Structural and Immunocytochemical Characterization of Keratinization in Vertebrate Epidermis and Epidermal Derivatives

Structural and Immunocytochemical Characterization of Keratinization in Vertebrate Epidermis and Epidermal Derivatives Lorenzo Alibardi Department of Experimental and Evolutionary Biology, University of Bologna, 40126 Bologna, Italy

This review presents comparative aspects of epidermal keratinization in vertebrates, with emphasis on the evolution of the stratum corneum in land vertebrates. The epidermis of fish does not contain proteins connected with interkeratin matrix and corneous cell envelope formation. Mucus‐like material glues loose keratin filaments. In amphibians a cell corneous envelope forms but matrix proteins, aside from mucus/glycoproteins, are scarce or absent. In reptiles, birds, and mammals specific proteins associated with keratin become relevant for the production of a resistant corneous layer. In reptiles some matrix, histidine‐rich and sulfur‐rich corneous cell envelope proteins are produced in the soft epidermis. In avian soft epidermis low levels of matrix and cornified proteins are present while lipids become abundant. In mammalian keratinocytes, interkeratin proteins, cornified cell envelope proteins, and transglutaminase are present. Topographically localized areas of dermal–epidermal interactions in amniote skin determine the formation of skin derivatives such as scales, feathers, and hairs. New types of keratin and associated proteins are produced in these derivatives. In reptiles and birds b‐keratins form the hard corneous material of scales, claws, beaks, and feathers. In mammals, small sulfur‐rich and glycine–tyrosine‐rich proteins form the corneous material of hairs, horns, hooves, and claws. Molecular studies on reptilian b‐keratins show they are glycine‐rich proteins. They have C‐ and N‐terminal amino acid regions homologous to those of mammalian proteins and a central core with homology to avian scale/feather keratins. These findings suggest that ancient reptiles already possessed some common genes that later diversified to produce some keratin‐associated protein in extant reptiles

International Review of Cytology, Vol. 253 Copyright 2006, Elsevier Inc. All rights reserved.

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0074-7696/06 $35.00 DOI: 10.1016/S0074-7696(06)53005-0

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and birds, and others in mammals. The evolution of these small proteins represents the more recent variation of the process of cornification in vertebrates. KEY WORDS: Vertebrates, Epidermis, Differentiation, Cytology, Immunocytochemistry, Immunoblotting, Proteins, Evolution. ß 2006 Elsevier Inc.

I. Introduction The present review deals with the process of keratinization in vertebrate epidermis and summarizes common aspects of this process from fish to amniotes. The focus of this review is on the cytological and comparative process of cornification of keratinocytes. Other cell types of the epidermis are not considered in the present review (see Chuong et al., 2002, for references on other functions of the epidermis). After the Introduction (Section I), the theme of the present review starts with the general processes of epidermal diVerentiation, illustrating the diVerence between keratinization and cornification (Section II). Cells of the special epithelium that constitute the epidermis (termed keratinocytes) move from the basal stratum to the external layers and eventually die: during this movement, followed by the production of new cells underneath and by the loss of superficial cells, a process of diVerentiation occurs in keratinocytes. DiVerent genes are activated and keratinocytes produce new proteins that are associated with the soft keratin network (Byrne et al., 2003; Fuchs, 1990; Kalinin et al., 2002; Resing and Dale, 1991; Steinert, 1998; Wu et al., 2004). The process of progressive increase in the amount of keratin filaments, isolated or associated as loose bundles, is termed keratinization and is common to most of the simple or stratified epithelia. In the epidermis, because of the particularly high mechanical and protective demands, keratin filaments are more abundant than in other epithelia, thus producing strong keratinization. In addition, glycoprotein, mucus, and complex lipids are produced in the upper intermediate cells of the epidermis (Elias et al., 1987; Matoltsy, 1987; Mittal and Banerjee, 1978; Rawlings et al., 1994). Terminal diVerentiation of keratinocytes determines nuclear loss and degradation of nuclear material that is mixed with the cytoplasmic components. When keratin remains the main component, and appears as single visible electron‐dense filaments, this type of terminal diVerentiation is still termed keratinization. This process is prevalent in the epidermis of fish and amphibians (Fox, 1986; Parakkal and Alexander, 1972; Warburg et al., 1996; Whitear, 1986).

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The morphology of the epidermis in the various vertebrates, from fish to mammals, and the specific organelles of keratinocytes, cytokeratins, and associated proteins are presented in Section II. In Section III of this review, the cytology of keratinocytes and of their proteins is discussed. In diVerentiating keratinocytes in amniotes various degrees of biochemical alteration occur in the keratin material during terminal diVerentiation. Keratin filaments are degraded and an increase in the quantity of interfilamentous, keratin‐associated proteins and of complex lipids occurs. Molecules are chemically bonded one to another, forming a dense, more or less homogeneous corneous mass: this process is termed cornification and produces a tougher and less pliable mass with respect to that produced by the process of keratinization. The granular layer in the mammalian epidermis represents the site of production of a histidine‐rich protein (filaggrin) that forms keratohyalin, and determines aggregation of keratin filaments (Fukuyama and Epstein, 1986; Holbrook, 1989). A granular layer is not seen in the epidermis of other vertebrates. Keratohyalin with mammalian characteristics (e.g., containing histidine‐rich proteins) is absent in the epidermis of fish, amphibians, most reptiles, and birds. This suggests that keratinization takes place without keratohyalin in the other vertebrates. In the corneous mass, the altered keratin filaments are more or less visible as electron‐pale filaments, 8–10 nm in diameter, surrounded by an electron‐ dense matrix: this is termed the ‘‘a‐keratin pattern’’ (Baden and Maderson, 1970; Fraser et al., 1972). The a pattern also produces a typical diVraction pattern using X‐rays, and has been found in the epidermis of all vertebrates. DiVerent forms of hard keratins or associated proteins have been produced in amniote epidermis, which produce an alteration in the X‐ray pattern of a‐keratin. The hard form of cornification occurs in scales, feathers, and hairs. Because of specific dermal–epidermal interactions, which begin during embryogenesis, specific skin derivatives are formed, among which are scales, feathers, and hairs: these three derivatives, which characterize, respectively, reptiles, birds, and mammals, are discussed in Section IV. Areas of dermal– epidermal interactions determine the production of specific epidermal proteins, among which are ‘‘hard keratins’’ and ‘‘keratin‐associated proteins.’’ In hard epidermal derivatives of mammals, such as hairs, horns, nails, and so on, an a pattern has been found. In sauropsids (reptiles and birds), large regions of scales and epidermal derivatives, including feathers, show a diVerent X‐ray pattern, termed the ‘‘b‐keratin pattern.’’ In this case ultrastructural examination shows more or less distinguishable, electron‐pale 3‐ to 4‐nm keratin fibrils present among scarce and denser matrix material. This matrix produces a hard, extremely resistant and inflexible corneous material made of ‘‘b‐keratin.’’ Therefore, it has been considered that reptilian and avian epidermis produces two major groups of keratin: a‐ and b‐keratins (Baden

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and Maderson, 1970; Fraser et al., 1972; Fraser and Parry, 1996; Maderson and Alibardi, 2000). The molecular nature of hard keratins in amniotes is compared in Section V to illustrate possible pathways followed by evolution in selecting these proteins to build the resistant skin derivatives in amniotes (horns, scales, beak, etc.). Finally, some trends and themes of research in comparative dermatology are indicated in the conclusions (Section V).

II. Comparative Cytology of Vertebrate Epidermis: From Keratinization to Cornification To follow the evolution of epidermal mechanisms in vertebrates we must deal first with the various structures of the epidermis in vertebrates, that is, cells, organelles, and their constituent molecules (keratins, associated proteins, lipids, and enzymes), as presented in the following sections. An overview of the histological and a‐keratin immunocytochemical patterns in the epidermis of various vertebrates is presented in Figs. 1 and 2.

A. Fish Epidermis: Adaptation to Water The epidermis in an aquatic environment presents less mechanical demand than in the terrestrial environment, because of the fluidity. The external epithelium covering the body of this first group of anamniotes therefore needs a strong keratin cytoskeleton and cell junctions to ensure cell cohesiveness under streamlined conditions. As a result the epidermis of most fish resembles a mucous epithelium, which instead covers the inner cavities of body organs in terrestrial vertebrates. At the same time, moving within a relatively dense fluid, mechanisms for sliding (cuticle, slimy surface) must be present at the surface of an exposed epithelium, otherwise superficial cells would be constantly damaged and removed by the movement of water currents. The histology, cytology, ultrastructure, and immunocytochemistry of the skin in fish, including keratinocytes and other cell types, have been extensively revised (Mittal and Banerjee, 1978; Sire and Huysseune, 2003; Whitear, 1986; Zaccone et al., 2001). The epidermis of actinopterygian fish is a variably stratified epithelium (Fig. 1A–C) in which cells can divide in all layers and are continuously lost at the surface, covered by a noncellular cuticle. Also, the skin of sarcopterygian fish is composed of a multistratified epithelium over a variably complex dermis. Superficial cells remain viable, little keratinized, no keratohyalin is formed in outer layers, and a mucous cuticle is produced on the surface (Fig. 3A). Corneous structures are limited to small areas in a few species.

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FIG. 1 Distribution of cytokeratins in vertebrate epidermis, using AE1 or AE3 antibodies. (A and B) Goldfish (Carassius carassius). Scale bar: 20 mm. (C) Lungfish (Neoceratodus forsteri). Scale bar: 10 mm. (D and E) Frog (Rana esculenta). Scale bar: 10 mm. (F) Turtle limb (Chrysemys picta). Scale bar: 10 mm. (G) Turtle carapace (C. picta). Scale bar: 10 mm. (H) Alligator (Alligator mississippiensis). Scale bar: 10 mm. (I) Lizard (Podarcis sicula). Scale bar: 30 mm. c, corneous layer; g, gland; h, hinge region. Arrows indicate the thin corneous layer; dashes outline the basal layer.

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FIG. 2 Distribution of cytokeratins in vertebrate epidermis, as determined with AE1 or AE3 antibodies. (A) Lizard (Podarcis sicula). Scale bar: 10 mm. (B and C) Tuatara (Sphenodon punctatus). Scale bar: 10 mm. (D) Ostrich scale (Struthio camelus). Scale bar: 10 mm. (E) Emu (Dromaius novaehollandiae). Scale bar: 10 mm. (F) Echidna (Tachyglossus aculeatus). Scale bar: 10 mm. (G) Platypus (Ornithorhynchus anatinus). Scale bar: 20 mm. (H) Rat (Rattus norvegicus). Scale bar: 10 mm. (I) Ultrastructural detail of labeled keratin bundles converging to a desmosome in a keratinocyte of lizard epidermis (Podarcis muralis). Scale bar: 100 nm. c, corneous layer; ds, desmosome; k, keratin bundles; s, suprabasal layers. Arrows indicate the thin corneous layer; arrowheads indicate the basal layer; dashes outline the basal layer.

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FIG. 3 Ultrastructural features of diVerentiative organelles of vertebrate epidermis. (A) Irregular mucus granules (arrows) and vesicles (v) discharged into the cuticle surface (cu) of lungfish (N. forsteri). Scale bar: 200 nm. (B) Dense small granule (arrowhead) near keratin bundle (k) in spinosus keratinocytes of toad (Bufo viridis). Scale bar: 100 nm. (C) Vesicular bodies (arrows) in precorneous (sebo‐)keratinocyte of zebra finch (Taeniopygia castanotis). Scale bar: 200 nm. (D) Lamellar body (arrowhead points to the lamellae) of upper spinosus keratinocyte of echidna (T. aculeatus). Scale bar: 100 nm. (E) Mesos/lamellar body (arrowhead points to the lamellae) of a‐keratinocytes in the tuatara (S. punctatus). Scale bar: 100 nm. (F) Detail of cornified cell membrane (arrow) in superficial corneous cells of toad (B. viridis). Scale bar: 200 nm. (G) Detail of loricrin immunolabeling (LOR) of keratohyalin (kh) in keratinocyte of the stratum granulosum of humans (Homo sapiens). Scale bar: 100 nm.

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The extracellular cuticle, together with superficial microridges, is the only indication of basoapical polarity in fish epidermis for the production of a specialized superficial structure. The modality of cell death seems to occur by a necrotic process due to metabolic wearing of cells, and no program for cell death has so far been demonstrated. Surface cells of many fish show microvillous structures sustained by an actin microfilamentous network on the surface of the epidermis. A three‐ dimensional study has shown that microvilli are in reality microridge structures with diVerent patterns (Bereither‐Hahn, 1979; Whitear, 1986). The plasma membrane of keratinocytes is generally not thickened as in terrestrial vertebrates, and deposition of dense material occurs in the outer plasma membrane of some specialized epidermal areas (Mittal and Banerjee, 1978; Mittal and Whitear, 1979). The proteins involved in the formation of cornified membrane in fish epidermis remain unknown. Fish mucus is produced mainly for secretion and covers the epidermal surface, where it forms an extracellular cuticle (Matoltsy, 1987) (Fig. 3A). The latter eases friction during swimming and functions as a protective barrier. The cuticle is composed of mucous, glycoprotein, and proteoglycan secretions from single cells, keratinocytes, and goblet cells (Lebedeva, 1999; Mittal and Banerjee, 1978; Zaccone et al., 2001). Mucus is a bactericide because of its content of C‐reactive proteases, antibacterial antibodies, hemoagglutinins, lysozyme‐like proteins, cholinesterases, and so on (Lebedeva, 1999; Zaccone et al., 2001). Some proteins of the cuticle possess a rich moiety of proline and serine, but their specific amino acid sequence is not known. Proteins linked to the activation or sequestration of calcium (e.g., calmodulin) have been reported. Enzymes such as nitric oxide synthase, glycosidases, alkaline and acid phosphatases, and succinic dehydrogenases indicate that fish epidermis is a living and respiratory epidermis. The presence of transglutaminases has been detected (Alibardi, 2002a; Rice et al., 1994). The latter enzyme may intervene in some cross‐linking of cuticle proteins to produce a more compact protein texture of the cuticle. The function of extracellular mucus is multiple, but in general it is a carrier of ions for osmoregulation, and contains bioactive molecules such as antibacterial molecules, agglutinins, lectins, bioactive agents, and also toxins. Mucus regulates ion and water movement through the epidermis, especially that of Naþ and Kþ, and it is the storage medium for alarm molecules and pheromones. Neither the function of intracellular mucus in the aggregation of keratin filaments nor its role in the mechanical protection of the epidermis seems to occur in fish epidermis. Keratinocytes contain keratin (intermediate) filaments but, except in specialized body regions in some species, no dense bundles or corneous material is produced. Most of the cytoplasm of superficial keratinocytes contains numerous isolated keratin filaments, generally not aggregated into a dense mass.

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A diVerent process has been described in the breeding tubercles of the fish Bagarius bagarius (Mittal and Banerjee, 1978; Mittal and Whitear, 1979), and in other horny formations of the epidermis of other fish. In the latter, a corneous cell envelope and dense keratin bundles are formed. The above examples show the potential of fish epidermis to form corneous material although little is known about the molecules involved in this process.

B. Amphibian Epidermis: Transition from Aquatic to Terrestrial Environment The histology, cytology, immunohistochemistry, and ultrastructure of the epidermis in amphibians (tetrapods or terrestrial anamniotes) have been extensively studied (Budtz, 1977; Fox, 1986, 1994; Warburg et al., 1996; Whitear, 1977). In the aquatic stage, the epidermis of amphibians presents characteristics similar to those of fish, adapted to streamline but incapable of oVering much mechanical protection from abrasion and other mechanical stress. The epidermis in this phase remains stratified and devoid of a corneous layer, like that of fish. This condition remains a permanent trait mainly in perennial aquatic forms, such as Amphiuma and Proteus. In terrestrial‐adapted species, during and after metamorphosis, the structure of the epidermis changes and a true (although generally monolayered) corneous layer appears (Fig. 1D and E). This determines the periodic shedding of the superficial, nonelastic corneous layer in relation to body growth. Molting allows the replacement of the dense corneous layer with a new, less cornified and little expandable corneous layer that maintains the oxygen exchange for cutaneous respiration. The latter would be impeded if more than two layers of corneocytes were accumulated on the surface of the epidermis. Molting has been divided into four stages: intermolt stage, preparative stage, early shedding, and late shedding (Budtz, 1977). The basal layer contains most dividing cells, but cell division also occurs in suprabasal layers. Mucus‐like granules start to accumulate in the upper stratum intermedium. Two types of granule have been described: large and small (Fig. 3B). Large granules remain intracellular, whereas small granules are extruded extracellularly. Mucus and glycoconjugates coat the surface of corneous and precorneous (replacement) layers, and protect the epidermis from desiccation in the terrestrial environment and from excessive water intake in aquatic species. Aside from mucous cells in the epidermis, the superficial mucous coat derives from dermal mucous and serous glands. Together with mucus, small granules contain glycoproteins, acid glycosaminoglycans, tyrosinase, and other enzymes. Glycoconjugates containing fucose‐N‐acetylglucosamine or galactosamine‐galactose have been found in

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the dorsal epidermis of frogs (Faszenski and Kaltenbach, 1995). This asymmetric distribution has been correlated with water intake (ventral skin) and loss (dorsal skin). In some species of anurans, highly keratinized cells (palisade) link basal with precorneous cells, and may act as holding points to the connective tissue, improving the mechanical resistance of the epidermis again stress. The more superficial layers include the precorneous and corneous layers, the latter generally forming a mono‐ or bilayer. Thickening of the plasma membrane occurs in corneous cells, and in some cytoplasmic areas keratin bundles are densely aggregated (Alibardi, 2001b, 2002a). The protein content of these modified corneous cell envelope is scarcely known, but some studies have shown cross‐reactivity for some cell corneous envelope proteins (Alibardi and Toni, 2004a,b). Dense material of unknown composition is deposited against the plasma membrane, which becomes thicker (Fig. 3F). Horny structures, as in fish epidermis, occurs only in some areas, but no data are available on their molecular composition. This occurs in keratinocytes of the beak of anuran larvae, and also in cocoons of estivating anurans. In the latter more than 40–50 layers of corneocytes are formed on the surface of the epidermis, an extraordinary case of stratum corneum stratification in amphibians (Fox, 1986; Pough et al., 2001). Another case is the disk‐like pads of urodeles and anurans (Fox, 1986). These organs are made by five to seven layers of epidermal cells, with the outermost cells forming apical fingers occupied by dense keratin bundles with axial orientation, among which dense granules are present. In addition, the nuptial pads of males of some anurans, which are temporary organs developed during the breeding seasons, are strongly keratinized structures. The outer six to eight cell layers organize into conical organs that accumulate dense bundles of keratin, among which dense granules are present. Therefore, like in fish, the epidermis of extant amphibians also has some potential for true cornification.

C. Reptilian Epidermis: Keratinization versus Cornification for a Terrestrial Ectotherm Lifestyle An intense process of cornification and extracellular lipidization is common only in reptiles, the first amniotes (Pough et al., 2001). This class represents a polyphyletic group, which is traditionally assembled into a single group. The skin of extant reptiles is generally scaled or presents a rough surface in nonscaled areas (e.g., in tortoises and turtles). Scales form a resistant but coarse surface that varies from an inflexible coat (crocodilians, many chelonians, and lepidosaurians) to a pliable and softer surface (in many snakes, and in lizards such as geckos).

