds typically have two kinds of scales: scutate and reticulate on their feet. In some reptiles, such as the crocodile, scales undergo continuous renewal.

Ping Wu,1 Ang Li,1 Jun Yin,2 Randall Widelitz,1 and Cheng-Ming Chuong1

1Department of Pathology, Keck School of Medicine, University of Southern California, Los Angeles, CA

2College of Life Science, Inner Mongolia Agricultural University, Huhhot, China

Figure 7.1. Evo-devo of ectodermal organ. (A) A conceptual animal with different forms of

ectodermal skin appendages. Endodermal organs are also shown. Modified from (Chuong 1998).

(B) Hair follicle. (B’) Feather follicle (adapted from Lin et al. 2006). (B”) Reptilian tooth family

unit. (C) Regeneration cycling of skin appendages. Episodic regeneration leads to alternative

growing and resting phases. Three follicles shown in B allow different topologies for regenera-

tion. (D) Hatchling chick and adult peasant with sexual dimorphism. (E) Evo-devo of ectodermal

appendages. The X coordinate represents new developmental mechanisms. The Y coordinate

represent new phenotypes. They all start from a flat layer of epidermis. Those in the upper right

quadrant used more novel developmental mechanisms and are more complex. Arrows indicate

possible topological transformation from one form to another, not necessarily indicating the

evolutionary process. The processes are in red, and the names of the forms are in blue (adapted

from Wu et al. 2004).

PHYsioLogiCAL REgEnERATion oF ECToDERMAL oRgAn sTEM CELLs 135

In snakes and lizards, the scale epidermis undergoes episodic molting and regeneration (Chang et al. 2009). In mammals interappendage epidermis undergoes continuous renewal. However, mammalian hairs and avian feathers undergo episodic regeneration. To achieve this, both hairs and feathers have evolved a follicle configuration which allows organo-genesis to occur within a protected space before the tissue differentiates (Figure 7.1B, B′). Differentiated hair or feather filaments grow distally and protrude out of the follicle and body surface. After a certain time, the appendages fall out, but the stem cells and dermal papilla remain in the follicle waiting for activation to enter the next growth phase. Thus, regeneration occurs cyclically (Figure 7.1C). It also offers an opportunity for the follicles to make new appendage phenotypes at different physiological stages (Chuong et al. 2012). For example, a baby galliform chick can grow up to show spectacular, sexually dimorphic feathers (Figure 7.1D).

Follicle Configuration for Stem Cell Niche

This follicle configuration evolved independently for mammals and birds. For hair folli-cles, stem cells are located at the bulge region beneath the sebaceous gland (Figure 7.1B) (Fuchs 2009; Morris et al. 2004). In catagen, the lower follicle undergoes apoptosis. The hair follicles then enter telogen. Later, epithelial stem cells are activated by factors secreted by the dermal papilla and the extrafollicular environment (see later), causing hair follicles to enter anagen. Feather stem cells are configured as a ring, sitting near the fol-licle base above the dermal papilla (Figure 7.1B′) (Yue et al. 2005). In the growth phase, collar epithelium cells proliferate and differentiate over time as they are pushed distally. After each feather reaches a certain length, it enters resting phase. The collar epidermis starts to keratinize. Stem cells shift down to the follicle base, tightly tucked next to the dermal papilla. Keratinization of the collar allows the feather shaft to molt. After that, the dermal papilla is re-epithelialized, and the feather follicle reenters growth phase (Figure 7.2A).

The tooth follicle is different. In the mouse, incisor stem cells are at the follicle base which allows for tooth elongation (Harada et al. 1999). However, if a tooth is plucked, stem cells are also removed and the tooth cannot regenerate. In reptiles, teeth can regener-ate, and they do this by forming a tooth family unit. The follicle becomes complex as multiple tooth germs within a family are at different developmental stages heterochron-ically (Figure 7.1B″). Typically, there are three components within a tooth family: a functional tooth, a replacement tooth, and a dental lamina. When a functional tooth is lost, the replacement tooth grows to become the new functional tooth. Simultaneously, the dental lamina, which contains stem cells, gives rise to a new replacement tooth but leaves a cluster of stem cells to replenish the dental lamina. This mechanism exists in reptiles and fish. The molecular pathway controlling tooth replacement in fish (Fraser and Smith 2011; Van der Heyden et al. 2000) and lizards (Handrigan and Richman 2010; Handrigan et al. 2010) has been studied. Putative stem cells have been localized in the gecko dental laminae (Handrigan et al. 2010).

Overall during evolution, ectodermal organs became more and more complex (Figure 7.1E). Originally, the ectoderm was smooth as in amphioxus. Later in development, fish developed scales and glandular structures. Reptile scales formed as effective barriers to protect movement within their terrestrial environment. In mammals and birds, follicular structures evolved. In feathers, further branching morphogenesis occurred (Wu et al. 2004). In the following part, we will discuss stem cells in feather follicles and then we will discuss the stem cell activity in hair waves.

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Figure 7.2. Feather cycling and morphogenesis. (A) Feather cycling from initiation to growth

then to resting phase. (B) Localization of transient amplifying cells (TA cells) in feather follicle at

growth and resting phases. (B’) Localization of label retention cells (LRC) in feather follicles at

growth and resting phases. in the epidermis, only stem cells are labeled (blue arrow). Pulp cells

are extensively labeled. (C) The branching morphogenesis in bilateral symmetric feather (adapted

from (Wu et al. 2004). (C’) Enlarged diagram showing branching morphogenesis. ap, axial plate;

bb, barbule; bp, barbule plate; br, barb ridge; gZ, growth zone; pe, pulp epithelium; rm, ramus.

FEATHER REGENERATION: STEM CELL HOMEOSTASIS AND MORPHOGENESIS

Stem Cell Homeostasis

Feathers are replaced episodically, and the feathers are known to cycle through different phases of initiation, growth, rest, and molting. During growth phase, the feather follicle

FEATHER REgEnERATion: sTEM CELL HoMEosTAsis AnD MoRPHogEnEsis 137

is vase-shaped with a cylindrical epithelial wall, a base made of the dermal papilla, and a core made of feather pulp (Figure 7.2B, B′). Proliferating cells are in the multilayered epidermis above the dermal papilla, named the epithelial collar. A small cluster of slow-cycling stem cells are located in the collar region named the collar bulge (Figure 7.2B′) (Yue et al. 2005). In three dimensions these stem cells are distributed in a ring configuration sitting at the bottom of the feather follicle. As feathers mature from the distal end, the basal epithelium together with the mesenchymal pulp retreats toward the proximal end and forms periodically positioned pulp caps. This involves the apoptosis of pulp cells allowing the filament cylinder to open and the feather vane to form. The mature feather, the part protrud-ing out of the body surface, is composed of cells that have keratinized and died. It is still connected to the follicle through the proximal end of the calamus (feather shaft proximal to the vane). Toward the end of the growing phase, feather stem cells descend from the collar bulge to be in close contact with the dermal papilla (Yue et al. 2005). Eventually, the collar epidermis thins, and the growth phase transits to the resting phase.

Feather Branching Morphogenesis

Feathers have distinct morphology with specialized functions covering different parts of the bird (Lucas and Stettenheim 1972; Yu et al. 2004). For example, the lower half of body feathers is plumulaceous and provides warmth for thermoregulation. In contrast, tail and crown feathers are pennaceous, forming a large vane used for communication. Wing feath-ers are also pennaceous and use their vane to move the air in order to fly. Not only do each of these feathers have a unique shape and texture, but they also are positioned in specific body regions to maximize their effectiveness in adapting each bird to its environ-ment. Other feather characteristics further enhance their function. For example, body feathers are radially symmetric, and flight feathers are bilaterally symmetric, which aids in aerodynamics.