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The histology, cytology, histochemistry, and ultrastructure of the epidermis of the diVerent reptilian groups have been extensively studied (Alibardi, 2003a; Alexander, 1970; Banerjee and Mittal, 1978; Landmann, 1986; Maderson, 1985; Maderson et al., 1972, 1998; Spearman, 1966). Compared with anamniotes, the corneous layer in reptiles becomes multilayered, but its thickness varies in diVerent species, body areas, histological structure, and biochemical composition. A more or less continuous desquamation occurs in the corneous layer of chelonian and crocodilian epidermis (Fig. 1F–H), whereas in lepidosaurian reptiles (lizards and snakes) a periodic shedding takes place (Figs. 1I and 2A–C).

1. Chelonians Two main types of cornification occur: soft (limbs, tail, and neck) and hard (shell and some scales). In the waved soft epidermis (limbs, tail, and neck) of chelonians, a smooth surface or a rough surface is present. In the latter, scales merely represent folds of the skin, and a regional diVerentiation along their surface is not present. The few layers of keratinocytes are covered by a thick, multilayered corneous layer (Fig. 1F and G). The basal layer varies from flat to columnar, probably in relation to the proliferative and synthetic activity during the year. Cytological and biochemical studies have shown that no keratohyalin‐like granules are present in the intermediate or transitional layer (Alibardi, 2003a; Alibardi et al., 2004b; Matoltsy, 1987). Keratin filaments are therefore aggregated in parakeratotic corneocytes. In addition, mucous granules and vesicular bodies, similar to those of avian keratinocytes (Fig. 3C), are produced in precorneous keratinocytes of the soft epidermis of chelonians. In the shell (carapace and plastron), and in hard scales of limbs of tortoises, b‐keratin is deposited or completely replaces a‐keratin, with the exception of the narrow hinge regions between scales. Scales or scutes (this term is most often used for shell scutes) do not overlap much and the dorsal surface is delimited by narrow hinge regions. The basal layer is cubic to columnar, and a monolayered suprabasal layer is present during resting seasons (in temperate species in fall and winter). More suprabasal layers are produced during shell growth (Alibardi, 2005a). b Cells maintain irregular boundaries within the newly generated corneous layer. In the more superficial corneous layer boundaries largely disappear as cells become more and more fused to form a compact layer. Many corneous cells are relatively large and thick (over 0.5 mm) at maturity. The latter is consumed by continuous wearing (many terrestrial and marine chelonians) or by periodic desquamation of the superficial horny layer of scutes. In the latter case, shedding layers are formed.

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b Cells form a resistant but thick corneous layer, which may also be heavy, especially when calcified (Alibardi and Thompson, 1999; Spearman, 1966): the resulting skin is dry, tough, and unpliable. Dense granules that mix with b‐keratin filaments are produced in the precorneous layers in some periods of shell growth but their chemical nature is unknown. b‐Keratin is also present in the soft epidermis of hard‐shelled species (Chrysemys, Testudo, and Pelomedusa) and also in the shell of a soft‐shelled species (Trionyx) (Alibardi et al., 2004b; Baden and Maderson, 1970; Wyld and Brush, 1979, 1983). 2. Lepidosaurians Scales in lepidosaurian reptiles are numerous in shape and degree of overlapping, from the tuberculate scales of some geckos or agamids, with almost no overlapping, to the extremely overlapped scales of snakes and scincids (Lillywhite and Maderson, 1982). A unique characteristic of lizards and snakes, among reptiles, is the periodic shedding of their multilayered epidermis (Maderson, 1985; Maderson et al., 1998). In the epidermis of most vertebrates, from the basal layer a single type of keratinocyte is generated and diVerentiates into a‐keratinocytes. In lizards and snakes, the basal layer instead produces diVerent types of keratinocytes, and they accumulate b‐ or a‐keratin and becomes cornified with diVerent characteristics. The first layer to be formed is called oberhautchen, and it is progressively followed by the b layer, mesos layer (or region), and a layer. A proper a layer is further distinguished into a lacunar layer and a clear layer: the latter two layers are evident only at the late renewal stage. Present day studies indicate that it is the amount and type of hard keratin (b‐keratin) deposited in these diVerent layers that determine their specific cytodiVerentiation (see below). The scaly epidermis of lizards and snakes is made of diVerent layers produced during a renewal phase underneath the old epidermis (outer epidermal generation; see Landmann, 1979, 1986; Maderson, 1985; Maderson et al., 1998) (see Figs. 1I and 2A–C). The latter is not completely mature until the new epidermal layers (inner epidermal generation) are produced underneath. The first suprabasal cells produced from the basal cubic to columnar epidermis diVerentiate into cells forming the oberhautchen layer. This layer produces spinules or surface microornamentation that form the typical, species‐specific, outer sculpturing of scales (Irish et al., 1981; Maderson et al., 1998; Price, 1982). In the special climbing scales of some geckos and anolid lizards extremely long protrusions (setae) of oberhautchen cells may extend over 60–70 mm in length (Alibardi, 2003b; Maderson, 1970). The second layer produced is made by a stratification of b‐keratin‐synthesizing cells that merge into a more or less complete syncytium, the b layer. b‐Layer cells become thin at maturity. The third layer is formed by cells that do not

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produce b‐keratin, and is termed the mesos layer or mesos region as it represents a transition to the following a layer. Mesos cells are extremely thin and contain lipids that are stored mainly in lamellate or mesos granules, similar to the lamellar bodies of mammalian epidermis (Fig. 3D and E). This layer constitutes most of the barrier against water loss. The latter layer recalls the corneous layer of the epidermis of the other vertebrates and its cells contain mainly a‐keratin. The cells of the inner generation do not further diVerentiate in this first renewal phase, while this process occurs in a cells of the above, outer generation, before it is shed from the inner generation. The lowermost a cells of the outer epidermal generation, destined to be sloughed, diVerentiate into a clear layer. In the meantime the first layer of the inner generation, the oberhautchen, also diVerentiates: clear and oberhautchen layers form the shedding complex. The split of the superficial keratinized epidermis takes place between the clear layer of the outer epidermal generation and the oberhautchen layer of the inner generation. It appears that the renewal phase lasts about 14 days in all species so far studied. This period should, however, be checked over many more species, as in some instances (some Agaminae lizards, and the tuatara Sphenodon punctatus) may have a diVerent length of shedding cycle. After the epidermis is shed, the germinal epithelium enters into a variable period of low activity or inactivity termed the resting phase. The alternation of renewal and resting phases is termed the shedding cycle (Maderson, 1985). This cycle allows expansion of the skin during the periodic somatic growth. In snakes the synchronous epidermal shedding determines the molt of the whole skin in a single piece but in most lizards this occurs as irregular flakes. In the archaic species Sphenodon punctatus, the a layer is made by multistratified, thinner and irregular cells than those of the b layer while lipids with a‐keratin filaments are accumulated (Alibardi, 2004a; Alibardi and Maderson, 2003a; Maderson, 1968). Lamellar granules are present in all a cells, not limited to mesos cells like in the epidermis of the other lepidosaurians. Large mucus granules often coexist with lamellar granules and may concentrate in true mucous cells containing large periodic acid–SchiV‐positive granules, a sign of primitive mucogenic activity. b Cells accumulate b‐keratin packets, and the mature (outer) b layer is made by a tile‐like association of individual cells, not fused into a syncytium like in modern lepidosaurians. Sphenodon punctatus, probably the most ancient extant lepidosaurian reptile, possesses the most epidermal layers compared with the other lepidosaurians with two exceptions: the oberhautchen and a specialized shedding complex (Alibardi, 2004a; Alibardi and Maderson, 2003a,b). This species probably sheds its epidermis (molt) along the intermediate region between the a layer of the outer epidermal generation and the inner b layer of the inner epidermal generation. This region does not produce a serrated or interdigitating, zip‐ fastened shedding complex like in the other lepidosaurians. It is also likely

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that the renewal stage is longer than the 14 days as in the other species, a period that allows the corneodesmosomes of the intermediate region to be digested. It is possible that the shedding complex of modern lizards and snakes evolved from a similar intermediate region, where a‐keratin is mixed with b‐keratin. 3. Crocodilians Extant archosaurians are represented by crocodilians and birds (Pough et al., 2001). Crocodilian scales are less varied in gross morphology than in lepidosaurians and they generally overlap very little (Alibardi, 2003a,b, 2005b; Alibardi and Thompson, 2000; Alexander, 1970; Parakkal and Alexander, 1972; Spearman, 1969). They have a large outer surface separated by narrow hinge regions. The outer scale surface will progressively expand into larger scutes during the growth of the animal, but this process is not known cytologically. The epidermis of the outer scale surface consists of a basal layer made of polygonal cells, three to six suprabasal layers made of flat cells, a transitional or precorneous layer, and a variably thick stratum corneum made of relatively thin corneocytes at maturity (Fig. 1H). Bundles of b‐keratin and a variable amount of melanosomes accumulate in cells of the transitional layer. Corneocytes in the dorsal part of the scale are 0.3–0.6 mm or thicker, show a spiny outline, and are composed mainly of b‐keratin. In hinge regions, the thickness of the epidermis decreases, and suprabasal keratinocytes accumulate lipid vesicles in their core while bundles of a‐keratin concentrate along the thickened plasma membrane. Mature corneocytes of hinge regions are much thinner than those on the outer scale and they resemble (sebo‐)keratinocytes of avian apteric epidermis. These corneocytes also resemble mesos or thin a cells of lepidosaurian epidermis. Their lipid‐rich corneous material does not show an a‐keratin pattern under the electron microscopic analysis.

D. Avian Epidermis: Adaptation to Flight and Homeothermy Although birds represent flying archosaurians, considered part of reptiles by many zoologists, the scaled integument has been largely lost in modern forms of the Cenozoic (Pough et al., 2001). The skin of birds presents many regional variations in apteric (nonfeathered) and pterylar (feathered) regions (Lucas and Stettenheim, 1972; Spearman and Hardy, 1985). Avian epidermis is therefore interrupted by feather follicles, and includes mainly interfollicular epidermis. Other areas include the scales tarsal–metatarsal areas (Sawyer et al., 1974, 1986).

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The histology, cytology, and ultrastructure of the epidermis in birds have been extensively analyzed (Alibardi, 2004b; Menon and Menon, 2000; Spearman and Hardy, 1985). The extremely soft and wrinkly apteric and interfollicular epidermis (Fig. 2D and E) seems to be associated with the need to maintain an elastic and flexible skin among feathers to favor their maneuverability. Because of the limited importance of apteric epidermis in the mechanical protection of the body, cornification is reduced in this type of epidermis. The delicate avian corneous layer has been termed ‘‘straw and mortar’’ in comparison with the ‘‘brick‐and‐mortar’’ mammalian corneous layer (Menon et al., 1986). The plumage takes up most of the mechanical protection (covering by flattening the feathers over the apteric epidermis), and acts as thermal insulation, leaving to the apteric epidermis a major role in the control of water loss (Menon and Menon, 2000; Menon et al., 1996). Apteric and interfollicular areas consist of a thin stratified epidermis made of a flat to cubic basal layer, two to four suprabasal layers (depending on body area), a transitional layer, and a thin corneous layer. The latter is made up of extremely thin corneocytes (0.05–0.2 mm). In the stratum basale, keratinocytes are cubic or flat and they contain sparse keratin bundles. The latter remain low in suprabasal cells, and increase only in precorneous (transitional) cells. Contrary to previous assertions, no keratohyalin is present in avian sebokeratinocytes, but keratin bundles tend to condense at the periphery of sebokeratinocytes. Complex lipids and waxes are synthesized in sebokeratinocytes, while the avian corneous cell envelope is probably simpler in comparison with that of mammalian corneocytes (Alibardi, 2004b, 2005c; Alibardi and Toni, 2004; Kalinin et al., 2002). The richness of lipids in avian epidermis (Menon et al., 1986) is related to water movement but may also be a means to protect the cutaneous surface from losing heat during flight. Lipids are accumulated in vesicular or multigranular bodies produced in spinosus keratinocytes (Landmann, 1980; Menon and Menon, 2000) (Fig. 3C). Sebokeratinocytes resemble mesos or thin a cells of reptilian epidermis. These lipid‐rich keratinocytes do not show an a‐keratin pattern under electron microscopic analysis because keratin‐ associated proteins are scarce or absent. The structure of scales and feathers, with their special type of keratinization, is presented in Section IV on epidermal derivatives.

E. Mammalian Epidermis: Adaptation to Different Environment and Homeothermy Although mammalian epidermis is used as a general example for vertebrate epidermis, it is in reality a specialized type of epidermis that evolved in the

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therapsid line of amniote evolution (Alibardi, 2003a; Maderson, 1972a; Spearman, 1964, 1966). The continuity of the epidermis is interrupted by hair follicles, so that most interfollicular epidermis covers mammalian skin. The morphologic variations and ultrastructure of mammalian epidermis have been extensively reviewed (Byrne et al., 2003; Holbrook, 1989, 1991; Matoltsy, 1986; Sokolov, 1982; Spearman, 1966). With respect to the dry and hard epidermis of extant reptiles, mammalian epidermis is soft, elastic, and moisturized and is coupled with the fine action of mammalian musculature. Mechanical protection has been taken over by a dense pelage that functions for thermal insulation (Findlay, 1970; Maderson, 1972a; Spearman, 1966). The softness of the mammalian epidermis has been related to the evolution of a granular layer to produce a soft stratum corneum (orthokeratotic). Parakeratosis (generally pathologic in extant mammals) has been considered a reversion to a more primitive form of cornification, possibly present in the first cotylosaurian reptiles from which therapsids and later true mammals derived (Spearman, 1964, 1966). The multilayered epidermis varies in thickness in diVerent mammals and body regions of the same individual (Sokolov, 1982). The epidermis can be linear but often presents folds or papillae that deepen into the dermis (Fig. 2F–H). It basically consists of a basal layer, where the mitotic activity is concentrated; suprabasal layers made by one to numerous layers of spinosus cells; one to three layers of granulosus cells; one or two layers of transitional cells; and a variably thick stratum corneum. The latter consists of a compact stratum (termed lucidum in some areas of the body) and a superficial stratum disjunctum, where corneodesmosomes are degraded, allowing desquamation. In the stratum granulosum, proteins (profilaggrin/filaggrin) aggregate into keratohyalin granules and function as an interkeratin matrix for cornification (Dale et al., 1994; Resing and Dale, 1991; Steinert, 1998). Other numerous proteins (involucrin, loricrin, sciellin, etc.) also mix with keratohyalin (Fig. 3G), and are eventually deposited along the plasmalemma to produce the resistant cell corneous envelope of mature corneocytes (Fuchs, 1990; Kalinin et al., 2002; Steinert and Marekov, 1995). Lamellar or Odland bodies are formed in cells of the upper spinosus and granular layers. They derive from the synthesis in the Golgi apparatus and smooth endoplasmic reticulum of multilamellar or Odland bodies (Elias et al., 1987; Menon and Norlen, 2002; Menon et al., 1986; Rawlings et al., 1994). These organelles release lipids into the extracellular space and contribute to the formation of the barrier in the stratum corneum (Elias et al., 1987; Matoltsy, 1986, 1987; Menon and Norlen, 2002). Vesicular or lamellar organelles have been found in the epidermis of birds and reptiles, and it is believed that these organelles have a similar role as a water loss barrier in all amniotes (Landmann, 1980).

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Ultrastructural studies of marsupial and monotreme epidermis have shown the unity of structure with the epidermis of placentals (Alibardi and Maderson, 2003c; Lyne et al., 1970). A broad survey on the skin of many mammalian species has shown some variation of the epidermis, in particular concerning the presence or absence of the stratum granulosum. The stratum granulosum is limited or discontinuous in the thin epidermis of monotremes, marsupials, chiroptera, and other placental mammals. It is likely that in the epidermis of other mammals where the stratum granulosum is not observed by light microscopy, keratohyalin granules may have submicroscopic dimensions (less than 0.5 mm).

III. Cytology of Keratinocytes and Their Organelles during Terminal Differentiation Aside from the general organelles present in most cells, keratinocytes possess specific organelles that allow the production of specific proteins used in the process of cornification and lipidization. The main cytoskeletal structures are represented by cytokeratin filaments that join desmosomes (or hemidesmosomes) present among keratinocytes of vertebrate epidermis (Fig. 2I). Bundles of denser tonofilaments accumulate in maturing keratinocytes. Other organelles are represented by intracellular mucus granules, and are produced from more apical keratinocytes and partially secreted on the epidermal surface of fish and amphibians (Fig. 3A). Mucus granules or dense protein granules form in upper keratinocytes of the amphibian epidermis, and remain in part associated with keratin bundles during the process of cornification (Fig. 3B). In the epidermis of amniotes, mucus granules disappear and lipid‐rich vesicular or lamellar bodies are formed in diVerentiating (upper spinosus) keratinocytes. Another peculiar diVerentiating product of maturing keratinocytes is the formation of a thickened cell membrane by the deposition of a denser material mainly on the inner leaflet of the plasmalemma, termed the cornified cell envelope (Fig. 3F). Finally, a typical organelle of mammalian keratinocytes, keratohyalin, is formed by the accumulation of nonkeratin and keratin proteins in the upper living layers (granular and transitional) beneath the corneous layer (Fig. 3G). Keratohyalin‐like granules of diVerent shape and dimension are present in lizard epidermis during the renewal phase. The above organelles are connected to the three main aspects of terminal diVerentiation of corneocytes in vertebrate epidermis (Alibardi, 2003a; Matoltsy, 1987). The latter are (1) increased concentration of keratin filaments with some mucoid matrix (stage of fish and amphibians); (2) increase

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in the amount of lipid‐rich component among relatively scarce keratin bundles (stage of birds); and (3) increased amount of matrix or interkeratin proteins among keratin bundles (stage of reptiles and mammals).

A. Soft Keratins or Cytokeratins in Vertebrates Cytokeratins constitute the main types of intermediate filaments ubiquitous in cells of vertebrates and invertebrates (Coulombe and Omary, 2002; Fuchs and Marchiuck, 1983; Fuchs and Weber, 1994; Fuchs et al., 1987; Luke and Holland, 1999; Steinert and Freedberg, 1991). More than 50 genes and various pseudogenes have been found to produce the diVerent types of keratin in mammals. Construction of the final keratin filaments derives from a progressive assembling of smaller subunits in mammalian keratinocytes. Aside from their general role in mechanical resistance of epithelia against mechanical sharing forces and tensile strength (‘‘hard principles’’), other roles (‘‘soft principles’’) have been ascribed to keratin filaments, including apoptosis and signal transduction (Coulombe and Omary, 2002). Keratin distribution in keratinocytes of the other vertebrates is still determined by use of panels of diVerent, more or less specific antibodies, against known mammalian keratins. Genomic and proteomic studies have allowed comparison of fish and amphibian keratin sequences with those of mammals (HoVmann and Franz, 1984; HoVmann et al., 1985; SchaVeld and Markl, 2005; SchaVeld et al., 2002a,b, 2003; Suzuki et al., 2001; Watanabe et al., 2001). These studies are completely missing for reptilian cytokeratins, and occasional studies are available on avian cytokeratins (Charlebois et al., 1990; Vanhoutteghem et al., 2004), although the chick genome is now available for screening. Mammalian keratins are therefore used as a general guide to the molecular biology of cytokeratins. Keratin filaments are made by the aggregation of intermediate (8‐ to 10‐nm‐thick) keratin proteins, made of a central a‐helical structure and two variable heads. These proteins aggregate according to the pair rule of one acidic keratin and one basic keratin (mainly type 1 keratin with type 10 keratin) in terminal keratinocytes (Cooper et al., 1985; Coulombe and Omary, 2002; Moll et al., 1982; O’Guin et al., 1987). The final X‐ray and ultrastructural a‐keratin pattern is of 8‐ to 10‐nm‐thick, electron‐pale filaments embedded in a denser matrix material. Cytokeratins constitute the 8‐ to 10‐nm‐thick intermediate filaments contributing to form the cytoskeleton of keratinocytes in all vertebrate epidermis. Two main types have been described according to the sequence of their genes, type I and type II, which have a diVerent genomic and protein structure in their main region (the central rod domain, made by about 310

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amino acids in mammals). Cytokeratins are formed by a highly conserved central rod region in which amino acid forms a‐helical secondary structure. The latter region further interweaves in a coiled‐coil conformation. Some regions within this conformation disrupt the coiled‐coil conformation. The remaining, shorter domains of cytokeratin consist of an N region (head) and a C region (tail), which have non‐a‐helical conformations. In tetrapods, type I includes cytokeratins with acidic isoelectric points (acidic cytokeratins, pI at 4.5–6.5, 40–59 kDa), and type II consists of those with basic isoelectric points (basic cytokeratins, pI at 6.5–8.5, 55–68 kDa) (Fig. 4). The main diVerences in acidic and basic cytokeratins are determined by the variable extension of the N region (head domain) and C region (tail domain), which varies extensively (from none to more than 150 amino acids). These regions are totally or partially regularly spaced on the sides along the main axis of cytokeratin filaments, made by tail–head joining of central rods. Type I (acidic) and II (basic) cytokeratins form heteropolymers according to the pair rule, forming the coiled‐coil domain of the basic unit (1 nm thick) that forms a supercoiled structure (the protofilament, 2–3 nm). Two protofilaments form a protofibril (4 nm), and four protofibrils form the 8‐ to 10‐ nm‐thick intermediate filament. Therefore, a keratin in modern terms should be considered a monomeric protein able to form long, filamentous structures within keratinocytes. This is a distinctive characteristic between keratins and keratin‐associated or matrix proteins (see below). In general, the selected pairs in the epidermis are characteristic and quantitatively prevalent relative to other possible combinations within the cytoplasm of keratinocytes involved in diVerent activities (e.g., K5/K14, K1/K10, and K6/K16; Cooper et al., 1985; Fuchs et al., 1987; O’Guin et al., 1987). So far, the highest number of protein spots and cytokeratin types have been isolated from humans and a few other mammalian species, than from other vertebrates. This status might change in future studies if cytokeratin pairs are found in the epidermis of reptiles, especially lepidosaurian reptiles, which apparently possess a more complex epidermis than that of mammals. In other amniotes, epidermis that lacks a specialized granular layer (birds, chelonians, and crocodilians) can be correlated to the presence of a minor number of keratin pairs (O’Guin et al., 1987). This will be determined only after the genomes of these organisms are known. 1. Fish Cytokeratins Cytokeratins in cyclostomes belong to the type reactive to antibody AE1 (K18 and K19, 40–45 kDa) and that are AE3 positive (K7 and K8, 54–56 kDa) (Alarcon et al., 1994; Zaccone et al., 1995).

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FIG. 4 Schematic presentation of protein patterns after bidimensional separation of epidermal proteins in various vertebrates. (A) Acidic keratins. a, a‐keratins (cytokeratins). (B) Basic keratins. b, b‐keratins; I, type I cytokeratins; II, type II cytokeratins. Numbers on the left indicate molecular weights. Fish (zebrafish, Danio rerio); lungfish (N. forsteri); amphibian (toad, Xenopus laevis); reptile (lizard, P. muralis); bird (chick, Gallus gallus); mammal (rabbit, Oryctolagus cuniculus).

Immunocytochemical and biochemical studies have shown the modality of keratinization in actinopterygian fish epidermis. The latter occurs by the synthesis of keratins with a narrower range of molecular mass (42–58 kDa) than in amphibians (47–69 kDa; Ellison et al., 1985) and amniotes (40–68 kDa). Although type I and II cytokeratins are present, they are all acidic

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(Alarcon et al., 1994; Alibardi, 2002a; Conrad et al., 1998; GroV et al., 1997; Markl et al., 1993; Markl and Franke, 1988; SchaVeld et al., 2003) (see Fig. 4). In fact, the pI of fish cytokeratins remains in the acidic range (4.2– 5.9). The lower specialization of keratins in fish may be correlated to the lower demand for mechanical resistance or specialization on their epithelia in the aquatic environment. More functions are required of epithelia in the terrestrial environment. This relative simpler specialization of fish keratinocytes is reflected in the simpler fish epidermis in comparison with that of terrestrial vertebrates. A broad presentation of fish cytokeratins, with their genes and derived proteins, has been done (SchaVeld and Markl, 2005; SchaVeld et al., 2002a,b). Actinopterygian fish epidermis presents AE1/AE3‐reactive cytokeratins in all or most layers (Fig. 1A–C). Homologs of cytokeratins 8/18 have been found, showing that fish epidermis is comparable to simple epithelia of amniotes. In sarcopterygian fish some conflicting results have been obtained. Whereas in the Australian lungfish (Neoceratodus forsteri) the epidermis is clearly reactive to the AE1 antibody, but less to the AE3 antibodies for mammalian basic keratins (Alibardi, 2001b, 2002a; Alibardi and Joss, 2003), this immunoreactivity is absent in the African lungfish (Protopterus aethiopicus) (SchaVeld and Markl, 2005). The immunoreactivity for AE2 (which recognizes K1/K10 keratins in mammalian epidermis) in N. forsteri is present in the outermost, more keratinized layers of the epidermis. Also in P. aethiopicus, a similar pattern of apical immunolocalization is observed when using antibody 79.14, which is directed toward cytokeratin‐8 (basic) of the amphibian Xenopus laevis. These results suggested some trend toward the amniote pattern of localization of these keratins (1, 8, and 10, although from diVerent vertebrates) in suprabasal layers of fish whose ancestors originated tetrapods. This is further supported by the isolation of keratin isoforms from the epidermis of N. forsteri with molecular mass ranging from 40 to 64 kDa and pI values of 5.7, 6.4, and 6.8 (M. Toni and L. Alibardi, unpublished observations) (Fig. 4). These values of molecular weight and pI are the highest so far detected in any fish keratin (ShaVeld and Markl, 2005). 2. Amphibian Cytokeratins Like fish keratins, those of the amphibian species studied so far indicate that cytokeratins are not uniquely present in the epidermis or other epithelia but are also present in mesenchymal or myoepithelial cells of the walls of blood vessels (Ferretti and Ghosh, 1997). Amphibian keratins belong to type I and II but, diVerently from fish, can also be divided into acidic and basic, although the basicity is lower than in amniote cytokeratins (Fuchs and Marchiuk, 1983; HoVmann and Franz, 1984; HoVmann et al., 1985) (Fig. 4). DiVerently from

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fish, type I already corresponds to acidic keratins (44–54 kDa, pI at 5.2–5.5) and type II to basic keratins (63–64 kDa, pI at 7.0–7.4). This characteristic is related to the formation of the corneous layer, and is a fundamental tetrapod characteristic toward the formation of the thicker corneous layer of the amniotes. In the newt Triturus vulgaris, bidimensional gel electrophoresis has identified keratins with molecular masses of 48–kDa (pI at 5.2, 6.2, and 7), 55–57 kDa (pI at 5.2, 6, 6.5, and 7.0), and 64 kDa (pI at 5.7, 6.2, 7, and 7.3) (M. Toni and L. Alibardi, unpublished observations). In the toad Bufo bufo, cytokeratins of 43–45 kDa (pI at 5.0 and 5.5), 50–52 kDa (pI at 4.8, 5.2, and 5.5), 55 kDa (pI at 5.8 and 6.0), 60 kDa (pI at 5.0, 5.3, and 5.9), and 63–64 kDa (pI at 5.6, 6.3, 7.0, 7.5, and 7.9) have been found (M. Toni and L. Alibardi, unpublished observations). The amino acid sequence of an acidic cytokeratin of 51 kDa in X. laevis has shown high similarity to human acidic keratin, and is almost devoid of a glycine‐rich region in the C‐terminal region. Conversely, from sequence analysis of three isoforms of a basic, type II cytokeratin of 64 kDa in X. laevis (HoVmann et al., 1985), the number of basic amino acids, especially in the variable external region of cytokeratins, increases. The lengthening of the C domain in particular determines both the increase in molecular weight and the shift of the pI toward a basic value, and contains glycine‐rich regions. The study also indicated the high conservation of amino acid domains in both the rod portion and tail portion (V2 region) of X. laevis and human cytokeratins. This indicates that this region can be involved in the formation of the stratum corneum or of the corneous cell envelope. The most recent studies on amphibian cytokeratins (Suzuki et al., 2001; Watanabe et al., 2001) indicate that eight genes for type I synthesize six acidic cytokeratins, and that five genes for type II synthesize four basic cytokeratins. These molecular data are confirmed by the distribution of acidic and basic cytokeratins determined by applying AE1, AE2, and AE3 antibodies on amphibian epidermis (Alibardi, 2001a, 2002b). While in aquatic and terrestrial urodele amphibians the uniform pattern of distribution of these cytokeratins recalls that of fish, and is immunonegative for AE2‐reactive cytokeratins, the pattern of cytokeratin distribution in the epidermis of terrestrial amphibians, especially anurans, often resembles that observed in mammals and reptiles. This suggests that in all tetrapods the sequence of production of keratin types is similar, with AE1 (acidic) keratins with low molecular weight synthesized in the basalmost layers. The AE3 (basic) keratins, with a higher molecular weight, are present in basal, suprabasal, and external layers, whereas the AE2‐positive keratins (61–68 kDa in molecular mass in mammals and lizards) are typical of keratinized layers. The production of larger keratins in more external epidermal layers has been found in X. laevis after metamorphosis (Ellison et al., 1985; Nishikawa et al., 1992).

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3. Reptilian Cytokeratins In general, a‐ or cytokeratins in reptilian epidermis appear to have molecular masses in the range of those in other vertebrates (40–70 kDa) (Carver and Sawyer, 1987; Fuchs and Marchiuk, 1983; Wyld and Brush, 1979, 1983). The distribution of AE1, AE2, and AE3 cytokeratins in the soft and hard epidermis of all reptilian orders has been studied (Alibardi, 2003a; Alibardi and Toni, 2005a,b; Alibardi et al., 2000, 2001, 2004a,b). Bidimensional gel electrophoretic studies have allowed the identification in reptiles of diVerent orders more specific types of both acidic and basic keratin fractions (L. Alibardi and M. Toni, unpublished observations; and see Fig. 4). The study suggests that diVerent isoforms of cytokeratin (K6, K16, and K17) are present in normal and regenerating epidermis (Alibardi and Toni, 2005c). Numerous cytokeratins have been found in the epidermis of the lizard Podarcis sicula, snakes (Elaphe guttata and Morelia carinata), molts of the tuatara (Sphenodon punctatus), and in the epidermis of a turtle (Chrysemys picta). No information is available on the cytokeratins of crocodilians. Because no data on the genome of any reptile are presently available, the number of reptilian cytokeratins remains unknown. Antibodies against cytokeratins (AE1, acidic cytokeratins; and AE3, basic cytokeratins) have been used to reveal specific a‐keratins. Bidimensional gel electrophoresis of lizard keratins (Fig. 4) shows AE1‐positive protein spots at 57–60 kDa with pI at 4.8–5.2, 5.8, and 6.3. The AE3 antibody showed numerous immunolabeled protein spots at 40–42 kDa, with pI at 6.2, 6.7, and 7.2. A main spot at 47–50 kDa and pI at 5.3 was seen. Main spots at 55–58 kDa with pI at 5.7, 6.0, 6.4, 6.8, 7.2, 7.7–8.0, and 9.6 were seen. The larger immunolabeled protein spots were seen at 60–66 kDa, with pI at 5.4–6.1, 6.8, 7.0, 7.5, and 8.3. Similar values have been found in Sphenodon, snake, and turtle cytokeratins isolated by bidimensional gel electrophoresis. These data indicate that at least 12–16 diVerent cytokeratins, basic and acidic, are present in reptilian epidermis. Three or four spots isolated at 10–24 kDa instead represent b‐keratins (Fig. 4). So far, no cytokeratin sequences are available to date for any reptilian cytokeratin. Therefore, although desirable, the direct comparison of sequences of reptilian cytokeratins with those from other vertebrates is not available yet. Information about the amino acid sequences of reptilian cytokeratins is needed before they are compared with those of mammalian epidermis and to trace the possible evolution and diversification of cytokeratins in amniotes. In general, the pattern of the tissue distribution of cytokeratins recalls that of mammalian epidermis, although the AE1 immunoreactivity is not restricted to the basal layer but also occurs in the suprabasal layers before they cornify (Figs. 1I and 2B and H). Immunocytochemical observations have indicated that acidic keratins of molecular masses 44–45 and 57–58 kDa tend

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to disappear in the a and b layers (Alibardi, 2000; Alibardi et al., 2001). Acidic keratins, typical of simple epithelia or precorneous cells, disappear where mature a‐ and b‐keratin layers are formed. Immunoblotting studies, after extraction and blotting of cytokeratins, however, indicate that also in corneous layers and molts of reptiles, acidic keratins are still detectable (M. Toni and L. Alibardi, unpublished observations). All together, these observations indicate that a‐keratins remain or are degraded in the corneous layers, especially in the a layer. Basic keratins (AE3) are instead retained in both corneous layers, including the b layer (Figs. 1F–H and 2A, C, and I).

4. Avian Cytokeratins Both acidic (type I) and basic (type II) cytokeratins have been described in apteric and interfollicular epidermis of avian epidermis (O’Guin et al., 1987; Sawyer et al., 2000) (Fig. 4). Acidic keratins (at least eight diVerent cytokeratins) have a molecular mass ranging between 41 and 59 kDa and their pI ranges between 4.0 and 6.5. Basic keratins (16 cytokeratins) have molecular masses ranging between 50 and 66 kDa and a pI that varies between 7 and 9. Cytokeratins K5/K14 and K1/10 have been detected in chick epidermis (SaathoV et al., 2004). A cytokeratin of deduced amino acidic sequence, reported in the literature for the chick embryo, shows high homology with type II keratins (Charlebois et al., 1990). The nucleotide and amino acid sequences of 10 cytokeratins containing 431 to 599 amino acids, and expressed in chick keratinocytes cultivated in vitro, have been published (Vanhoutteghem et al., 2004). They encode five cytokeratins each of type I (acidic) and type II (basic), and show 62–70% homology with mammalian cytokeratins. Three (IIA, IIB, and IIC) are basic cytokeratin members with homology to either mammalian K5 and K6 (however, it is uncertain whether they can be orthologs for K5 and K6). Two other acidic members instead appear to be the avian orthologs of mammalian K14 and K15. Although type I and other type II cytokeratins have some homology with those of mammals no further characterization and specific studies on avian cytokeratins, their genomic and molecular structure, localization, and so on, have been published. General antibodies against mammalian ‘‘prekeratins’’ (various cytokeratins including K1/K10) immunostain the epidermis (especially the stratum corneum) and the folded barb ridges of developing feather (Schmidt et al., 1979). The AE1 immunolabeling for acidic keratins is variably intense in avian epidermis, but is generally present in the basal and suprabasal layers, and is not limited to the basal layer as in human epidermis (Alibardi, 2004b, 2005b;

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Alibardi and Toni, 2004b). Acidic keratins (AE1 positive) are coupled to basic keratins (AE3 positive) according to the pair rule (one acidic keratin molecule is bound to one basic keratin molecule; see O’Guin et al., 1987). Reactivity to AE1 is lost in the uppermost epidermal layers. AE2 immunolabeling (for 56.5‐ and 66‐ to 67‐kDa mammalian keratins) in the corneous layer of the epidermis often shows a pattern similar to the pattern observed with antibodies against filaggrin. This is probably because the AE2 antibody also cross‐reacts with common epitopes between keratins and filaggrin (Dale and Sun, 1983). Immunolabeling with AE2 indicates the presence of a small amount of high molecular mass keratins, used for cornification (K1, 66–67 kDa and K10, 56.5 kDa; O’Guin et al., 1987) in apteric keratinocytes. Basic keratins (AE3 positive) of variable molecular weight (Fig. 4) are also present in the transitional layer, and partially in the corneous layer (Alibardi, 2004b; Alibardi and Toni, 2004b; O’Guin et al., 1987). 5. Mammalian Cytokeratins In mammalian epidermis, diVerent types of acidic (I) and basic (II) keratins associate in pairs to form tonofilaments of diVerent properties and mechanical strength. Only some general comments on their tissue distribution in the epidermis of diVerent mammalian species are reported here. These keratins are present in the basal layer (K5, 58–60 kDa; to K14, 50–53 kDa), in the suprabasal layer, and in the granular layer (58 and 67 kDa). Finally, other types of keratins are localized in the stratum corneum (57.5–58.5 to 62–64 kDa) (Bowden et al., 1987; Cooper et al., 1985; Moll et al., 1982) (see Fig. 4). The general pattern of cytokeratin distribution in human epidermis shows that the AE1 antibody specifically labels the basal layer. AE2 antibody labels keratins involved in cornification of the upper spinosus layer and final corneocyte layer of the epidermis (K1, 67 kDa; and K10, 56.5 kDa). AE3 antibody (for basic keratins) tends to stain most living layers. The human AE1–AE2–AE3 pattern, however, is not precisely observed in all mammalian epidermis because the AE1 antibody not only stains the basal layer but also various spinosus layers (Alibardi and Maderson, 2003c). Testing the above‐described antibodies over several species of monotreme, marsupial, and placental mammals has often shown a similar pattern of staining between AE1 and AE3, therefore diVerent from that of human epidermis (Fig. 2F and G). In other species the human pattern is present. These variations are probably related to the type of keratin pair present in keratinocytes of diVerent mammalian species (O’Guin et al., 1987). The localization of low molecular weight AE1‐positive keratins in the basalmost layers is also typical of early proliferating or stratifying epithelia. In addition, the AE2 pattern, associated with the presence of K1 (65–67 kDa,

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basic) and K10 (56.5 kDa, acidic) keratins, can sometimes overlap with that of filaggrin, as the two proteins have a common antigenic determinant (Dale and Sun, 1983). These initial studies have been replaced by the use of monospecific antibodies that have clarified the localization of specific keratin pairs in diVerent epidermal layers (Bowden et al., 1987; Cooper et al., 1985; Moll et al., 1982). Studies with knockout mice have shown that artificial cytokeratin pairs (e.g., K5/K14 replaced by K5/K16, or K8/K18 replaced with K8/K19 or K8/ 20) have less stability than the original, natural pair (Coulombe and Omary, 2002; Paladini and Coulombe, 1999), leading to epidermal impairment and diseases. It also known that cytokeratins can vary their electrophoretic pattern and pI by process of phosphorylation, glycosylation, transglycosylation, proteolysis, or aggregation with other proteins (Boweden et al., 1987; Coulombe and Omary, 2002; Steinert and Freedberg, 1991).