Recent progress in molecular and developmental biology has allowed us to dig into the molecular regulation of different transition steps in this complex morphogenetic process (Figure 7.2C, C′; Figure 7.3A–E). Our lab has developed a novel powerful model to analyze adult feather follicle morphogenesis (Yu et al. 2002). RCAS viruses have been used to express candidate genes or dominant negative forms of genes. The virus is added to plucked feather follicles and feathers are allowed to regenerate. The regenerated feathers carry the mis-expressed genes and may exhibit abnormalities if the tested genes are involved in morphogenesis (Yu et al. 2002). We reported that BMP4 has a critical role in feather branching. Mis-expression of BMP2 and BMP4 caused barb fusion, while mis-expression of Noggin promoted further branching of barbs. Shh was found to induce apoptosis of the marginal plate epithelia (Yu et al. 2002). Therefore, it is likely that the transition of unbranched to branched feathers happened through a periodic inhibition of BMP signaling and activation of Shh signaling.

The normal growth of feathers involves temporal and spatial regulated apoptotic events. The effects of imbalanced Shh on apoptosis were tested (Chang et al. 2004). Shh suppression reduced marginal plate apoptosis and caused abnormal differentiation of barbule plates. Shh overexpression enhanced proliferation in barb ridges. Expression of Patched in the barbule plate epithelia implies a paracrine mechanism. Shh suppression or overexpression resulted in aberrant proliferation, apoptosis, and differentiation, and thereby formed abnormal feathers (Chang et al. 2004).

In chicken and duck embryonic feather studies, interactions between Sonic hedgehog (Shh) and Bone morphogenetic protein 2 (Bmp2) were suggested to be involved in feather

Figure 7.3. Molecular basis of evolution novelty during feather evolution. (A) Feathers with

different shapes. (B–E) illustration depicting important stages in the formation of modern pen-

naceous feather. (B) opening of the epithelial cylinder via localized apoptosis produces branches.

(C) Anterior-posterior axis forms through a Wnt3a gradient—combination of (B) and (C) converts

a three-dimensional appendage into a two-dimensional, planar structure. (D) Regulating rachis/

barb ratio through BMP activity. (E) Modulation of stem cells to form different morphology along

the distal-proximal-distal axis, which is made temporally from the distal to the proximal end.

Panel A is from Lucas and stetteheim (1972); Panels B–E are from o’Connor et al. (2012).

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branching morphogenesis (Harris et al. 2002). The polarized anterior-posterior expression of Shh and Bmp2 in the primordia of feathers, avian scales, and alligator scales is con-served. This suggests that integrative signaling between Shh and Bmp2 is involved in regulating the growth and differentiation of skin appendages.

In radially symmetric feathers, the stem cell ring is placed horizontally. In bilaterally symmetric feathers, the stem cell ring tilts toward the anterior rachis position (Yue et al. 2005). Molecular gradients were also found in feather follicles. In bilaterally symmetric feathers, a Wnt3a gradient formed from anterior to posterior positions. This gradient does not exist in radially symmetrical feathers (Yue et al. 2006). Flattening the Wnt 3a gradient converted bilaterally symmetric feathers to radially symmetric feathers (Yue et al. 2006). Furthermore, surgically exchanging dermal papillae between radial and bilaterally sym-metric feathers produced chimeric feathers whose phenotype was in accord with the origin of the dermal papilla (Yue et al. 2006).

We next wondered what determines the proximal versus distal feather forms. Ectopic expression of Spry4, a negative regulator of receptor tyrosine kinases, converts proximal regions of the feather follicle to more distal fates. In other words, the region of the collar epithelium abnormally produces barbs. Ectopic barbs can even be seen within the follicle sheath (equivalent to the hair follicle outer root sheath). Feathers which develop have an expanded vane with a greatly diminished calamus (the unbranched, bottom section of a feather). Feathers overexpressing Spry4 have an expanded pulp and reduced or missing dermal papillae. Since epithelial–dermal papilla interactions are essential for feather initia-tion of a new feather cycle, these feathers often cannot regenerate. Ectopic expression of FGF10 has the opposite effect. The dermal papilla is increased in size, and the collar epithelium and adjacent mesenchyme are expanded. Branching morphogenesis and keratin differentiation are inhibited (Yue et al. 2012).