B. Keratin‐Associated Proteins Keratins constitute the fibrous, elongated part of the maturing cytoplasm of keratinocytes, and tend to form parallel filaments in which single filaments, 8–12 nm thick, are distinct (Fig. 3B). Various nonkeratin proteins, also present in the cytoplasm of keratinocytes of diVerent vertebrates, simply fill the space between keratins and have no specific or known interaction: possibly mucins and complex lipids. Other nonkeratin proteins have instead a specific interaction with keratin filaments: the latter are termed keratin‐ associated proteins. 1. Mucins and Lipids: Water and Thermal Barrier The production of mucus and glycoproteins occurs in the Golgi apparatus of keratinocytes, although most mucus is produced in specialized goblet cells in fish epidermis. In general, mucus contains neutral and acidic mucopolysaccharides that prevail over proteins and lipoproteins (Lebedeva, 1999). Small and large mucus‐like granules of amphibians are produced in the upper stratum spinosum (Fig. 3B). The large granules remain mainly intracellular, suggesting they have some matrix function. They contain neutral and sulfated mucopolysaccharides but lack sialic acid, and glycoproteins are made for more than 70% of glycans (Toledo and Jaret, 1992). Mucus and glycoproteins are produced in fish and amphibian keratinocytes among keratin filaments: it is unknown whether this glycoprotein–keratin complex has specific cytoskeletal eYciency greater than that of the keratin network. Present information, however, excludes a specific, interkeratin or matrix role

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for mucus/glycoproteins in organizing the polymerization of cytokeratins. It is possible that lateral chains of glycoconjugates have been progressively replaced with glycans bonded to lipid molecules to form nonpolar glycolipid molecules typical of amniote epidermis (Elias et al., 1987). Therefore mucus indirectly regulates the movement of water across the epidermis by specific bonding of sodium and potassium ions, as previously indicated. In amphibians more adapted to terrestrial life, complex lipids (waxes, glycosphingolipids, etc.) have been added to the keratin in mucus secretions to form a horny material with more mechanical and water loss resistance (Lillywhite and Maderson, 1982; McClanahan et al., 1978). In an American tree frog neutral and polar lipids (phospholipids) are produced, most of which are wax esters, followed by triglyceride esters with oleic, linoleic, and palmitic acids. Hydrocarbons and cholesterol esters are reduced and polar lipids are nearly absent (glycolipids and phospholipids). Glycoproteins, glycolipids, and other lipids are generally extruded into the extracellular spaces among corneocytes, between the horny and replacement layers, and cover the external surface of the stratum corneum. In amniote epidermis lipids become more complex and abundant than mucus, and accumulate intra‐ and extracellularly for barrier purposes, including coating of the cell surface with waxes (Elias et al., 1987). A new organelle, the vesicular or lamellar body, is present in amniote keratinocytes (Elias et al., 1998; Landmann, 1980; Menon and Norlen, 2002; Menon et al., 1986, 1992, 1996; Rawlings et al., 1994) (Fig. 3C–E). These organelles are isolated in the cytoplasm or connected to the smooth endoplasmic cisternae to form a network in functional continuity with the trans‐Golgi vesicles. They contain high levels of glucosyl ceramides, phospholipids, and cholesterols, but also store lipases, sphingomyelinases, b‐glucosyl cerebrosidase, phosphodiesterase, and proteases in corneodesmosomes (Menon et al., 1992). Specific markers for these organelles, at least in mammals, appear to be specific basic proteins of 25 kDa (O’Guin et al., 1989), elafin (Nakane et al., 2002), a special ceramide and glucosyl ceramide (Vielhaber et al., 2001), and caveolin
1 and
3 (Sando et al., 2003). Caveolin‐1 protein, with the same molecular mass of that in mammalian epidermis (22–23 kDa), has been detected in the epidermis of the soft‐shelled turtle, Trionyx spiniferus (Alibardi and Toni, 2006). In the latter species the richness of smooth endoplasmic reticulum and associated vesicular bodies (the analogous turtle organelle for lamellar bodies) is related to the formation of a waterproof, lipid barrier, for water adaptation in this fully aquatic turtle. Lipids in the mesos and a layers of snakes, among reptiles, show a composition that already recalls that present in mammals (Roberts and Lillywhite, 1980, 1983): cholesterol esters, wax esters, free fatty acids (palmitic, oleic, and methyl derivates), phospholipids (phosphatidylethanolamine, phosphatidylserine, phosphatidylinositol, and phosphatidylcholine), and sphingomyelin.

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In avians the lipid composition also shows an abundance of sphingolipids and neutral lipids, and scarce phospholipids, glycosphingolipids, and triglycerides (Menon et al., 1986). A high amount of enzymatic activity involved in the synthesis of lipids [3‐hydroxybutyrate dehydrogenase (BDH), a‐glycerophosphate dehydrogenase (a‐GPDH), etc.] has been detected by histochemical methods. In mammals the stratum corneum represents the main site of the barrier against water loss, microbial penetration, and chemical and mechanical protection. The barrier to water loss is particularly strong in the stratum compactum, where bilayered lipid forms extracellular lamellae after lipids have been released from lamellar bodies. Main lipids are represented by glycolipids, in particular some types of acyl‐glucosyl ceramides. Most of the latter form acylceramide‐1, a metabolic derivatives that is eVective in forming the barrier. These lipids form a ‘‘gel phase’’ in the extracellular space that eVectively limits water movement. Minor components are represented by long‐chain fatty acids (mainly saturated) and cholesterols: these two components may be free among corneocytes or bonded to the extracellular side of the corneous cell envelope. Lipids are bonded to proteins of the outer side of the plasma membranes by transesterification catalyzed by transglutaminase‐1 (epidermis) to glutamine/glutamic residues, especially of involucrin (Kalinin et al., 2002). 2. Matrix Proteins and Formation of Intracellular Corneous Material Although keratin filaments condense within corneocytes in some areas of fish epidermis (Mittal and Banerjee, 1978; Mittal and Whitear, 1979) and in amphibian corneocytes (Alibardi, 2001b, 2002a; Forbes et al., 1975), an a‐keratin pattern is generally absent or limited to small areas of corneocytes. In electron‐dense areas at the periphery of horny cells, isolated keratin filaments are no longer visible, suggesting that matrix proteins are present and form a true corneous material. The only keratin‐associated interkeratin proteins known so far are histidine‐rich proteins (HRPs) produced in the mammalian granular layer: profilaggrin and filaggrin (Dale et al., 1994; Fukuyama and Epstein, 1986; Holbrook, 1989; Resing and Dale, 1991). Filaggrin is produced in the stratum granulosum or transitional layer from a much higher molecular weight precursor called profilaggrin (Dale et al., 1994; Harding and Scott, 1983). Filaggrins aggregate keratin filaments into dense bundles that trap other degrading products and form the dense intracellular corneous material of corneocytes. This occurs by regularly distributed polar charges of filaggrin that interact with those of keratin (Mack et al., 1993). This association retards or inhibits the degradation of keratins by proteolytic enzymes in the

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transitional layer and lowermost part of the corneous layer. The enzyme arginine deiminase in the stratum corneum gradually transforms arginine residues into citrulline, and determines loss of the link between filaggrin and keratin. Filaggrin is then degraded in the stratum corneum to form a mix of peptides and amino acids termed ‘‘natural moisturizing factor’’ that seems to be involved in maintaining the hydration and softness of the stratum corneum. Immunological and sequence analyses of filaggrins have shown that the molecular weight of these proteins is variable among species and that these proteins present poor cross‐reactivity among even close species. To evaluate the possible presence of HRPs in the epidermis of nonmammalian vertebrates, the epidermis of fish, amphibians, chelonians, lepidosaurians, and birds was treated with tritiated histidine (Alibardi, 2003a) (Fig. 5A–F). Both autoradiographic and electrophoretic analyses of protein fractions were conducted to reveal the presence of putative HRPs. Tritiated histidine is generally incorporated into proliferating regions of the epidermis, and is a general indication of increased protein synthesis: this occurs exclusively in the epidermis of fish, chelonians, and birds. On the other hand, in the renewal epidermis of lizards and snakes, and in the renewal epidermis of amphibians, labeling is enhanced in epidermal layers connected with shedding (Fig. 5A, D, and E). The labeling suggests that, aside from the intense metabolism of these cells, more specific proteins involved in cornification of these cells may be present. This is indicated by the presence of labeled nonkeratin protein bands in the epidermis (Alibardi et al., 2003a,b). Bidimensional gel electrophoretic studies in newt, toad, and lizard epidermis indicate that small amounts of basic, nonkeratin proteins are present in their epidermis (L. Alibardi and M. Toni, unpublished observations). Detailed ultrastructural analysis has shown in both amphibian and reptilian epidermis a preferential labeling of dense areas among keratin filaments or over keratohyalin‐like granules among keratin filaments (Fig. 5G and H). However, even in cases of labeling of keratin bundles at the ultrastructural level, it is not certain whether true keratins or mixed keratin‐associated proteins are labeled. The lack of filaggrin‐like immunolabeling and of keratohyalin‐like granules in fish and amphibian epidermis further indicates the absence of HRP‐like molecules. In the soft epidermis of lizards, turtles, and birds, a filaggrin‐like immunoreactivity similar to that of mammalian epidermis has been detected (Fig. 6A–E). This immunolabeling overlaps that produced by the AE2 antibody, which cross‐reacts with an epitope in common with filaggrin (Dale and Sun, 1983). This suggests that the filaggrin‐like immunoreactivity of nonmammalian epidermis is due to some cross‐reactive epitopes present in keratins of the stratum corneum in these species. In clear cells of lizard epidermis, small keratohyalin‐like granules show weak but localized immunolabeling for filaggrin and are surrounded by a

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FIG. 5 Autoradiographic detection of tritiated histidine in vertebrate epidermis. (A) Labeling is present mainly in the precorneous and corneous layers (arrow) of frog (R. esculenta). Scale bar: 10 mm. (B) DiVuse labeling is visible through the soft limb epidermis of turtle (C. picta). Scale bar: 10 mm. (C) Labeling is concentrated mainly in the transitional layer beneath the growing corneous layer of turtle carapace (C. picta). Scale bar: 20 mm. (D) Labeling is concentrated mainly in the forming shedding line of snake scale (Natrix natrix). Scale bar: 10 mm. (E) Labeling (arrows) is concentrated mainly along the shedding line occupied by keratohyalin‐like granules in lizard (P. muralis). Scale bar: 10 mm. (F) DiVuse labeling is visible in apteric skin of zebra finch (T. castanotis). Scale bar: 10 mm. (G) Ultrastructural detail of labeled dense material (arrowhead) among keratins in a precorneous keratinocyte of toad (Bufo bufo). Scale bar: 100 nm. (H) Labeled keratohyalin‐like granules in the clear layer of lizard (P. muralis). Scale bar: 200 nm. c, stratum corneum; d, dermis; e, epidermis; k, keratin bundle; khl, keratohyalin‐like granule.

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FIG. 6 Immunocytochemical detection of filaggrin‐like reactivity in vertebrate epidermis. (A) Turtle limb (C. picta). Scale bar: 10 mm. (B) Lizard epidermis (P. muralis). Scale bar: 20 mm. (C) Epidermal peg of regenerating lizard scale (P. muralis). Scale bar: 20 mm. (D) Human epidermis (H. sapiens). Scale bar: 10 mm. (E) Rat epidermis (R. norvegicus). Scale bar: 10 mm. (F) Ultrastructure of double‐gold immunolabeling of lizard epidermis (P. muralis). Weakly immunolabeled keratohyalin‐like granules are evident (arrowhead to small gold particles), while the surrounding keratin filaments are labeled (arrows on larger gold particles) for AE3‐positive keratins. Scale bar: 100 nm. a, a layer; b, b layer; c, corneous layer; g, granular layer; khl, keratohyalin‐like granules; Dashes underline the basal layer.

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cytokeratin mass of keratin bundles (Fig. 6F). The formation of a shedding layer in lizard, snakes, and amphibians seems to be associated with the synthesis of histidine‐rich molecules in the range of 22, 29–30, and 38 kDa (Alibardi and Toni, 2004a, 2005a,b; Alibardi et al., 2003a,b; and our unpublished observations). Filaggrin‐like labeling is also present over granular or vesicular bodies in turtle and avian keratinocytes of the transitional layer contacting the stratum corneum (Alibardi and Toni, 2004b). Immunoblotting cross‐reactive bands at 33 and 42 kDa are present in lizard epidermis, and at 50–52 and 62–64 kDa in turtle epidermis. Bands at 32, 38, and 45–48 kDa appear in avian soft epidermis. 3. Cornified Cell Envelope Proteins During terminal diVerentiation of mouse and human corneocytes numerous proteins participate in the formation of the 15‐ to 25‐nm‐thick ‘‘cornified cell envelope’’ (CCE) (Ishida‐Yamamoto and Iizuka, 1998; Steinert, 1998; Steinert and Marekov, 1999; Kalinin et al., 2002). Many proteins (about 20 diVerent main types recognized so far, such as sciellin, involucrin, small proline‐rich proteins, keratolinin, and loricrin) compose the envelope. Immunoreactivity to these proteins, however, already appears in upper spinosus, granular, and corneous layers (Figs. 7 and 8). Most of these proteins have a structural role in mechanicochemical protection (involucrin and loricrin) but others, in addition to cross‐linking to stabilize the cornified membrane, form a ‘‘physiological barrier.’’ In particular, some proteins such as cystatin‐ A and elafin, although cross‐linked, have an enzymatic action, respectively, as pepsinogen inhibitor of bacterial enzymes (protecting against microbial invasion), or as inhibitor of elastase and proteinase derived from polymorphonuclear leukocytes. These proteins are represented in human and mouse chromosomes (e.g., 1q21 in humans) as linear gene sequence clusters in a special region termed the ‘‘epidermal diVerentiation complex.’’ Finally, many of the proteins clustered in the epidermal cornification complex appear to share common amino acid sequences at their N and C terminuses, indicating that these proteins evolved from a common precursor (Backendorf and Hohl, 1992; Kalinin et al., 2002). Instead, the central motif of the molecule of this family of proteins has specifically diversified in order to produce various proteins with specific functions in the formation of the CCE. Involucrin is an abundant, acidophilic, and hydrosoluble protein that participates in the formation of the physical barrier of the CCE. It is initially free in the cytoplasm of keratinocytes of the upper stratum spinosum and linked to the amorphous material present among keratohyalin granules of the stratum granulosum (Fig. 8B). It is a 585‐amino acid protein with a

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FIG. 7 Immunocytochemical detection of loricrin‐like reactivity in vertebrate epidermis. (A) Regenerating lizard epidermis (P. muralis). Scale bar: 10 mm. (B) Mature lizard epidermis (P. muralis). Scale bar: 10 mm. (C) Snake epidermis (N. natrix). Scale bar: 10 mm. (D) Turtle tail epidermis (C. picta). Scale bar: 10 mm. (E) Zebra finch apteric epidermis with reactive transitional (arrow) and corneous layers. Scale bar: 10 mm. (F) Rat epidermis (R. norvegicus). Scale bar: 10 mm. (G) DiVuse immunogold‐labeled corneous layer of turtle tail epidermis (C. picta). Scale bar: 100 nm. (H) Immunolabeled vesicular body in transitional layer of ostrich epidermis. Scale bar: 100 nm. c, corneous layer; g, granular layer; t, transitional layer. Dashes underline the basal layer.

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FIG. 8 Immunocytochemical detection of cornified cell envelope proteins in vertebrate epidermis. (A) Sciellin labeling of corneous layer (arrow) of cat epidermis (Felis catus). Scale bar: 10 mm. (B) Involucrin labeling in the upper spinosus, granular, and corneous layers of human epidermis (H. sapiens). Scale bar: 10 mm. (C) Transglutaminase labeling of rat epidermis and corneous layer inside a hair canal (R. norvegicus). Scale bar: 10 mm. (D) Transglutaminase labeling of the corneous layer (arrow) of cat epidermis (F. catus). (E) Isopeptide labeling in the corneous layer (arrow) of the epidermis and of the hair canal. Scale bar: 10 mm. (F) Ultrastructural detail of transglutaminase‐labeled granule (arrow) within an a‐corneocyte of lizard epidermis. Scale bar: 100 nm. c, corneous layer; g, granular layer; h, hair. Dashes underline the basal layer.

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molecular mass estimated, by diVerent methods, as between 68 and 92 kDa. The protein, believed initially to be present only in primates, is now recognized more generally in mammalian epidermis (Kubilius et al., 1990). Small proline‐rich proteins also participate in building the physical barrier of the CCE. In fact, they are particularly abundant in epidermis with a thick corneous layer (e.g., plantar epidermis). They contain more than 30% proline, which is localized mainly in repeated domains of the central part of these small and basic proteins (10–30 kDa, more than 10 types). They have amino acid consensus sequences recognized by transglutaminases (TGases) that catalyze their bonding to other proteins of the CCE. These proteins are initially soluble in upper spinosus keratinocytes but become associated with the CCE in the transitional layer. Keratolinin or cystatin‐A is a 10‐ to 12‐kDa monomer protein that organizes into a 36‐kDa active polymer, which can further polymerize in forming the CCE. The protein is present in the cytoplasm of spinosus keratinocytes and becomes rich in citrulline in the CCE, where it is bonded mainly to loricrin. Elafin is a 6‐ to 7‐kDa serine‐protease inhibitor that is derived from a 12‐kDa precursor of 117 amino acids. It is rich in cysteine and proline, has transglutaminase amino acid motifs, and becomes cross‐linked to other proteins of the CCE by isopeptide bonds and also by disulfide bonds. Sciellin is a 75‐ to 82‐kDa protein consisting of 668 amino acids, pI 8.0–9.4, and rich in glutamine and lysine. This protein is relatively diVerent from the other proteins of the CCE (Champliaud et al., 2000; Kvedar et al., 1992). Keratins (especially K1/K10) and filaggrin interact with the other proteins of the CCE. In particular, the glycine‐rich segments of keratins (V1 and V2) may interact with the glycine‐rich loop of loricrin. The latter is the major component of the CCE (more than 70% of all the proteins; Mehrel et al., 1990) and presents a high amount of glycine (more than 55% of the total amino acids), followed by cysteine and serine. The molecular weight varies among mammals, from 26 kDa in humans to 36 kDa in mouse, because of variation in internal repeats (Hohl et al., 1993). Loricrin is present in round (L)‐granules of keratohyalin or over the amorphous matrix of keratohyalin granules (Figs. 3G and 7F). The protein is later dispersed along the plasma membrane of corneocytes of the stratum corneum (Hardman et al., 1998; Ishida‐Yamamoto et al., 2000; Steven et al., 1990). Larger, specialized keratins expressed in the stratum corneum (K1/K10) interact through their glycine‐rich regions (variable regions, near the N or C end) with the glycine‐rich sequences of loricrin. Two other large proteins, periplakin (200 kDa) and envoplakin (230 kDa), do not localize within the epidermal diVerentiation complex. The former protein forms chemical bonds with lipids on the outer plasma membrane, and the latter protein binds keratins on the cytoplasmic side of the plasma membrane.