EVOLUTION OF FEATHERS

Diverse Forms in Feathered Dinosaurs

The view of feathers as features unique to birds has recently been overturned by the dis-coveries of many nonavian dinosaurs with feather or feather-like integuments from the Jehol Biota. The Jehol Biota in northeastern China contains Middle-Upper Jurassic and Lower Cretaceous deposits. The frequent volcanic eruptions and widespread distribution of lakes made this area a Mesozoic Pompeii that preserved the imprints of delicate ancient integuments (Zhang et al. 2008b). Here we summarize these findings and show the gradual evolution of new feather forms (table 1; Chuong et al. 2003; O’Connor et al. 2012; Prum 2005; Wu et al. 2004).

Sinosauropteryx prima, a small theropod dinosaur, was first found to have short (∼20 mm on average), slender, and unbranched filamentous structures on the head, neck, back, and both sides of the tail (Chen et al. 1998). The base of the main filaments was hollow, resembling a modern feather’s calamus (Chiappe 2007). Some investigators thought these structures might represent subcutaneous collagen fibers (Lingham-Soliar et al. 2007), but today, most paleontologists acknowledge that these structures were protofeathers (Chiappe 2007). Protofeathers are morphologically different from modern feathers and from the rigid, tubular filaments found on Psittacosaurus and Tianyulong (Mayr et al. 2002; Zheng et al. 2009).

Sinornithosaurus millenii and Beipiaosaurus inexpectus had large patches of filament-like integumentary structures preserved on the forelimbs, hind limbs, and body (Xu et al.

140 ECToDERMAL oRgAn sTEM CELLs

1999a, 1999b). These primitive filaments appeared to be hollow, and some of them showed branched distal ends. Similar protofeathers with simple branching patterns were also found on the tail of a basal tyrannosauroid Dilong paradoxus (Xu et al. 2004), which suggests that protofeathers are not a unique feature of Maniratora and may be present quite early in Coelurosauria. Sinornithosaurus showed the presence of compound structures with multiple joined filaments (Xu et al. 2001). Symmetric vaned feathers (distinct rachis and barbs but no barbules) were discovered on a juvenile Sinornithosaurus. Beipiaosaurus remains show two types of integuments. One is the short slender filamentous protofeathers and the other is the elongated, tubular, unbranched, planar broad filamentous feathers. The latter may have grown from the epidermal collar without barb–interbarb differentiation (Xu and Guo 2009).

Elongated feathers were also reported in Epidexipteryx hui, a new clade at the base of Avialae. This animal exhibited four elongated ribbon-like tail feathers with a central rachis and unbranched vanes which projected out from its short tail. These feathers are thought to have served as ornamental plumage for mate attraction. In addition, this animal also had nonshafted feathers whose distal part is composed of filamentous parallel barbs which unite proximally into an unbranched membranous structure (Zhang et al. 2008a).

The fossils of theropods with modern feather shapes were reported for Protarchae-opteryx robusta and Caudipteryx zoui) (Qiang et al. 1998). Caudipteryx evolved different types of feathers over different body regions, indicating the establishment of feather tracts as an evolutionary novelty. Spectacular pennaceous feathers were found in both the wing (remiges) and tail (rectrices) with tapering shafts (Chuong et al. 2003). Additionally, detailed examination of two juvenile Similicaudipteryx specimens revealed that their rec-trices and remiges are ribbon-like at the proximal part but become fully pennaceous at late juvenile stage. This developmental diversity is not seen in modern birds (Xu et al. 2010). Protarchaeopteryx also has well-preserved vaned feathers close to its tail. The barbs at the proximal region of these feathers are plumulaceous, similar to the aftershaft of modern contour feathers (Chiappe 2007). However, the vaned feathers on Caudipteryx and Pro-tarchaeopteryx are bilaterially symmetric. The lack of bilaterally asymmetric primary remiges indicates poor flight capability and consistent with this idea, the forelimbs of the two dinosaurs are proportionally too small for the animals to have flown (Chiappe 2007; Zhang et al. 2008a).

A slightly more modern type of remige was discovered on Anchiornis huxleyi (Hu et al. 2009). Similar to basal avians, its distal primary remiges have curved rachides. But the size is relatively small and the rachides are thinner. The vanes are still symmetric and blunt at the end. Interestingly, this animal had long pennaceous feathers on its hind limb, a feature only known in Microraptor, the famous four-winged dromaeosaurid dinosaurs.

A Microraptor family member had long narrow feather-like integuments which con-tained rachis-like structures (Xu et al. 2000). The dromaeosaurid “four-winged” dinosaur had long and asymmetric vaned feathers attached to both the forelimbs and hind limbs, making extensive airfoils (Xu et al. 2003). The distribution of feathers across the forelimb also resembles that of modern birds, indicating that this animal may have flown well (Chiappe 2007). Hind wings were also found on another basal Avialae called Pedopenna daohugouensis, yet the wings were smaller and round in shape. The feathers bear the more primitive bilaterally symmetric form (Xu and Zhang 2005).

Recently, the ornithomimosaur (bird mimic dinosaur), a theropod dinosaur, was recov-ered from Upper Cretaceous deposits of Alberta, Canada. It presented filamentous feathers

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covering both juvenile and adult specimens. An adult specimen also possessed a penni-brachium which is a forelimb bearing long feathers (Zelenitsky et al. 2012).

The feather-like structures are not restricted to the small size therapods. A gigantic basal tyrannosauroid, Yutyrannus huali, was found with long filamentous feathers (Xu et al. 2012). This new finding provided direct evidence for the presence of extensively feath-ered gigantic dinosaurs in the Jehol biota. These findings have been summarized (Prum 2005; Prum and Brush 2002; Xu and Guo 2009).

Beside the imprints of primitive feathers in fossil dinosaurs, amber containing Mesozoic feathers was also found recently in Canada. This amber provides an excellent chance to observe microscopic feather structures. Upon careful examination, several different feather morphologies were discovered from the most primitive hollow singular filaments to rachis containing feathers with barbules specialized for discrete functions (McKellar et al. 2011).

Evo-Devo of Feather Forms

Here we combine the cellular/molecular laboratory work described in the “Feather Branch-ing Morphogenesis” section with the diverse proto-feather fossil forms described in the “Diverse Forms in Feathered Dinosaurs” section to produce the model in Figure 7.3, which was first published in O’Connor et al. (2012).

Process 1. Opening the epithelial cylinder via localized apoptosis (Figure 7.3B). Here apoptosis (i.e., programmed cell death) of the feather sheath and pulp epithelium allows the feather cylinder to open and to become a two-dimensional epithelial plane. The apop-tosis of marginal and barbule plate epithelia leads to the formation of space between barbs and barbules, respectively (Chang et al. 2004). It is possible that in the Mesozoic birds, these processes may be uncoupled (i.e., apoptosis occurs in the feather sheath and pulp epithelium without barb ridge formation) and thus potentially form the undivided sheet-like structure observed in the proximal rectrices in some enantiornithines and confuciusor-nithiforms (O’Connor et al. 2012).

Process 2. Forming an anterior (location of rachis)—posterior axis by a Wnt 3a gradient (Figure 7.3C). Feather stem cells lie horizontally at the base of a downy feather follicle but tilt downward toward the anterior side of a flight feather follicle (Yue et al. 2005). Wnt 3a protein has an anterior-posterior gradient in flight, but not in downy feathers. Global inhibition of the Wnt 3a gradient transforms bilaterally symmetric feathers into radially symmetric feathers (Yue et al. 2006). Combining processes 1 and 2 enables the radially arranged barbs in downy feathers to insert into a rachis with bilateral symmetry, thus converting a three-dimensional appendage into a two-dimensional plane.