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The formation of a complete CCE (which is activated by cytoplasmic influx of calcium) takes place by the cross‐linking action of TGase (especially type 1). The initial steps involve the link with periplakin–envoplakins, followed by involucrin, small proline‐rich proteins, elafin, envoplakin, filaggrin and glycine‐rich keratins, cystatin‐A, and loricrin (Kalinin et al., 2002; Steinert and Marekov, 1995). It is not known whether an epidermal diVerentiation complex is present in chromosomes of the other vertebrates in which a CCE is present. To address this problem, it is essential first to detect the presence of proteins that cross‐ react with antibodies for mammalian CCE proteins. Granules containing nonkeratin sulfur components are present in the precorneous or replacement layer of amphibians (Alibardi, 2003a). Most sulfhydryl groups and sulfur are present along the plasma membrane of amphibian corneocytes. No loricrin has, however, been detected by immunocytochemistry in fish and amphibian corneocytes so far. However, using immunoblots, bands at 25 and 52 kDa have been found (Alibardi and Toni, 2004a). Positive immunodetection (both by immunocytochemistry and immunoblotting) for loricrin, involucrin, and sciellin has been obtained for reptilian and avian epidermis (Alibardi, 2003a; Alibardi and Toni, 2004a,b, 2005a,b) (Figs. 7A–F and 8A, B, and E). The latter results suggest that epitopes are masked in tissues used for immunocytochemistry. Loricrin immunoreactivity appears over the a layer of normal and regenerating reptilian epidermis, including keratohyalin‐like granules of lizards. In the transitional layer of reptilian and avian epidermis labeling is diVuse but is present mainly over electron‐pale and heterogeneous electron‐dense vesicles, which resemble vesicular bodies or dense granules (Fig. 7G and H). The labeling becomes diVuse in the mature corneous layer but, unlike corneocytes of placental mammals, does not concentrate along the cell corneous membrane. Immunoblotting results indicate that reptilian loricrin‐like proteins show molecular mass mainly in the range of 52–57 kDa for turtle and lizard epidermis. In avian epidermis a range mainly at 48–54 kDa has been found. By immunogold cytochemistry, loricrin‐like immunoreactivity is not detected in the CCE but only in vesicular‐like bodies discharged on the corneocyte surface. Sciellin has not been detected in anamniote epidermis by immunocytochemistry. Conversely, by immunoblotting main bands at about 42 kDa have appeared in fish; at 45, 52, and 80 kDa in amphibians; at 45, 53, 60, and 62 kDa in reptiles; and at 45 and 52 kDa in birds. Using immunocytochemistry, sciellin‐like immunoreactivity is present, although with broad variation in intensity, in the soft epidermis of turtle and lizards, but not in that of birds (Fig. 8A). Isopeptide bonds, detected by immunoblotting, are also

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present in the epidermis of amphibians, reptiles, and birds, indicating the presence of TGase responsible for the formation of the bond (Fig. 8E). Future, more detailed proteomic and genomic studies will better illustrate the characteristics of these proteins in nonmammalian vertebrates. 4. Enzymes of Cornification and Apoptosis The process of terminal diVerentiation of keratinocytes in their progressive advancement from the basal layer to the external or corneous layer eventually produces dead cells: in terrestrial vertebrates there is a basal–apical polarity, and cell death is reached in the precorneous layers. Conversely, in fish keratinocytes death occurs both in intermediate and external layers (tiers). Cell necrosis with organelle disruption and vacuolization involves lysosomal degradative enzymes (Whitear, 1986; Zaccone et al., 2001). From amphibians onward a true programmed cell death, more than a specific apoptotic process, is the rule (Alibardi, 2003a; Haake and Polakowska, 1993; McCall and Cohen, 1991; Polakowska and Goldsmith, 1991). The outcome is the formation of one (amphibians) or more layers of dead corneocytes stratified on the epidermal surface. Enzymes in the epidermis, especially those involved in cornification, are represented mainly by TGases and sulfhydryl oxidases (Hashimoto et al., 2000; Kalinin et al., 2002; Polakowska and Goldsmith, 1991; Rice et al., 1994; Yamada et al., 1987) (Fig. 8C, D, and F). These enzymes determine the formation of both isopeptide binding and disulfide binding that stabilize the CCE (Fig. 8E), anchor the latter to inner keratin‐degraded proteins, and form a cross‐linked network of dense keratin filaments. Both in the epidermis and inner root sheath of hairs a special type of resistant chemical bond, an E‐(N‐g‐glutamyl)‐lysine or isopeptide bond, is formed among proteins of the cornified cell envelope. This reaction is catalyzed by epidermal transglutaminase on substrates such as involucrin, sciellin, loricrin, and so on. Two forms of transglutaminase have been detected, soluble and membrane bound, the latter especially involved in the formation of the cornified cell envelope of the epidermis. Comparative biochemical and immunoblotting analyses have shown that transglutaminases are present in the epidermis of fish, amphibians, reptiles, and birds (Alibardi and Toni, 2004a,b, 2005a,b; Polakowska and Goldsmith, 1991; Rice et al., 1994). The molecular mass of transglutaminases varies from 50 to 60 kDa in most tested vertebrates. Immunolocalization of the enzyme shows a diVuse labeling in a‐corneocytes, or is present in round organelles of unclear nature, and with dimension of 0.1–0.2 mm (Fig. 8F). Other enzymes, such as peptidylarginine deiminase and matriptase, intervene in keratin degradation (Coulombe and Omary, 2002) or take part in the

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alteration of keratin‐associated proteins such as filaggrin and trichohyalin (Harding and Scott, 1983; Resing and Dale, 1991; Rogers, 2004; Rogers et al., 1999). Other important enzymes are represented by proteases involved in terminal diVerentiation such as caspase‐14 (Alibardi et al., 2005; Eckart et al., 2000; Rendl et al., 2001). In both avian and mammalian epidermis, caspase‐ 14 is present mainly in precorneous and corneous layers, or in the inner root sheath of hairs (Fig. 9A–D). This enzyme is absent in the epidermis of fish and amphibians, but some immuno‐cross‐reactivity seems to be present in lizard, snake, and turtle epidermis (L. Alibardi and M. Toni, unpublished observations). Caspase‐14 is a cysteinyl‐protease that cleaves substrates in a specific tetrapeptide sequence. In mammals, the protein consists of two proenzyme forms, one at 11–12 kDa and another at 17–18 kDa, which form a dimeric and active form at 28–30 kDa. Some immunolabeled bands appear at 17–18 kDa in avian epidermis, especially in the corneous layer of scales, and in the sheath of developing feathers (L. Alibardi and M. Toni, unpublished observations). In some cases, weak but cross‐reactive protein bands at 16–18, 22, and 27–30 kDa are present in protein extracts of turtle, lizard, and snake epidermis. This enzyme, unlike other caspases, is not activated by apoptotic signals and appears to be involved in the process of terminal diVerentiation of keratinocytes. The enzyme is distributed in the cytoplasm of suprabasal and granular cells, in the latter associated with keratohyalin (Alibardi et al., 2005a). A nuclear localization has also been found, in particular in heterochromatin clumps of transitional corneocytes of reptilian, avian, and mammalian epidermis (Fig. 9E). The enzyme is eventually diVuse in corneocytes, including their corneous cell envelope, with a loricrin‐ or involucrin‐like distribution (Fig. 9F). The specific substrates of caspases are not yet known. Numerous enzymes are involved in epidermal desquamation of the outermost cells of the stratum corneum (Menon and Norlen, 2002; Menon et al., 1992; Milestone, 2004; Rawlings et al., 1994). The process is caused by lytic enzymes, many of which are derived from lamellar bodies after they have discharged their lipid contents extracellularly. These enzymes comprise glycosidases and proteases (stratum corneum tryptic enzymes/cathepsins/thiol protease‐cathepsin‐2), and degrade the proteins of desmosomes (desmoglein, desmocollin, desmoplakin, and corneodesmosin). During the process of dissolution, activation of the general lysosomal marker acid phosphatase has been described in the epidermis of reptiles (Alibardi, 1998; Goslar, 1964), birds (Menon et al., 1986; Spearman, 1966), and mammals (Freinkel et al., 1983; Spearman, 1966). The enzyme mainly indicates activation of diVerent lysosomal proteases for degradation of cell organelles during cornification, including keratin filaments.

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FIG. 9 Immunocytochemical detection of caspase‐14 in vertebrate skin. (A) Immunoreactive corneous layer of chick scale (G. gallus). Scale bar: 10 mm. (B) Slightly immunoreactive corneous layer of echidna epidermis (T. aculeatus). Scale bar: 10 mm. (C) Immunoreactive inner root sheath (arrow) of cross‐sectioned hamster hair (Mesocricetus auratus). Scale bar: 10 mm. (D) Longitudinal section of cat hair (F. catus) showing reactivity in the inner root sheath (arrow). Scale bar: 20 mm. (E) Details of immunolabeled chromatin in a cell of the stratum corneum of lizard epidermis (P. muralis). Scale bar: 100 nm. (F) Immunolabeled corneocyte of human epidermis (H. sapiens). Arrow indicates a desmosome remnant. Scale bar: 200 nm. bu, hair bulb; c, corneous layer; e, epidermis; h, hair; nu, nucleus; m, medulla; o, outer inner sheath.

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C. Hard Keratins or Keratin‐Associated Proteins Cytokeratins are present in most epidermal layers, where they play a role in general mechanical resistance, impart form to cells, or determine their changes in shape. Small ‘‘hard keratins’’ are instead proteins present in hard structures (scales, scutes, claws, beak, feathers, etc.), and are specifically produced in the precorneous layer of the epidermis or in specialized skin derivatives. In addition to soft cornified epidermis, hard corneous layers of scales, claws, beak, feathers, and so on, are formed in reptilian and avian skin. Also in mammalian scales, hairs, horns, hoof, nails, claws, and so on, hard corneous layers are produced. Keratinocytes of hard keratinized appendages produce proteins termed ‘‘b‐ or ‐keratins’’ in reptiles and birds (Fraser and MacRae, 1978; Fraser et al., 1972; Gregg and Rogers, 1986; Sawyer et al., 2000, 2005; Wyld and Brush, 1979, 1983). In mammalian appendages hard proteins are termed ‘‘keratin‐associated proteins’’ (high glycine–tyrosine‐rich proteins, high‐sulfur proteins, ultrahigh‐sulfur proteins in mammals) (Gillespie, 1991; Marshall et al., 1991; Powell and Rogers, 1994). Unlike cytokeratins, or the specific fibrous keratins of hairs (hair keratins) or the other skin derivatives, keratin‐associated proteins form the amorphous, nonfibrous interkeratin matrix. The modality of biosynthesis, accumulation among cytokeratin bundles, molecular weight, amino acid sequence, and secondary structure of these small proteins are completely diVerent from those of cytokeratins. Evidence indicates that, despite the ‘‘b‐keratin pattern’’ of reptilian scales and claws, b‐keratins are in reality ‘‘keratin‐associated proteins’’ (KAPs). These proteins might have some amino acid homologies with those of mammals (Dalla Valle et al., 2005; Gillespie et al., 1982; Inglis et al., 1987; Knapp et al., 1991; Marshall and Gillespie, 1982; Powell and Rogers, 1994; Wyld and Brush, 1979, 1983). These proteins are packed in the cytoplasm of scale b‐layer cells (and in feather cells), among the cytokeratin framework, and eventually replace cytokeratins. Hard keratins of extant reptiles and birds diVer from those present in extant mammals, and possess diVerent chemicophysical and biochemical properties, such as molecular weight, amino acid composition, secondary conformation, degree of elasticity of stretching, packing modality, X‐ray diVractography pattern, solubility, and biosynthesis. The infiltration of mineral salts among keratin bundles has been reported in reptilian hard cornified structures such as claws and scutes (Alibardi, 2003a; Spearman, 1966). The biochemical contribution of minerals, and their molecular organization with proteins to increase the hardness of epidermal structures, remain to be further analyzed. In the following sections the three main types of KAP in reptiles, birds, and mammals are discussed.

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1. Reptilian b‐ or f‐Keratins In upper spinosus and precorneous keratinocytes of scales of reptiles, b‐keratins replace a‐keratins, suggesting that a‐keratins disappear or are degraded, diluted, or masked by the deposition of b‐keratin (Alexander, 1970; Alibardi, 2000, 2003a; Alibardi and Sawyer, 2002; Landmann, 1979, 1986; Maderson, 1985). Figure 10 shows the immunolocalization of b‐keratin in the epidermis of reptiles and birds. b‐Keratins have diVerent chemicophysical, biochemical, and cytological characteristics than cytokeratins (Baden and Maderson, 1970; Fraser and Parry, 1996; Fraser et al., 1972; Maderson, 1985; Sawyer et al., 2000; Wyld and Brush, 1979, 1983). These small proteins form a hard and inflexible layer, show little extensibility, impart durability, and increase the resistance of scales to mechanical stress. The above‐cited studies have indicated that these proteins possess amino acid regions organized in a b‐pleated sheet, aggregate into a densely packed lattice, and produce resistant microfibrils with a typical X‐ray and ultrastructural pattern of 3‐ to 4‐nm electron‐pale filaments in a denser matrix. b‐Keratins of reptiles and birds have a lower molecular mass (10–22 kDa) than cytokeratins (40–68 kDa) (Alibardi et al., 2000, 2001, 2004a,b; Fuchs and Marchiuk, 1983; Wyld and Brush, 1979, 1983). The result of aggregation of b‐keratin molecules produces resistant microfibrils and filament masses, initially termed b‐keratin packets, and, after they merge into tangled masses, b‐keratin filaments. The mechanism of aggregation of b‐keratins, whether due to these proteins or to the participation of (unknown) keratin‐associated proteins, is not known. The b layer in the epidermis of most reptiles (chelonians, crocodilians, but also many lepidosaurians) is also the depository of most epidermal pigmentation. DiVerent b‐keratins are present in reptilian epidermis, and they probably present a specific amino acid composition (rich in glycine), sequence, and spatial conformation (Table I). The cytology and synthesis of b‐keratins are presented in Section IV.A. Biochemical, immunocytochemical, and autoradiographic information about b‐keratin (Alibardi and Sawyer, 2002; Sawyer et al., 2000, 2005) has indicated that these proteins are heterogeneous. In diVerent reptiles, despite some common immunological cross‐ reactive epitopes three diVerent biochemical groups are likely: that of chelonians (turtles, terrapins, and tortoises), that of lepidosaurians (lizards, snakes, amphisbaenids, and sphenodontids), and that of archosaurians (crocodilians and birds). Immunocytochemical studies, using diVerent antibodies, have indicated broad (chicken scale b antibodies) or limited (turtle‐specific or lizard‐specific antibodies) cross‐reactivity among avian and reptilian b‐keratins. The broad immunoreactivity with b1 (Fig. 10), b‐general, and b‐universal antibodies indicates the presence of general, evolutionarily conserved epitopes in all

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FIG. 10 Immunocytochemical detection of scale b‐keratins in sauropsid scales (b1 antibody) and feather keratin (Fbk antibody) in barbs and barbules of a feather. (A) Tortoise (Testudo hermanni); (B) lizard (P. sicula); (C) setae of climbing scales of gecko lizard (Hemidactylus turcicus); (D) snake outer (arrowhead) and inner (arrow) b layer (N. natrix); (E) tuatara outer (arrowhead) and inner (arrow) layer (S. punctatus); (F) crocodile (Crocodylus porosus); (G) zebra finch scutate scale (Taeniopygia castanotis); (H) cross‐sectioned barbs of zebra finch feather (T. castanotis); (I) ultrastructural detail of immunogold‐labeled b‐keratin bundles (arrowhead) of b cell of lizard scale (P. sicula). Scale bars: (A and C–H) 10 mm; (B) 20 mm; (I) 200 nm. c, stratum corneum; h, hinge region; se, setae as a continuation of the oberhautchen/ b layer. Dashes underline the basal layer of the epidermis.

TABLE I Variation in Percentage of Five Representative Amino Acids in Hard Keratins of Reptiles and Mammals Amino acid (%) Species

Cys

Tyr

Ser

Pro

Glys

References

Turtle (Pseudemys picta)

6.2

10.4

6.7

8.3

32.8

Baden and Maderson, 1970

Tortoise (Chelonia midas)

5.5

12.7

5.6

10.1

30.3

Baden et al., 1974

Tuatara (Sphenodon punctatus)

4.0

—

9.9

9.5

29.7

Baden and Maderson, 1970

Boa snake (Boa constrictor constrictor)

6.8

—

6.6 Monitor lizard (Varanus gouldii)

219

Fraction V Low fraction

2.4

13.3

10.7

16.2

Baden and Maderson, 1970

14.7

15.9

18.6

Baden et al., 1974

7.8

13.9

7.3

9.5

23.5

Marshall and Gillespie, 1982

9.1

18.6

8.6

7.4

27.6

Marshall and Gillespie, 1982

6.9

4.1

7.5

7.2

12.8

Marshall and Gillespie, 1982

Fraction

13.0

4.1

7.5

9.1

28.2

Gillespie et al., 1982

Fraction

12.7

4.6

7.2

8.8

28.4

Gillespie et al., 1982

Fraction IV

12.5

4.8

4.6

10.7

31.4

Marshall and Gillespie, 1982

Fraction D

14.2

5.8

8.5

5.5

41.2

Marshall and Gillespie, 1982

Published primary sequence

16.0

2.1

9.0

8.0

29.0

Inglis et al., 1987

11.3

4.6

6.0

9.8

29.4

Frenkel and Gillespie, 1976

4.3

4.8

7.4

8.0

29.4

Dalla Valle et al., 2005

Monitorlizard (Varanus sp.) Lizard, scale keratin (Podarcis sicula) Chick beak (Gallus gallus)

4.6

9.4

6.3

9.6

33.9

Gillespie et al., 1982

Chick claw

4.9

9.0

7.1

9.4

30.8

Gillespie et al., 1982

Chick feather

8.3

3.9

10.7

11.1

15.6

Gillespie et al., 1982 (continued)

TABLE I (continued) Amino acid (%) Species

220

Cys

Tyr

Ser

Pro

Glys

References

Echidna quill (Tachyglossus aculeatus)

9.1

18.9

5.3

3.9

40.1

Gillespie et al., 1982

Type I, high‐tyrosine proteins of matrix fraction

6.0

15.1

11.9

5.3

27.9

Gillespie et al., 1982

Type II, high‐tyrosine proteins of matrix fraction

9.8

20.5

10.9

3.0

33.9

Gillespie et al., 1982

High glycine–tyrosine protein II (Mus musculus) hair

12.1

18.5

8.5

4.2

32.8

Marshall and Gillespie, 1982

Merino wool high‐sulfur protein (Ovis sp.)

18.9

2.1

12.7

12.5

6.9

Gillespie et al., 1982

Human high‐sulfur protein (Homo sapiens)

27.2

1.5

11.9

12.7

6.1

Gillespie et al., 1982

Mouse hair high‐sulfur protein

31.9

1.2

11.7

13.7

6.0

Gillespie et al., 1982

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b‐keratins (probably epitopes made of glycine‐rich amino acid sequences). The limited cross‐reactivity, obtained with antisera against lizard or turtle b‐keratins of 15–16 kDa, indicates that the latter polyclonal antibodies recognize subclass‐specific epitopes (for lepidosaurians, for chelonians, and for archosaurians; L. Alibardi and M. Toni, unpublished observations). The antibodies label the dense keratin bundles, made up of 3‐ to 4‐nm‐thick filaments, uniquely present in b cells (Fig. 10I). The variability of molecular weight patterns for b‐keratins is supported by detailed biochemical studies on keratins. The latter were extracted from the claw of a monitor lizard (Gillespie et al., 1982; Inglis et al., 1987; Marshall and Gillespie, 1982). Others came from geckos (Alibardi and Toni, 2005a,b; Thorpe and Giddings, 1981), lizard embryos (Sawyer et al., 2000), and regenerating epidermis (Alibardi and Toni, 2006; Alibardi et al., 2004a). Other keratins were derived from normal scales of snakes, lizards, chelonians, and crocodilians (Alibardi and Toni, 2005a,b; Alibardi et al., 2004b; Holmer et al., 2001; Wyld and Brush, 1979, 1983; and our unpublished observations). Bidimensional gel electrophoretic analysis of b‐keratins in all reptilian groups (exemplified by the lizard bidimensional pattern in Fig. 4) shows that three or four b‐keratin spots, mainly with basic pI, are present. b‐Keratins are therefore small, mainly basic proteins, typical characteristics for KAPs (see Section IV. D.3 on the evolution of glycine‐rich proteins). In a monitor lizard, two main fractions were found: 13‐ to 14‐kDa glycine– cysteine‐rich keratins and a minor fraction containing 8–20 proteins of 8–70 kDa. Among the latter, 7‐ to 8‐kDa keratins and a 13‐ to 14‐kDa keratin were glycine–tyrosine rich but cysteine poor, resembling mammalian or avian glycine–glycine‐rich proteins. The sequence of a 13‐kDa protein, constitutively expressed in the sodium dodecyl sulfate (SDS) component 1 fraction from lizard claw, was found to be 142 amino acids with a deduced acidic pI of 5.2. This protein contains regions homologous to both mammalian high‐ sulfur and high‐tyrosine proteins, and avian b‐keratins (Inglis et al., 1987). Chromatographic and electrophoretic analyses of claw proteins have shown the presence of other protein fractions (Gillespie et al., 1982; Inglis et al., 1987; Marshall and Gillespie, 1982). Among the diVerent fractions two main groups have been identified: (1) one is rich in lysine and glycine, and members resemble b‐keratin of avian claw and beak; (2) another resembles high‐tyrosine/glycine‐ rich matrix proteins of mammalian hard‐keratinized structures (or keratins). Conformational analysis of a 142‐amino acid lizard claw b‐keratin sequence showed that amino acid residues 36–67 determine a b‐pleated conformation that is the counterpart of amino acid 24–55 in the b‐keratin of emu feathers (Fraser and Parry, 1996). On the other hand, reanalysis of the secondary structure of this protein, using computer analysis programs, indicates that no canonical b‐pleated sheet is present. Instead, the presence of

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four‐strand conformations in the molecule may explain the origin of a b‐pleated sheet pattern on X‐ray analysis (Dalla Valle et al., 2005). Sequencing and cloning of the first reptilian scale keratin, from regenerating scales of a lizard (Dalla Valle et al., 2005), revealed that this is a 163‐ amino acid protein of 15.5 kDa, with a pI of 8.2. mRNA for this protein is localized in diVerentiating b cells, and ultrastructural in situ hybridization showed that the mRNA is also associated with forming b‐keratin packets (Fig. 11). The probe, against the whole 585‐nucleotide sequence of the b‐keratin mRNA, forms clusters or linear arrays of gold particles in the cytoplasm (see insets in Fig. 11). Also in this case, despite the presumed ‘‘b‐keratin pattern’’ for this b‐keratin, the computer program for secondary structure analysis indicates that no b‐pleated sheet is present in this protein. Most of the sequence is present as random coil conformation and three small regions present a strand conformation. The latter regions may be the source of the ‘‘b‐keratin pattern,’’ but possibly after homophilic aggregation with other molecules: this point is presently under analysis. The study of lizard scale protein reveals that the N‐ and C‐terminal regions share high homology with mammalian KAPs whereas the central block shows homology with feather and claw proteins. Computer phylogenetic analysis of this protein shows that it is closer to mammalian KAPs than are claw and feather keratins (M. Toni and L. Alibardi, unpublished observations). All avian and alligator b‐keratins, and that of lizard claw and scale, have a similar amino acid sequence from amino acids 39 to 53 (Dalla Valle et al., 2005; Fraser and Parry, 1996; Sawyer et al., 2000). Most b‐keratins share a common epitope of 22 amino acids, which is recognized by a specific antibody, called b‐universal (L. Alibardi and M. Toni, unpublished observations; Sawyer et al., 2000, 2005). A study of diVerent species of gecko (Thorpe and Giddings, 1981) showed bands at low molecular mass (11–12 and 15 kDa), belonging to a low‐sulfur fraction, whereas the high‐sulfur fractions showed higher molecular mass (16–22 kDa). This has been confirmed in other species of gecko (Alibardi and Toni, 2005b). These b‐keratins are present where elongated structures are formed, suggesting that they can associate into a tridimensional conformation that readily polymerizes into linear, long, and parallel cables supporting cell elongation. This occurs in gecko setae and in barbule cells of feathers (microfibrils of b‐keratins) (Fraser et al., 1972; Gregg and Rogers, 1986; Sawyer et al., 2003). Higher molecular weight scale b‐keratins cannot assembled into long‐axial cables to form finger‐like or elongated cells. Scale b‐keratins associate into b‐packets that merge irregularly to form a dense but irregular (not oriented) mass of keratin. Although genes for b‐keratins are becoming known (Dalla Valle et al., 2005; and unpublished polynucleotide and amino acid sequences), the chromosomal positions of these genes are presently unknown. The deduced

KERATINIZATION IN VERTEBRATE EPIDERMIS

223

FIG. 11 In situ hybridization detection of mRNA for a b‐keratin in b cells of regenerating lizard epidermis (P. sicula). (A) Immunofluorescence‐labeled fusiform cells of the diVerentiating b layer. Scale bar: 10 mm. (B) Ultrastructural view of clusters of gold particles (arrows) in the cytoplasm or near b‐keratin filaments (arrowheads) in a b cell. Scale bar: 200 nm. (C) High‐ magnification detail of cluster‐shaped cytoplasmic labeling. Scale bar: 100 nm. (D) High‐ magnification detail of linear cytoplasmic labeling. Scale bar: 50 nm. cy, cytoplasm; n, nucleus; w, wound epidermis. Dashes underline the basal layer.

proteins, completely or partially sequenced from lizard, snake, and turtle epidermis, reveal homologous but nonidentical amino acid regions (L. Dalla Valle, V. ToVolo, M. Toni, and L. Alibardi, unpublished observations). The general gene and protein structures found in lizard scale glycine–proline‐rich protein, with a central avian motif and two external mammalian motifs, are

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ALIBARDI

present in all the proteins of other reptiles. The emerging concept is that b‐keratin proteins are constitutive in all reptilian keratinocytes (b but also a), but their b‐keratin genes are expressed only in b‐keratinocytes. The molecular mechanism that controls the more or less continuous (chelonians and crocodilians) or cyclical (lepidosaurians) expression of b‐keratin genes is not known, but is probably of direct or indirect dermal origin (see Section IV, on dermal–epidermal interactions). The sequence homology of glycine‐rich proteins, derived from the determination of reptilian sequences, is not yet detectable in extant reptiles. The present sequence information, however, indicates that similar amino acid sequences were present in ancient glycine‐rich proteins of the epidermis of basic amniotes (see Section IV.D.3 on the evolution of glycine‐rich proteins). The preceding fragmentary information does not yet allow the tracing of a possible phylogenetic relationship between lizard b‐keratins and those of birds or with the hard matrix proteins in mammalian skin. Despite b‐keratin being phylogenetically more recent than a‐keratin (Maderson and Alibardi 2000), no information is presently available on the evolution of b‐keratin from a previous a‐keratin sequence. 2. Avian b‐Keratins Avian b‐keratins are hard proteins expressed uniquely in avian scutate scales, feathers, claws, beak, tongue tip, and barb‐bristles of avian epidermis (Brush, 1993; Brush and Wyld, 1980; Sawyer and Knapp, 2003; Sawyer et al., 2000, 2003, 2005). The genomic organization, protein structure, and nucleotide and amino acid sequences of avian hard keratins are well known (Frenkel and Gillespie, 1976; Gregg and Rogers, 1986; Gregg et al., 1983, 1984; Molloy et al., 1982; Presland et al., 1989; Whitbread et al., 1991). Avian b‐keratins comprise a large multigene family of small, glycine‐rich proteins. A group of at least 18 genes (20 proteins) for feather keratins, 4 genes for claw keratins, and 3 genes for claw‐like scale keratin, are localized in a linear array in the avian genome. At least another nine scale keratin genes (with a minimum of three expressed scale keratins) are present in the chick genome. All b‐keratin genes might have originated by duplication and mutation from one ancestral sequence. In general, avian b‐keratins proteins have been classified as scale (14–16 kDa), scale‐like (claw, beak, etc., 14–16 kDa), feather (10–12 kDa), and feather‐like (10–12 kDa) keratins (Sawyer and Knapp, 2003; Sawyer et al., 2003). The main diVerence between scale and feather keratins is the deletion of four repeats of 13‐amino acid, glycine‐rich regions present in scale keratin, to produce feather keratin. This observation has strengthened the hypothesis that feathers derived from the modification of archosaurian scales (Gregg et al., 1984; Maderson, 1972a; Spearman, 1966; Wu et al., 2004). However,

KERATINIZATION IN VERTEBRATE EPIDERMIS

225

the localization of a feather‐specific antibody (Fig. 10H) in embryonic scales of alligator and avian epidermis has indicated that feathers and scales may have diVerent evolutionary histories (Sawyer and Knapp, 2003; Sawyer et al., 2003, 2004). Feather‐like keratin is present in both scale and feather embryonic epidermis, thus suggesting that the simpler feather‐like keratin precedes during development the expression of larger b‐keratins (scale, claw, and beak; see Alibardi and Toni, 2004b; Sawyer et al., 2003, 2004). Whether the earlier expression means that feather‐like keratins precede the other b‐keratins is also indicated by a detailed study on feather development and regeneration (Alibardi, 2005a). A new member of the b‐keratin family, CK‐b‐keratin, has been found in avian keratinocytes growing in culture medium (Vanhoutteghen et al., 2004). This 13‐kDa keratin of 125 amino acids is diVerent from the above‐described b‐keratins. The presence of this protein in morphologically undiVerentiated avian keratinocytes further stresses the concept that b‐keratins are constitutive for avian keratinocytes (Sawyer and Knapp, 2003). Other lines of thought have instead indicated that feather keratins have evolved specifically in birds for feather formation, and have no relationships with those of most reptiles, or even with those of avian scales (Brush, 1993; Brush and Wyld, 1980). All studies agree that scale, claw, beak, and feather keratins are diVerent proteins. The smaller dimension of feather keratins indicates that this is a specially designed keratin, with chemicophysical properties adapted to produce filamentous proteins for the elongation of cells (barbules and barb) of feathers (Alibardi, 2002b, 2005c; Matulionis, 1970). Despite the loss of the central, 52‐amino acid glycine–glycine‐rich region, feather keratins remain glycine‐rich proteins with a lower percentage of glycine (13– 16%; see Table I). This indicates that these proteins belong to the generalized family of the glycine‐rich proteins of epidermal derivatives, although the sequences are diVerent (Table I).

3. Mammalian Hard Keratin‐Associated Proteins In epidermal derivatives of mammalian skin (horns, nails, hairs, etc.) numerous matrix proteins, present among the filaments of specialized cytokeratins (trichocytic keratins; see Section IV.C on hairs), have been found (Gillespie, 1991; Marshall et al., 1991; Powell and Rogers, 1994; Rogers, 2004). At present, matrix proteins are classified as intermediate filament‐associated proteins (IFAPs) or keratin‐associated proteins (KAPs) (Powell and Rogers, 1994). They are present among trichocytic keratins, mainly in the hair cortex, and a few are localized in the cuticle of hairs, where they are best known. However, the same or similar number of KAPs may be present in other derivatives (nails, claws, horns, scales, etc.).

226

ALIBARDI

In the hair cortex and cuticle more than 100 diVerent KAPs have been found. They are classified into three main types: high glycine–tyrosine (HGT‐ KAPs), high sulfur (HSP‐KAPs), and ultrahigh sulfur (UHSP‐KAPs). Between 35 and 60% of the amino acids of HGTs are glycine and tyrosine, and these proteins can be subdivided mainly into HGT I (poor in cysteine and rich in phenylanine, less soluble) and HGT II (rich in cysteine and poorer in phenylalanine, more soluble). Between 20 and 27 diVerent proteins are included in the mammalian HGT family. HSP‐KAPs include three families, members of which contain 30% or less of cysteine. UHSP‐KAPs include five families, members of which contain more than 30% (up to 55%) cysteine. According to current knowledge of the human genome, the genes for all these proteins are clustered in two main chromosome loci. Seventeen genes for HGTs and 7 genes for HSPs are localized in chromosome‐21q22; 37 genes for HSPs and UHSPs, and 7 genes for HGTs, are localized in chromosome 17q12‐2 (Rogers, 2004). The HGT proteins have a molecular mass ranging between 6 and 9 kDa, and are made up of 81–131 amino acids. Secondary conformation in these proteins is incompletely known, but they produce an a‐keratin pattern when studied in tissues or when isolated.

IV. Areas of Dermal–Epidermal Interactions and Epidermal Derivatives The dermis beneath the generalized epidermis is made of more or less homogeneously distributed mesenchymal cells. In some areas of embryonic skin, mesenchymal cells become morphologically and functionally diVerent from the surrounding cells, and form the dermal papillae of skin derivatives (some scales, hairs, feathers, etc.). Dermal–epidermal interactions during embryogenesis determine the formation of scales, hairs, and feathers (Chuong and Widelitz, 1999, 2000, 2003; Hardy, 1992; Millar, 2002; O’Guin and Sawyer, 1982; Olivera‐Martinez et al., 2004; Philpott and Paus, 1999; Sengel, 1975, 1986; Sire and Huysseune, 2003; Widelitz et al., 2003; Wu et al., 2004). These skin derivatives are capable of cyclical regeneration (e.g., hair cycling, feather replacement, and scale molting). Comparative studies on the distribution and shape of regions where dermal–epidermal interactions occur have indicated that these interaction are not randomly distributed in the epidermis but instead are organized in areas of dermal–epidermal interactions (ADEIs; Alibardi, 2004c). The continuity of the basement membrane is partially interrupted, so that dermal–epidermal interactions occur via direct contact by cell bridges or by

KERATINIZATION IN VERTEBRATE EPIDERMIS

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intimate contact, with production of active molecules exchanged between dermis and epidermis. The best known ADEIs are those of hairs and feathers. The mesenchyme underneath the outer scale forms the ADEIs of scales, whereas this is not the case for the germinal epithelium of the inner scale surface and hinge region. This also occurs between the dermal papilla and collar cells of feathers or in the dermal papilla and matrix cells of hairs. The above‐cited studies have shown that ADEIs are characterized by a specific pattern of distribution of extracellular matrix proteins, adhesion and signaling molecules, and growth factors and their receptors. The dermal and epidermal components of ADEIs move in relation to each other during development (Alibardi, 2004c). Variations in the extension of these regions may form dermal condensations and influence the morphogenesis of skin appendages, determining the formation of scales, hairs, feathers, nails, hooves, and other derivatives. The location of ADEIs on the skin of diVerent vertebrates and their modifications (reduction, extension, and shaping) might have determined the origin and evolution of epidermal appendages (Figs. 12–14) (Alibardi, 2004c). This hypothesis is presented in the following sections. The scaled pattern of the skin of sarcopterygian fish originated the scaled skin of ancient amphibians and basic amniote amphibians of the Carboniferous Period (Chudinov, 1968; Romer and Witter, 1941) (Fig. 12A and C). Modern amphibians lost scaled ADEIs, which might have become internalized around dermal glands (Fig. 12B). The scale pattern in the skin of ancient amphibians was replaced by a patterned or localized distribution of glands. During the evolution of synapsid reptiles in the upper Paleozoic Era, and of therapsids in the Mesozoic Era, ADEIs became progressively reduced in surface and internalized into the dermis to generate hairs and vibrissae (Fig. 12C–G). The conformation of hair ADEIs is represented by the dermal papilla associated with the surrounding hair germinal epidermis (Fig. 13J). Other derivatives such as nails, claws, hooves, and horns might also have derived from complex disposition of ADEIs (Fig. 12D and D1–4). The epidermis of these derivatives produces specific proteins for hard cornification, probably in relation to the dermal influence on the epidermis. The evolution of reptilian diapsids and anapsids in the upper Paleozoic and Mesozoic Eras was characterized by extended and superficial ADEIs that originated scales, claws, and beaks (Fig. 12I and I0 ). Interaction between the dermis and epidermis is directly or indirectly responsible for the formation of a laminar layer of hard or b‐keratin. In archosaurians, ADEIs became progressively reduced and internalized in the dermis to form feathers (Fig. 12J–M). The conformation of the feather ADEI is represented by the dermal papilla contacting the collar epidermis (Fig. 13B and D).

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FIG. 12 Drawing summarizing the distribution of various proteins in the epidermis of vertebrates during the evolution of epidermal derivatives (scales, feathers, and hairs; see text). (A) Fish scales. (B) Amphibian epidermis. (C–H) Hypothetical evolution of hairs from the reduction of large areas of dermal–epidermal interactions (ADEIs) into small intradermal areas in hairs. (C and I) Hypothetical evolution of reptilian scales from enlarged ADEIs. (J–M) Evolution of feathers from the reduction of large ADEIs of scales into a truncated cylinder (collar) among large apteric areas. Complex ADEIs have produced other epidermal appendages

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Specific proteins were produced from the epidermis of these derivatives, probably under direct or indirect dermal influence. Scales, hairs, and feathers of extant amniotes are unique derivatives, and they have followed diVerent evolutionary histories from a scaled, diVerent integument of basic amniotes (Fig. 12).

A. Reptilian Scales In reptilian scales ADEIs are large and have roughly the shape of their outer surfaces (Fig. 12I and J). Large or small, flat or tuberculate, nonoverlapped or extremely overlapped scales in reptiles can be structurally divided into two main types: those of chelonians and crocodilians and those of lepidosaurians. These diVerent scales have been previously described in the sections on reptilian epidermis. The typical protein of scales of both reptiles and birds, and of feathers, is b‐keratin. These keratins have common epitopes among all reptilian (and avian) b‐keratins that are recognized by broad‐spectrum antibodies (Fig. 10). During terminal diVerentiation of b‐layer cells, b‐keratin filaments coalesce into a homogeneous mass and most of the other organelles and cytoplasm disappear. In lepidosaurians b cells tend to merge into a syncytium and a compact b layer is formed. In Sphenodon, crocodilians, and chelonians b cells remain initially separated in the lower part of the corneous layer but for the cell membrane, and are partially digested in the upper part. The deposition of electron‐dense material of unknown composition onto the plasma membrane is frequently observed. The formation of a b layer in reptilian scales is directly or indirectly induced by the mesenchyme of ADEIs during embryogenesis, normal cycling, or scale regeneration (Alibardi, 2003a, 2004c). This is indicated by the intimate dermal–epidermal contacts that occur exclusively in this area and by the scaleless mutant (Licht and Bennett, 1972). In the latter case, the lack of mesenchyme under the epidermis correlates with lack of b‐layer formation. The diVerentiation of b cells is also induced when avian or mammalian dermis is combined with epidermal epidermis in organ culture (Dhouailly, 1975). Therefore, like in avian scales (O’Guin and Sawyer, 1982), reptilian scales in which a b layer diVerentiates are also considered skin appendages (in the sense of Maderson, 1972a). in mammals (D1–D4) and birds (I0 and M). Fbk, feather b‐keratin; cADEI, condensed areas of dermal–epidermal interaction; HGP, high glycine–tyrosine proteins; HK, hair keratins; HRPs, histidine‐rich proteins; HS, high‐sulfur proteins; KAPs, keratin‐associated proteins; KI, keratins of the inner root sheath; KO, keratin of the outer root sheath or epidermal keratins; LOR, loricrin; TG, transglutaminase; TH, trichohyalin; UHS, ultrahigh‐sulfur proteins.

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FIG. 13 Drawing illustrating the organization of areas of dermal–epidermal interaction (ADEIs) in feather (A–D) and hair follicles (F–J; see text). (A–C) Development of the first feather generation (down feathers). (A–E) Development of the second and successive feather generations (juvenile and adult feathers). The ring‐shaped ADEI of feather (collar and dermal papilla) interacts with inductive fibroblasts. In this case epidermal cells proliferate along the ring and remain separated from the central dermal papilla to form folds (barb ridges). In down feather barb ridges remain separated and reach the collar at the same point, producing separated barbs inserted into the calamus. In pennaceous feathers barb ridges merge into a rachis that grows axially, producing the planar vane. In hair the ADEI is shaped as a inverted cup (J) containing the dermal papilla interacting with the hair placode. DiVerently from the ring‐shaped collar of feather, epidermal cells of the cup‐shaped epidermal placode proliferate around the tip of the dermal papilla and coalesce internally above the papilla, forming the hair

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Other experiments on lizard skin maintained in an in vitro medium have shown that the epidermis, without direct contact with the dermis, is capable of b‐keratin diVerentiation and production of successive epidermal generations (Flaxman et al., 1968). However, the culture medium contained serum and chick extracts so that numerous growth factors and unknown dermal factors were actually continuously present in the medium. These dermal factors might have been responsible for triggering the mechanism of cycling of the epidermis: this would indicate that direct dermal–epidermal contact is not needed for epidermal diVerentiation in lizard epidermis. It is likely that, as in all the other derivatives in vertebrates, in the case of reptilian scales a dermal influence is also needed to stimulate cell diVerentiation.