Process 3. Regulating the rachis/barb ratio with BMP activity (Figure 7.3D). Once the periodic barb ridge formation process is triggered, cells in the epithelial cylinder will become either rachid, barbs, or the space (following apoptosis) between barb branches (Yu et al. 2002). Feather morphogenesis results from activator–inhibitor interactions that occur between the bone morphogenetic proteins (BMP2 and BMP4), noggin, Sonic hedgehog (Shh), and other molecules, which form a hierarchical epithelial branching pattern leading to the diverse feather morphologies seen in living birds (Harris et al. 2002; Yu et al. 2002). Thus, modifying the expression of these molecules can produce primitive or novel mor-phologies. For example, changes in BMP activities can result in numerous barb ridges or an enlarged shaft, as seen in the elongate rectrices of enantiornithines and confuciusor-nithiforms (O’Connor et al. 2012).

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Process 4. Modulating stem cells to form different morphologies along the proximal-distal axis (Figure 7.3E). Stem cells were found to be located in the proximal follicle. They proliferate to generate new cells which are pushed upward and differentiate as they move toward the distal end (Lucas and Stettenheim 1972). Thus, the distal end represents more differentiated structures that were formed earlier. Since the micro-environment for stem cells can change at different times, it is possible to generate different structures during the course of feather development, enhancing the diversity of feather morphologies through changes in the signaling environment. For example, overexpression of sprouty genes leads to overt branching, while overexpression of Fgf 10 leads to collar-like proliferative and undifferentiating epithelia (Yue et al. 2012). With this understanding, it is possible to envisage that the unique morphology of a feather can be shaped by incremental changes of the micro-environment for feather stem cells.

HAIR REGENERATION: POPULATION BEHAVIOR IN REGENERATION

In a way analogous to feathers, hairs also cycle through periods of growth (anagen), invo-lution (catagen), and rest (telogen) (Figure 7.4A). The hair cycle can then reenter anagen to begin a new cycle. Hair follicle stem cells within a specialized niche (the hair bulge) must be activated in order for the hair cycle to renew. Once activated, the stem cells pro-liferate to generate transient amplifying cells and a new progenitor cell (Hsu et al. 2011). In this way, they can maintain their stem cell population throughout the life of the organism.

New Hair Cycle Phases Due to Micro- and Macro-Environmental Interaction for Stem Cells

Activation of progenitor cells is regulated by factors localized within the hair follicle. This micro-environment is known to contain Wnt signaling among other regulatory factors. Surprisingly, we found that hair follicles not only respond to micro-environmental signals but can communicate between themselves using macro-environmental (extrafollicular) communication.

This results in the ability of hair follicle populations to cycle in waves. We can deter-mine the hair cycle stage by shaving the mice. Using a black mouse, anagen skin produces pigment which can be easily observed, while skin in telogen is pink. Using this approach, we traced the hair wave as it progressed across the skin for 1 year. The waves move in a coordinated fashion caudally. Initially, the wave progresses as a single domain. As a mouse ages, the domain fragments into multiple subdomains. We found that the wave traverses the skin with sharp boundaries. The boundaries are formed by refractory regions that cannot be induced to reenter anagen. (Figure 7.4C).

Our study enabled us to identify four hair follicle stages during mouse hair regenera-tion. During anagen, there is a signal-propagating (P) phase where competent neighboring follicles prompt can be induced to enter anagen. Next is an autonomous (A) phase where neighboring follicles cannot be stimulated.

Plikus et al. found that the periodic expression of dermal bone morphogenetic protein 2 (Bmp2) and Bmp4 inhibited progression of the hair wave (Plikus et al. 2008). Furthermore, in comparing the timing of Wnt and BMP expression, they found that BMP expression was out of phase with that of WNT/beta-catenin. Since Wnt/beta-catenin stimu-lates hair follicle progression to anagen and BMP inhibits this event, this explained how telogen could now be divided into refractory and competent phases of hair regeneration.