B. Feathers and Their Evolution Among avian derivatives feathers are the most complex (Lucas and Stettenheim, 1972). Modern scales and feathers have probably evolved from ancient archosaurian scales (Chuong and Widelitz, 1999; Chuong et al., 2000, 2003; Maderson, 1972b; Prum, 1999; Prum and Brush, 2002; Spearman, 1966; Wu et al., 2004). The reduction of ancient scales might have produced apteric areas (Alibardi, 2003a, 2005c) (Fig. 12J–L). The pliable and stretchable apteric and interfollicular epidermis transformed the rigid and scaled reptilian armor into a dynamic skin to favor feather movements. The feather areas (or tracts) resulted from the formation of localized rows of feather germs that grew into filaments (Fig. 12K, K0 , and L). Although the ADEI was initially localized in all the surface of germs, the ADEI became restricted at the base of the growing feather filament (Alibardi, 2004c). The ADEI eventually sank into follicles and became internalized with a ring shape (the collar) (Fig. 13B and D). The conformation of this ADEI and the intense proliferation of the upper part determined the increase in epithelial area and the formation of folds or barb ridges (see Figs. 13B and 14). The latter permitted the evolution and diversification of feathers (Alibardi, 2005a), as is indicated in the following hypothesis. From tuberculate scales hair‐like appendages evolved (Fig. 14A and B), which progressively lengthened into feather filaments containing a dermal,

cone (F and G). The latter is surrounded by the inner root sheath. The surrounding cells of the hair peg (in gray) have been removed for visualization of the hair cone. B1 and B2, barb ridges 1 and 2 (merging); BA, barb; BL, barbules; BR, barb ridges; CO, collar; CX, cortex; CU, cuticle; DP, dermal papilla; E, epidermis; FB, active fibroblasts of the ADEI; FO, follicle; GH, growing hair; HP, hair placode; IRS, inner root sheath; MB, merging barb ridges; ORS, outer root sheath; R, rachis.

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FIG. 14 Drawing presenting the hypothetical evolution of down feathers from a tuberculate scale (A) to a pointed outgrowth (B), and an elongated, hair‐like feather filament (C) with dermal core (see text for details). After the formation of barb ridges inside the feather filaments (D), the diVerent displacement of subperiderm cells to form the biplanar branching of a typical/ modern down feather is shown (G–G2). Other possible barb ridge phenotypes are represented in (E)–(F2). The process of fusion of barb ridges into a rachis (H) originates the pennaceous feathers (H1). AP, axial plate; BA, barb; BL, barbule; BP, biplanar barb ridge; BR, barb ridge; BAP, beginning of formation of the axial plate; BRO, barb ridge outline (dashes indicate disappearing barbules); CBP, condensed barb/barbule plate; CO, collar; E, subperiderm cells

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vascularized core (Fig. 14C). Along the perimeter of the epidermis periodic folds were formed, originating the barb ridges. The epidermal folding derived from the resistance of the sheath over the internal expansion of the epidermis, which folded toward the less resistant, central mesenchyme. The formation of regularly spaced barb ridges represented an evolutive innovation in mechanism of epidermal morphogenesis and keratinization that led to the origin of feathers. Only detailed knowledge of the cell structure and three‐dimensional organization of barb ridges during feather development allowed an understanding of feather evolution (Alibardi, 2005c). A new hypothesis on feather evolution may be based on a variation of the process of morphogenesis in barb ridges. The embryonic epidermis consists of an external or primary periderm, a secondary periderm, a subperiderm layer, and the basal or germinal layer. The sequence and stratification of epidermal layers in developing feathers are altered by the formation of barb ridges, from which barb and barbules are later formed. Barb ridge simply represent a fold of the epidermal sheet inside a narrow cylinder, terminating in a dome‐like end (Fig. 14C). Under these conditions it is likely that the epidermal cells will take a conformation that reduces the ‘‘tension’’ between the epidermal cylinder and the domed surface at the tip of the hollowed tube. This mechanical stress determines the formation of epidermal folds inside the apical part of the hollowed tube that increase the epidermal surface within the cylinder. Barb ridges form by the tip of the feather filaments and propagate down toward the base of the feather filaments, near the collar region (Fig. 13A). Barb ridges remain independent and isolated barbs form a plumulaceous feather (Fig. 13A–C). In each barb ridge only the original four embryonic layers remain: primary and secondary periderm, subperiderm, and basal layer. The secondary epidermis is distorted by the folding process and forms the two to four layers of cells of the sheath and, likely, supportive cells (barb ridge vane cells). These cells store a common organelle, the periderm granule. Cells of the subperiderm are displaced by the folding movement but tend to remain contiguous and form the alar or barbule plate and the central, median, barb area (Fig. 14D1). The basal layer stops proliferation and forms the marginal plates of barb ridges, destined to degenerate. When barb ridges

concentrating into a unique central mass within the barb ridge; E1, derived unbranched barb; E2, down feather without barbules; FO, follicle. (F) Initial stage with most centrally aggregated subperiderm cells within the barb ridge; (F1) intermediate stage with the formation of a central barb and short barbules; (F2) resulting down feathers with short barbules; (G) initial bilateral displacement; (G1) derived bilateral branching; (G2) down feather (only 3 barbs attached to the collar are represented; there can be more than 10 barbs in chick down feathers). MC, mesenchyme; MV, mesenchyme with blood vessels; RA, rachis; RM, ramogenic zone; S, sheath; SBL, short barbule.

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begin to merge before reaching the collar a rachis and a pennaceous feather are formed (Fig. 13A, D, E, and H). Feathers may be derived from tuberculate or coniform scales present in preavian archosaurian reptiles (Maderson, 1972a; Prum and Brush, 2002; Wu et al., 2004) (Fig. 14A and B). The latter were hairy‐like scales that perhaps performed an insulatory role for thermoregulation (Fig. 14C). Inside these filaments the epidermis started to form foldings from which barb ridges originated. The morphogenesis of barb ridges was an evolutionary novelty necessary for the origin of feathers in birds and therapods (Alibardi, 2005a). Barb ridges are bilaterally symmetric epidermal folds (Fig. 14D1). After the degeneration of supportive cells among barbules and of the sheath, barb ridges open up to form the branched structure made of a central barb and lateral barbules (Fig. 14F–F2). As a result branched down feathers are formed. Barb and barbules are formed contemporaneously, and only an alteration of this scheme of morphogenesis produced feathers with barbs and no barbules. DiVerent feather phenotypes, such as those seen in fossils, might be derived from alternative morphogenetic processes of cell displacement within barb ridges. For instance, the fusion of barbule plates with the central barbs (condensing axial plate) could have produced the partial (Fig. 14F–F3) or complete (Fig. 14G–G2) disappearance of barbules in down feathers. The simple branching in primitive feathers found in ancient fossils such as Sinosauropteryx, Beipiaosaurus, Shuvuuia, and Sinornithosaurus (Prum and Brush, 2002; Wu et al., 2004) could be due to this process. All these plumulaceous feathers served an insulatory and thermoregulatory function, the primary role for feathers. The fusion of barb ridges to form a rachis and a pennaceous vane originated pennaceous feathers (Fig. 14H and H1). Modulation of the pattern of barb ridge formation in the follicles of regenerating feathers was responsible for the production of the diVerent types of pennaceous feather (semiplumes, contour, bristles, filoplumes, etc.). Finally, the formation of hooklets permitted the formation of close vanes with aerodynamic properties, later exploited for flight.

C. Hairs and Their Evolution The progressive reduction of ADEIs of synapsids into the dermis of therapsids might have originated the mammalian pelage (Fig. 12C–H). The specific conformation of ADEIs at the base of hair pegs and definitive hairs (Fig. 13I and J) determines the confluence, fusion, and growth of cells from the hair placode epidermis (matrix zone) into a central mass. This produces the hair shaft made of cortical (medullary in some hairs) and cuticle cells (Fig. 13F, G, and I). The latter are surrounded by cells of the inner root

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sheath (IRS), which are also produced from the more external cell columns of the hair matrix (Orwin, 1979; Rogers, 2004; Stenn and Paus, 2001). It is probable that, as a result of the dermal–epidermal interaction (Botchkarev and Paus, 2003; Millar, 2002), diVerent genes are activated in cells that belong to the various epidermal columns (or better cones) originated from the matrix. These genes encode specific trichocytic keratins in the cortex, other keratins in the cuticle, IRS, and companion layer cells (Langbein and Schweizer, 2005; Powell and Rogers, 1994; Rogers et al., 1999). Furthermore, special matrix or hard keratin‐associated proteins are sequentially produced in the proximal to distal direction along the cortex and cuticle. At first trichocytic keratins are synthesized, and then high glycine–tyrosine‐rich protein (HGT) followed by high sulfur‐rich proteins (HSPs), and by ultrahigh sulfur proteins (UHSP) produced in the cortex and in the cuticle (Fig. 12H). Within each main class of proteins diVerent subsets of genes for HGT, HSP, and UHSP are activated at progressive levels of the diVerentiating hair fiber up to the maturing region. The formation of papillate epidermis, commonly found only in the mammalian epidermis or in the context of diseases such as psoriasis, suggests that the epidermis has the potential to migrate into the dermis. This morphogenetic process may represent a preadaptation for hair bulb formation. The stratification of cells around the growing cone of the hair (the forming inner root sheath, IRS) suggests that these cells serve to carve out the hair canal or/and allow the upward movement of the hair fiber within the mass of epidermal cells of the hair peg (Fig. 13F and G). A shedding or protective layer destined to be sloughed around the hair fiber would avoid the creation of discontinuities in the epidermis excluding water‐loss and inflammatory reactions. The variation in the number of layers in the IRS is probably related to the papilla and to hair size and shape. Normally one cuticle, one Huxley layer, and one Henle layer (the more external) are present. It is known that large hairs often have a thicker Huxley layer than that of thinner hairs. The Huxley, Henle, and external companion layers participate in the formation of the slippage plane along which the hair fiber moves to exit through the epidermal surface (Langbein et al., 1999; Orwin, 1979; Stenn and Paus, 2001). 1. Hair (Trichocytic) Keratins The outer coverage of hairs, the outer root sheath (ORS), is mainly an extension of the epidermis, and presents a similar pattern of keratin distribution (Fig. 15A). Acidic and basic keratins stain most of the layers, although the K1/K10 pair tends to stain the corneous layer lining the hair canal. These a‐keratins have a basic molecular structure similar to that epithelial cytokeratins, with a central rod domain and an N‐ and C‐external domain (Gillespie, 1991; Powell and Rogers, 1994). The latter, however, is diVerent from that of

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FIG. 15 Immunocytochemical detection of hair proteins. (A) AE3‐immunofluorescent outer root sheath around the inner root sheath and the negative hair shaft in cross‐sectioned wombat hair (Vombatus ursinus). Scale bar: 10 mm. (B) Trichohyalin‐immunoreactive inner root sheath and medulla of cross‐sectioned large mole hair (Talpa europaea). Scale bar: 20 mm. (C) Longitudinally sectioned cat hair showing immunofluorescent medulla and inner root sheath (arrow) for trichohyalin (F. catus). Scale bar: 10 mm. (D) Obliquely sectioned rat hair showing immunoreactive medulla (arrowhead) and inner root sheath (arrow) for trichohyalin. Scale bar: 20 mm. (E) Ultrastructural detail of immunolabeled trichohyalin granule (arrowhead) of echidna hair (T. aculeatus). Scale bar: 100 nm. bu, hair bulb; h, hair; k, keratin filaments; i, inner root sheath; m, medulla; o, outer root sheath.

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cytokeratins, as the glycine‐rich extended V regions are replaced by cysteine‐ rich E regions. In the latter the secondary conformation is modified with respect to the V regions and more intermolecular disulfide bonds are formed during polymerization. Initially in human and bovine hairs four main type I (acidic) and four type II (basic) trichocytic keratins were found in the cortex and cuticle, and less abundantly in the medulla (Heid et al., 1988). Two minor components of types I and II were also found mainly in nails, filiform papillae of the tongue, and thymus. More detailed studies of human hairs eventually defined nine type I (acidic) trichocytic keratin members and six type II (basic) members (Langbein and Schweizer, 2005; Langbein et al., 1999, 2001). The pairing of type I and type II appears more random than the stricter pairing of epidermal cytokeratins. These hair cytokeratins are localized in human chromosome 17q12‐21 (type I) and in chromosome 12q13 (type II). Other, more specific keratins (hK6 and hK16) have been found in the inner root sheath (Bawden et al., 2001; Langbein et al., 2001; Winter et al., 1998) and in the companion layer that surrounds the Henle layer (Mahony et al., 1999; Rothnagel and Roop, 1995). Nail contains cytokeratin of epidermal and hair types, sometimes coexisting (Heid et al., 1988). Hair keratins have been found also in other cell types, such as those of the filiform tongue papillae and epithelial cells of the thymus (Heid et al., 1988). Gene activation for the synthesis of hair keratins occurs early in the matrix zone of the hair bulb, before activation of genes for keratin‐associated or matrix proteins (Aoki et al., 2002; Powell and Rogers, 1994). 2. Hair Trichohyalin Aside from specific cytokeratins, the inner root sheath (IRS) presents a peculiar interkeratin‐associated or matrix protein: trichohyalin (Rogers, 2004; Rogers et al., 1999; Rothnagel and Rogers, 1986). This basic protein of 190–220 kDa in diVerent mammalian species, which grows in sheets, is made up of 458 amino acids, among which prevail glutamate, arginine, glutamine, and leucine (Fietz et al., 1993; Hamilton et al., 1993). The protein has common epitopes in all mammals, from monotremes to marsupial– placentals. It was probably the matrix protein selected in therapsids and ancient mammals before the splitting between prototherians and eutherians for the special form of cornification of the IRS around the hair fiber (Alibardi, 2003a, 2004d; Rogers et al., 1999) (Fig. 15B–D). Trichohyalin is the main protein of the cornified matrix of the IRS and medulla (Rogers, 2004; Rogers et al., 1999; Rothnagel and Rogers, 1986; O’Guin and Manabe, 1991). The protein is concentrated in trichohyalin

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granules, which are large (0.2–3 mm) and dense organelles, formed by a meshwork 10–12 nm thick (keratin) associated with trichohyalin. Initially, in the diVerentiation of IRS cells, trichohyalin granules enlarge and occupy large portions of the cytoplasm of Huxley and Henle layer cells. These granules quickly disappear in cornifying cells and are replaced by a fine cornified matrix. Immunolabeling for trichohyalin is reduced or absent in the corneous cytoplasm of IRS cells, and limited to only a few areas where the 10‐ to 15‐nm‐ thick coarse filaments are still irregularly organized (Fig. 15E). During the process of cornification of the IRS in all mammalian species, the special keratin filaments organize themselves into parallel rows, whereas trichohyalin is dispersed among them and cross‐linked by the transglutaminase present in maturing IRS (Steinert et al., 2003; Taresa et al., 1997). Transglutaminase localization in the IRS probably indicates terminal diVerentiation before full cornification of the IRS. The enzyme arginine deiminase determines the transformation of trichohyalin into a form that loses its interaction with keratin, and is then degraded during cornification of IRS cells. The IRS possesses a diVerent consistency with respect to the hair cuticle, based on special trichocytic keratins, high sulfur proteins, and disulfide bonds formed by the action of sulfhydryl oxidases (Langbein et al., 1999, 2001; Marshall et al., 1991; Rogers, 2004; Rogers et al., 1999). The above diVerence determines the detachment of the hair fiber from the IRS along the serrated interface of the fiber cuticle and the IRS cuticle. The two cuticles separate one from the other after cell junctions are degraded by enzymes and by sebaceous secretions (Orwin, 1979; Stenn and Paus, 2001). Trichohyalin in the IRS is enzymatically degraded in the sloughing zone of hair (near the exit of the duct of the sebaceous gland), whereas sulfur‐rich proteins of the cuticle are resistant to the lytic enzymes (O’Guin and Manabe, 1991; Orwin, 1979). The origin of the IRS was probably essential during the evolution of hairs as a shedding layer and the slippage for hair exit. The stratification and retarded cornification of IRS cells have some functional analogies with the shedding layer of lizard scales. In the latter, a particular epidermal tissue, the lacunar layer, is sandwiched between the cornified layers of scale epidermis and is located in a ‘‘slippage plane’’ for the lateral growth of scales. The lacunar layer forms a plastic cushion during the distension of the new growing epidermis underneath the old epidermis of scale before molt. Beneath the lacunar layer, a specialized shedding layer is formed. In both IRS and the shedding layer of reptiles the continuity of the epidermis is maintained and infection/inflammation is avoided. 3. Hair Evolution The integument of mammalian‐like reptiles of the Permian Period might have progressively reduced scales and the skin became smooth (Chudinov, 1968;

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Findlay, 1968; Maderson, 1972a; Spearman, 1964). The epidermis probably lacked a granular layer but the progressive accumulation of HRPs originated some keratohyalin granules that transformed the transitional layer into a granular layer (Alibardi, 2003a; Alibardi and Maderson, 2003c). A primitive skin patterned into scales explains the derivation of regularly distributed epidermal appendages such as hairs. The progressive reduction and migration of ADEIs into the dermis and the reduction into smaller units determined the progressive loss of scales. Internalized ADEIs might have induced the formation of a dermal papilla near the former hinge region (Fig. 12D, D0 , E, and E0 ). Dermal papillae with inverted cup geometry (Fig. 13J) induced cell proliferation at the base of epidermal pegs with the formation of hair‐like outgrowths (Fig. 13F–H). Shedding of the epidermis around the growing hair fiber was necessary to avoid damaging the continuity of the epidermis with consequent inflammation and microbe invasion. Cells around the growing hair formed the IRS by means of a protein (trichohyalin) producing a diVerent corneous material with respect to that of the epidermis. Trichohyalin might have derived from duplication and evolution of the gene for profilaggrin (Fietz et al., 1993). The known profilaggrin and trichohyalin genes are closely localized in the same chromosomal epidermal diVerentiation complex, a cluster of genes encoding several epidermal proteins and that share some common sequences (Fietz et al., 1993; Taresa et al., 1997). Trichohyalin and filaggrin coexist in some epidermal areas or in mixed granules, suggesting they are coexpressed in the same keratinocytes (Hamilton et al., 1993; Manabe and O’Guin, 1994; O’Guin and Manabe, 1991). Filaggrin and trichohyalin might have been initially mixed in the hair canal. Then trichohyalin was selected in keratinocytes of the hair canal and produced a type of cornification diVerent from that present in the epidermis. This diVerence allows the separation between epidermis and the hair canal. The last modification was the formation of an inverted, cup‐like dermal papilla that favored the growth of keratinocytes into a central, rod‐like hair (Figs. 13F–H and 16F and G) (Alibardi, 2003a, 2004d).

D. Evolution of the Stratum Corneum and Epidermal Derivatives The epidermis of extinct vertebrates is renewed by more or less continuous activity with loss of the external layers (fish, anapsid and archosaurian reptiles, birds, and mammals). In other vertebrates periods of rapid proliferation (renewal phases), ending in a process of molt, alternated with periods of resting (terrestrial amphibians and lepidosaurian reptiles) (Budtz, 1977; Maderson, 1985; Maderson et al., 1998). Molting is possible because of the

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FIG. 16 Drawing illustrating the hypothesized evolution of hard keratins (b‐keratins and keratin‐associated matrix proteins) from the duplication and diversification of the gene (or exon) encoding the glycine‐rich V regions of an a‐keratin (see text for further explanation). E1 and E2, E regions; H1 and H2, H regions; HGP, high glycine–proline‐rich proteins; HGPT, high glycine–proline–tyrosine‐rich proteins; HGSP, high glycine–cysteine–proline‐rich proteins; HSP, high‐sulfur proteins; UHSP, ultrahigh‐sulfur proteins; RD, rod domain; V1 and V2, V regions.