HAiR REgEnERATion: PoPuLATion BEHAvioR in REgEnERATion 143

Figure 7.4. Regenerative hair waves of mice. (A) The hair cycle includes anagen (early and late),

catagen, and telogen phases. (B) Model of macro- and micro-environment-mediated activation

of hair follicle stem cells (from supplement of Plikus et al. 2011). (1) in the R-phase of telogen,

WnT antagonists, such as Dkk1 and sfrp4, and inhibitory Bmp2/4 ligands collectively keep hair

follicle stem cells in a quiescent state; (2) Loss of these WnT antagonists from the macro-envi-

ronment in the C-phase of telogen enables spontaneous, low-frequency WnT activity in the

dermal papillae; (3) The stochastic activation of large groups of WnT positive dermal papillae is

necessary to stimulate adjacent stem cells and to induce spontaneous anagen reentry events (4

and 5 on panel B). (C) Progression of regenerative hair waves is affected by the ratio of activators

and inhibitors in the dermal macro-environment. Panels B and C are from Plikus et al. (2011). A,

autonomous anagen; C, competent telogen; P, propagatory anagen; R, refractory telogen.

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The competent (C) telogen phase had low BMP signaling while the refractory (R) telogen phase had high BMP expression in the macro-environment, which kept the hair follicle stem cells in a quiescent state and blocked progression toward anagen reentry.

This novel system helps to maintain stem cell homeostasis. It also enables these pro-genitors to respond to the sum of all micro-environmental and macro-environmental signals impinging upon them (Figure 7.4B). So hair follicles can respond to macro-environmental signals to enter anagen during the (P) phase in which case the hair follicles initiate from a center and subsequently propagate. Hair follicles also can respond to micro-environmental signals to enter anagen during the (A) phase and initiate at random locations. This observation supports our understanding that hair follicle activation occurs in a stochastic manner (Plikus et al. 2011).

Activator/Inhibitor Modulation

Manipulating levels of BMP signaling can alter both the length of (R) and the speed of propagation during the regenerative wave. BMP signaling was inhibited using KRT14-noggin transgenic mice which express Noggin, a BMP inhibitor, in the basal epithelium. These mice had a shortened refractory telogen. In contrast, KRT14-Wnt 7a mice that have elevated Wnt activity showed a propensity toward hair follicle propagation (Plikus et al. 2011). This nicely demonstrates that hair follicles can integrate micro-environmental and macro-environmental stimuli to regulate stem cell homeostasis (Figure 7.4C). The Wnt inhibitors, DKK and sfrp4, were also shown to act as macro-environmental inhibitors (Plikus et al. 2011), while PDGF acts as a macro-environmental activator of anagen reentry (Festa et al. 2011).

REGENERATIVE HAIR WAVES IN TRANSGENIC MICE AND DIFFERENT MAMMALIAN SPECIES

Wave Patterns in Different Animals

In mouse skin, hair cycling is patterned and appears coordinated. Hair cycling patterns are complex and change over time. However, we observed the hair wave of a laboratory mouse housed under constant environmental conditions without seasonal variation. To examine whether the hair wave pattern in mice also exists in wild animals, we observed three species of wild rodents which were kept in a wild environment with natural temperatures and day–night cycles during three seasons (Spring, Summer, and Fall). We observed the Mongolian gerbil (Meriones unguiculatus), hamster, and chipmunk for 353 days, 57 days, and 33 days, respectively (Figure 7.5). Similar to lab mice, wild Mongolian gerbils (Figure 7.5A) and hamsters (Figure 7.5B) show hair waves. The Mongolian gerbil hair waves change slowly with advancing age. The hamster hair wave initially has almost equal hairs in anagen and telogen, and the wave becomes disordered after several waves. The chip-munk showed a very rapid change in pattern, but the hair wave was not clear (Figure 7.5C). These versatile hair regenerative patterns may help animals adapt to different physi-ological conditions.