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formation of weak, nonuniform intraepidermal regions (e.g., made by a diVerent modality of keratinization), separated by more or less specialized shedding layers. A shedding mechanism operates when structures formed inside the epidermis (neogenic scales, hairs, glands, etc.) must exit on the epidermal surface without producing barrier disruption. Shedding complexes evolved independently along diVerent amniote lineages (lepidosaurian reptiles, birds, and mammals) but was already present in amphibians, the first tetrapods. Shedding or scission layers are also formed in the turtle carapace and plastron of some species (Alibardi, 2005a), and in avian scales (Spearman, 1966). Also, molting hairs and feathers present shedding zones formed during catagen or regressive phases. By this mechanism, the continuity of the epidermis is maintained, water loss is minimized, and infection/inflammation is avoided. In mammalian epidermis, a mechanism of shedding works along the slippage plane between the hair fiber, the inner root sheath, and the epidermis on top of the hair canal (Alibardi, 2004d; Orwin, 1979; Stenn and Paus, 2001). A similar phenomenon of delayed cornification is found in the stratified lacunar cells of the epidermis of lizards and snakes before molt (Landmann, 1986; Maderson, 1985). Lacunar cells form within the epidermis a stratified layer that is also localized between two cornified layers: the corneous layer of the outer epidermis and that of the underlying inner epidermis. Along this interface the inner epidermis is expanding horizontally (growing) to form a larger, new scale surface underneath the old one (Maderson et al., 1998). The presence of incompletely keratinized and stratified cells in the lacunar layer probably gives plasticity to the expanding movements and ensures the continuity of the epidermis in case of microfractures. A shedding layer with a serrated grip similar to that of the inner sheath cuticle and fiber cuticle cells is also formed in the growing area of reptilian scales. The formation of intraepidermal shedding layers (that may involve cell death) allows a perfect sculpturing of epidermal structures in relation to progressive body growth. The diVerentiation of programmed shedding layers in conjunction with the action of immune cells in vertebrate epidermis may represent a physiological process that avoids pathological damage to the skin such as inflammatory responses and microbial invasion. With the previous background we are in the position to conclude our discussion with some hypothesis on the evolution of the process of keratinization and cornification of the epidermis during the transition from water to land, and its following adaptation. 1. Initial Stages of Land Transition from the Aquatic Phase The establishment of an eYcient stratum corneum in land vertebrates in the Carboniferous Period (basic amniotes) occurred by numerous changes from

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the epidermis of amphibians (Alibardi, 2003a; Matoltsy, 1987; Toledo and Jaret, 1992). While ancient amphibians (stegocephalians; Pough et al., 2001; Romer and Witter, 1941) were scaled, modern forms (Lissamphibia) lost scales but developed dermal glands to protect the epidermal surface (Fig. 12B). Aside from the multilayered stratum corneum, new cell organelles and molecules also appeared during the transition from the scaled epidermis of amphibians to that of amniotes: coarse filaments, vesicular or lamellar bodies, matrix proteins, and numerous proteins for the formation of the cornified envelope (the ‘‘protective system’’; Matoltsy, 1987). These basic modifications in the protective system of the general epidermis increased the vital functions for survival on land: eYcient in limiting water loss and enhanced mechanical and antimicrobial protection of the skin in the terrestrial environment. The fossil record indicates that basic amniotes possessed a scale integument (Chudinov, 1968; Pough et al., 2001), probably inherited by their piscine ancestors (Fig. 12A and C). The first basic amniotes probably possessed an epidermis made of more layers of dead horny cells accumulated on the surface, like in the wall of amphibian cocoons. The stratification increased the eYciency of the protective system but the epidermis lost completely its respiratory capability: the improvement of respiratory and circulatory systems in amniotes overcame this loss. A process of terminal diVerentiation was present and corneocytes were lost by desquamation or by periodic shedding. The mucous epidermis of amphibians was replaced with a dry epidermis in amniotes. In both sauropsid and therapsid amniotes the ubiquitous cytokeratins of the epidermal cells increased their complexity and genomic makeup, so that more specialized keratins were formed in comparison with those present in aquatic and amphibian vertebrates (Luke and Holland, 1999; SchaVeld and Markl, 2005). Cytokeratins became more and more basic and hydrophobic, especially for the lengthening of their variable (V) regions (Klinge et al., 1987; Steinert and Freedberg, 1991). However, from the duplication and divergent evolutions of the exons encoding these hypervariable regions, glycine‐rich regions of shorter length than in cytokeratins could be formed in both sauropsids and theropsids (Fig. 16). Examples of glycine‐rich regions are underlined in Fig. 16. These regions are present in proteins of diVerent localization and species: in V1 and V2 regions of a basic human cytokeratin, in lizard scale b‐keratin, in monitor lizard (Varanus) claw keratin, in sheep hair high‐glycine tyrosine, in human hair high‐sulfur protein, a chick scale keratin, and a feather keratin. Despite the diVerences all these proteins contain a high percentage of glycine that gives hydrophobic and insoluble characteristics to form resistant protein aggregations. These glycine‐rich, small proteins could initially combine with other glycine‐rich proteins present in corneous cells or with matrix proteins: this

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combination strengthened the corneous material and the cornified envelope of superficial corneocytes. The molecular evolution and diversification of new types of cell corneous proteins (involucrins, loricrins, etc.) were probably underway, but only knowledge of the nucleotide and amino acid sequences of these proteins in diVerent vertebrates, especially reptiles, will allow tracing of their molecular diversification and derivation. Characteristic of an apoptotic or terminal diVerentiation program of the epidermis (Haake and Polarowska, 1993) is the nuclear condensation and production of basic proteins (HRPs) or sulfur‐rich proteins for keratohyalin formation (Fukuyama and Epstein, 1986; Ishida‐Yamamoto et al., 2000). Nuclear loss and keratohyalin formation coincide with the transition from a parakeratotic to an orthokeratotic horny layer. 2. Origin of Epidermal Derivatives Two main types of evolving transformation progressed from the Carboniferous Period to the Mesozoic Era and originated the scaled integument and feathers in sauropsids and a pelage in therapsids (Maderson, 1972a,b). The appearance of epidermal derivatives such as feathers, hairs, nails or claws, horns, beak, and so on, in vertebrate history was, however, determined by the establishment of specific areas of dermal–epidermal interactions (ADEIs) of variable dimension and shape (Figs. 12 and 13). During the evolution of the two lines of amniotes (sauropsids and therapsids), glycine‐rich proteins acquired diVerent amino acidic composition in sauropsids versus therapsids. In specific derivative of the skin, under direct or indirect dermal influence, the activation of genes for glycine‐rich proteins progressively evolved in one direction in sauropsids, forming the so‐called b‐ or ‐keratins (Fig. 12I–M), and in another direction in therapsids, forming the HGTs (Fig. 12F–H, D–D4). In reptiles of the Mesozoic Era, a further elaboration of these genes for ‘‘b‐keratins’’ gave rise to high glycine–tyrosine–proline‐rich proteins (HGTPs) in chelonians, to high glycine–proline‐rich (HGPs) or high glycine–cysteine–proline‐rich proteins (HGCPs) in lepidosaurians, and to high glycine–cysteine–proline‐rich proteins (HGCPs) in archosaurians. From the latter, small high glycine–proline–serine–cysteine‐rich proteins (feather keratins) evolved in birds. Another line of evolution in therapsids gave origin to trichocytic keratins and to HSPs and UHSPs (Fig. 12H). In both lineages of amniote these small proteins, associated with specific filamentous keratins, built a scaled integument in reptiles, a feather integument in theropsids and birds, and a hairy integument in mammals. It has been hypothesized (Spearman, 1964, 1966) that parakeratosis or cornification without stratum granulosum was the common modality of cornification of early synapsids. In therapsids, parakeratosis was later replaced by orthokeratosis after the accumulation of HRPs to form a stratum

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granulosum. Therefore, the hard epidermis of synapsids was gradually replaced by a pliable and flexible epidermis. A thin and flexible epidermis allowed muscles to undergo plastic deformation that became sensitive, typical characteristics of mammalians (Alibardi and Maderson, 2003c; Findlay, 1970; Maderson, 1972a; Spearman, 1964). In some areas of the body where friction or mechanical resistance was needed, hard keratinization was obtained by the evolution of HGT or HSP matrix molecules (nails, horns, claws, and hoofs; Gillespie, 1991; Marshall et al., 1991; Powell and Rogers, 1994) (Fig. 12D1–D4). The molecular evolution of mammalian hard keratins and their matrix proteins in skin derivatives was the last step in the evolution of the process of cornification in mammals. The sauropsids evolved a mechanically strong, scaled integument. Before diapsids (archosaurians and lepidosaurians) and anapsids (chelonians) diverged, a new type of keratin or of keratin‐associated protein (‘‘b‐keratin’’) appeared. A harder corneous material oVered protection to the skin and dermal bones were reduced, so making lighter and more agile archosaurians (e.g., dinosaurs and birds) and lepidosaurians (lizards and snakes). The resistant glycine‐rich proteins able to replace cytokeratins in reptilian keratinocytes was later selected in therapods and proavians to produce a small keratin (feather keratin). The latter was used to form the fine branching of feathers (Chuong et al., 2000, 2003; Maderson and Alibardi, 2000; Maderson, 1972b; Prum, 1999; Prum and Brush, 2002; Wu et al., 2004). Another type of small b‐keratin is that used to make the climbing setae of geckos (Alibardi, 2003a; Alibardi and Toni, 2005b). Other types of HRP were also produced, especially in feathers, but their role is still enigmatic (Rogers et al., 1999; Sawyer et al., 2000; Sawyer and Knapp, 2003). 3. Evolution of Glycine‐Rich Proteins of the Matrix In cytokeratins of all vertebrates, a central and more conserved helical sequence is flanked at the two extremities by nonhelical and often hypervariable N and C heads (see Section III.A). These latter regions are responsible of most of the variation in a‐keratins, and in particular become richer in repeated sequences of glycine–glycine–serine or other, more basic amino acids. These end domains might have been important for the further evolution of more basic or larger keratins, or the isolation and subsequent evolution of genes encoding these glycine‐rich sequences originated the smaller keratin‐associated proteins. These regions do not possess an a‐helical conformation but a random coil, strand, or even b conformation in some regions. It has been suggested (Fuchs et al., 1987; Steinert and Freedberg, 1991) that the evolution of epidermal cytokeratins took place in three main stages: (1) origin of the central domain, (2) duplication and mutation to form type I and type II, and (3) origin of the end domains. In the first step, occurring

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more than 1 billion years ago, an ancestral gene (exon) made of a heptad motif with an a‐helical conformation duplicated repeatedly to form a new gene with at least four repeated regions interrupted by introns: this was an ancestral type I cytokeratin. The four repeats allowed the coiled‐coil conformation necessary to form the dimers that determine filament formation and the derived mechanical stability of the keratin filaments. Some mutations, perhaps causing a sliding in the intron–exon boundaries, produced the non‐a‐ helical interruptions (linkers) within the initial gene. By subsequent mutation, deletion, or loss of introns, the rod domain of type I and type II cytokeratins originated, well before vertebrates appeared. The following evolution of cytokeratins occurred by the incorporation of genes for the H and E regions, and later for the V1 and V2 regions. These regions are good candidates for the evolution of glycine‐rich proteins from cytokeratins. Cytokeratins in the epidermis of fish are relatively few, whereas they increase in isoforms in terrestrial vertebrates in relation to the strong keratinization required for the formation of the stratum corneum. Therefore cytokeratins have modified their original role as cytoskeletal elements (e.g., types K5/K14) to become an essential component and scaVold (K1/K10) for the deposition of other proteins destined to form the intracellular corneous mass and the peripheral cell corneous envelope. This might have occurred in more complex cytokeratins of suprabasal, precorneous, and corneous layers of amniote epidermis, by the increase in hydrophobic amino acid sequences rich in glycine and containing basic amino acids. This process therefore increased the basicity of these small, cytokeratin‐derived proteins. It is possible that the exons for V1 or V2 of an ancestral cytokeratin were duplicated and mutated independently of the remaining gene (Fig. 16, bottom). The hypervariable domain of cytokeratins possesses no introns (V1) or three introns (V2). A lack of introns seems to be the case for reptilian glycine‐ rich proteins isolated so far (L. Dalla Valle, personal communication). This small but independent (at least functionally) gene in basic amniotes could have further duplicated and mutated (Klinge et al., 1987) to produced various types of glycine‐rich proteins in both the derived therapsids and sauropsids (Fig. 16). This hypothesis is suggested from the analysis of the glycine–proline‐rich and glycine–cysteine–proline‐rich proteins of two lizards (Dalla Valle et al., 2005; Inglis et al., 1987) (Fig. 16). They both appear as chimeric proteins made of a mosaic of sequences present in epidermal proteins of birds and mammals. The central part of these proteins contains most of the prolines and have high homology with avian feather–scale–claw keratins. The lateral regions toward the N and C ends have high homology with mammalian KAPs of hairs. This suggests that reptilian basic amniotes, the ancestors of modern reptiles, birds, and mammals, possessed glycine‐rich proteins in their epidermis. The evolution of these glycine‐rich sequences perhaps derived from the same or similar genes encoding the glycine‐rich

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regions of the V domains of the N or C terminals of a‐keratins (Klinge et al., 1987; Steinert and Freedberg, 1991). These ancestral genes evolved diVerently in the lineage of therapsid amniotes (synapsids ! therapsids ! mammals) versus that of sauropsid amniotes (anapsids ! diapsids, i.e., reptiles and birds). In therapsids, from these glycine–serine‐rich sequences, a base change in the codon for serine originated the insertion of proline or of cysteine. The latter change allowed proline and cysteine that replaced the initial glycine– serine‐rich sequences and originated the E regions of trichocytic keratins (Klinge et al., 1987; Steinert and Freedberg, 1991) (see Fig. 16). The duplication, mutation, and independent evolution of the gene encoding these sequences (E regions) may have produced HSPs and UHSPs (Fig. 16). The selection of other amino acids within the glycine‐rich framework shifted the secondary structure and produced an a‐helical conformation. The secondary structure for these proteins, when studied under X‐ray, at least when they are aggregated with cytokeratins, produces an ‘‘a pattern.’’ In sauropsids (reptiles and birds), the initial glycine–serine‐rich sequence shifted for the insertion of proline. Their secondary structure, studied under X‐rays, produces a ‘‘b‐keratin’’ pattern. The selection of certain amino acids might have produced a typical b‐pleated conformation or a mixed random coil and strand conformation (Dalla Valle et al., 2005; Fraser and Parry, 1996). Small glycine‐rich sequences can form denser aggregates (Klinge et al., 1987). The latter can produce the high mechanical resistance of sauropsid scales, claws, beaks, and feathers. Although it remains to be further verified, it seems that more glycine‐rich proteins have been produced in mammals versus those of reptiles and birds (Gillespie, 1991). Ongoing studies have already shown at least 10 distinct genes involved in the production of lizard ‘‘b‐keratins,’’ and the number of genes for b‐keratins is growing and extending to other suborders of reptiles such as serpents and chelonians. Therefore we may expect more and more new genes and b‐keratins in reptiles, as indicated by biochemical studies (Marshall and Gillespie, 1982; Marshall et al., 1982; Wyld and Brush, 1979, 1983).

V. Conclusions and Future Directions Although the present knowledge of genomes and proteomics of nonmammalian vertebrates is limited, emerging concepts and generalizations are possible concerning the evolution of the epidermis in vertebrates. Superficial cells of the epidermis, in contact with the environment, became polarized and more specialized in land vertebrates to produce a corneous

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layer. The number of proteins and complex lipids progressively increased in the stratum corneum during land colonization, and later for further specialization (flight and homeothermy). During epidermal evolution, keratinocytes became initially heavily keratinized and a specific matrix formed intracellularly among keratin filaments. Underneath the plasma membrane numerous cell envelope proteins accumulated to form a resistant cornified cell envelope. The stratum corneum increased in thickness and eYciency for land survival. Cytokeratin complexity and number changed from simple to complex epithelia and produced nonkeratin‐associated proteins implicated in the formation of more resistant corneous material in the cytoplasm and/or deposited against the cell membrane of corneocytes. A diversification occurred in sauropsid amniotes (reptiles and birds) versus therapsid amniotes (mammals). This was made possible by new forms of dermal–epidermal interactions of diVerent shape and extensions that produced cutaneous derivatives, including scales, hairs, and feathers. It may now be possible to classify both sauropsid and therapsid keratins as soft or epidermal keratins (i.e., intermediate filaments, ubiquitous to all cells, where they constitute a large part of the cytoskeleton). Other specialized keratins have formed in the special derivatives of amniotes (hairs, nails, scales, feathers, etc.). Finally, keratin‐associated proteins (i.e., specialized cornification proteins, mammalian KAPs and reptilian/avian ‘‘b‐keratins’’) were formed to increase the mechanical resistance in these skin derivatives. Future studies in the area of molecular biology of the epidermis of nonmammalian vertebrates, especially reptiles, are needed in conjunction with localization of nucleotide sequences and proteins in epidermal cells, using immunological and in situ hybridization methods. A search for mammalian ortholog genes in the genomes of fish, amphibians, reptiles, and birds is needed, with subsequent genomic analysis and comparison, and chromosomal and protein localization. This knowledge will help increase our understanding of the process of evolution of specialized cytokeratins and of glycine‐rich keratin‐associated proteins, including loricrin and filaggrin, among others. Several questions remain to be addressed, especially on the genomic, molecular, and phylogenetic relationships between KAPs among amniotes, on HRPs or other proteins associated with keratinization, and on HSPs involved in the formation of the cell corneous envelope in nonmammalian vertebrates.

Acknowledgments Most of the financial support for this work was from grants (60% and traveling grants) from the University of Bologna. Personal funding has also been important, especially for animal collection, and the collection of samples of particular rarity (skin from lungfish, monotremes,

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marsupials, alligators, crocodiles, and tuatara). Thanks are due to many overseas colleagues for donation of various antibodies over the years (Drs. R. H. Sawyer, L. W. Knapp, G. Rogers, T. T. Sun, B. Dale, E. Fuchs, H. Baden, and L. Echart). Special thanks go to Dr. M. Toni (University of Bologna) for the intense collaborative eVort in the characterization of epidermal proteins and for some drawings. Mrs. L. Dipietrangelo and Mr. N. Mele (University of Bologna) helped in producing the drawings and figures. The collaboration with Dr. L. Dalla Valle and Dr. V. ToVolo (University of Padua) has allowed the study of reptilian b‐keratins and their genes.

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Further Reading Alibardi, L. (2003c). Immunocytochemistry and keratinization in epidermis of crocodilians. Zoolog. Stud. 42, 346–356. Aoki, N., Ito, K., and Ito, M. (1997). Isolation and characterization of mouse high‐glycine/ tyrosine proteins. J. Biol. Chem. 272, 30512–30518. Lavker, R. M., and Matoltsy, A. G. (1971). Substructure of keratohyalin granules of the epidermis as revealed by high resolution electron microscopy. J. Ultrastr. Res. 35, 575–581. Schnyder, U. W., and Roop, D. R. (1993). Expression patterns of loricrin in various species and tissues. DiVerentiation 54, 25–34.