Compared with mice, rabbits have more robust hair growth. The skin surface area of rabbits is 30 times larger than that of mice. Rabbits have compound hair follicles, with each follicle containing multiple bulge stem cell clusters, hair germ cells, and dermal papillae, which are activated in a time sequence. This configuration increases the ability

REgEnERATivE HAiR WAvEs in TRAnsgEniC MiCE AnD DiFFEREnT MAMMALiAn sPECiEs 145

Figure 7.5. Regenerative hair waves in three wild rodents. (A) Mongolia gerbils. in adult gerbils,

the anagen period is very variable. in dorsal region, the anagen period could last 3–45 days in

winter and 7–31 days in spring. (B) Hamster. The hamster hair wave initially has almost equal

time in anagen and telogen. Both of them last 8–12 days. The wave becomes disordered after

several cycles. (C) Chipmunk. Two-month-old chipmunk starts to show five black stripes. When a

two-month-old animal was shaved, a 15-day anagen period was observed, followed by a 30-day

telogen period. The hair wave patterns in adult chipmunks become complex. They appear in the

regenerating wave and domain, similar to lab mice. Adult chipmunk underwent one to two

anagen period then enter hibernation. The hibernation period could last 6–8 months. The hair

in the hibernation period is in telogen phase.

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of follicles to initiate and propagate the hair wave, leading to a fractal-type wave pattern (Figure 7.6A) (Plikus et al. 2011).

Regenerative Behavior in a Large Population of Stem Cells

During hair follicle evolution, epithelial stem cells underwent topological clustering where all stem cells within a cluster either remain quiescent or become activated relatively

Figure 7.6. The spectrum of hair regenerative patterns and the management of the large stem

cell population. (A) The spectrum of hair regenerative patterns. The x-axis is the probability of

coupled activation, whereas the y-axis is the probability of intrinsic activation. (B) The topology

and hierarchical management of the large stem cell population. The x-axis is the probability of

coupled hair follicle regeneration, while the y-axis is the strength of intrinsic stem cell activation.

Different animals or different physiological conditions in the same animal can significantly alter

the global dynamics of hair regeneration by modulating the x- and y-axes (from Plikus et al. 2011).

REFEREnCEs 147

synchronously. This allows for the conversion from the continuous renewal mode observed in epidermis to the episodic regeneration mode observed in the hair cycle.

Stem cell clustering and episodic hair cycling represent evolutionary novelties and enable new ways for the large-scale coordination of regeneration. Each cluster of hair stem cells can become activated by the intrinsic hair cycle clock (Figure 7.6A, y-axis). Intrinsic activation alone can ensure sufficient levels of hair regeneration if it can occur with a high probability. This is seen in the adult human scalp, where hair follicles regenerate autono-mously based on intrinsic activation (Figure 7.6A, y-axis). Hair regeneration based on this mechanism alone can become easily deficient when the probability of intrinsic activation drops, such as upon alopecia. Additionally, this mechanism does not allow for any coor-dination of regeneration among neighboring hair follicles. Diffusible signaling molecules used for regulating hair stem cell activities within each hair follicle can be co-opted to mediate interactions between neighboring hair follicles. Such signaling couples activation events among many stem cell clusters at once (Figure 7.6A, x-axis). By modulating the strength of intrinsic stem cell activation (Figure 7.6A, y-axis) and the probability of coupled activation (Figure 7.6A, x-axis), different animals or different physiological condi-tions in the same animal can significantly alter the global dynamics of hair regeneration. As a result, versatile hair growth patterns in rabbits, mice, wild rodents, normal, and alo-pecic human scalps can be all be explained within the same patterning framework, which is based simply on how hair stem cell activities are “managed.” Stem cell clusters can now be regulated as one entity allowing organ regeneration to occur episodically with an intrinsic rate (Figure 7.6B).

Thus, we can appreciate ectodermal organ stem cells are highly plastic. They are regulated by the micro-environment and macro-environment. The micro-environment helps regulate feather regeneration and morphogenesis. The macro-environment mediates the effect of the outside environment (e.g., season) or body physiological conditions to modulate the micro-environment (Chen and Chuong 2012). During evolution, ectodermal organ stem cells show high adaptability to accommodate changes in the ecological envi-ronment over a large timescale (Figure 7.1E) to help species survive and flourish.

ACKNOWLEDGMENTS

This work was supported by grants from US NIH AR42177, AR47364, and AR 060306. J. Y. is supported by China NSFC 30960244.

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