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Scale keratin in lizard epidermis reveals amino acid regions homologous with avian and mammalian epidermal proteins.

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Scale Keratin in Lizard Epidermis
Reveals Amino Acid Regions
Homologous With Avian and
Mammalian Epidermal Proteins
Dipartimento di Biologia Evoluzionistica Sperimentale, University of Bologna,
Bologna, Italy
Dipartimento di Biologia, University of Padova, Padova, Italy
Small proteins termed ␤-keratins constitute the hard corneous material of reptilian scales. In order to study the cell site
of synthesis of ␤-keratin, an antiserum against a lizard ␤-keratin of 15–16 kDa has been produced. The antiserum recognizes
␤-cells of lizard epidermis and labels ␤-keratin filaments using immunocytochemistry and immunoblotting. In situ hybridization using a cDNA-probe for a lizard ␤-keratin mRNA labels ␤-cells of the regenerating and embryonic epidermis of lizard.
The mRNA is localized free in the cytoplasm or is associated with keratin filaments of ␤-cells. The immunolabeling and in
situ labeling suggest that synthesis and accumulation of ␤-keratin are closely associated. Nuclear localization of the cDNA
probe suggests that the primary transcript is similar to the cytoplasmic mRNA coding for the protein. The latter comprises
a glycine-proline-rich protein of 15.5 kDa that contains 163 amino acids, in which the central amino acid region is similar
to that of chick claw/feather while the head and tail regions resemble glycine-tyrosine-rich proteins of mammalian hairs.
This is also confirmed by phylogenetic analysis comparing reptilian glycine-rich proteins with cytokeratins, hair keratinassociated proteins, and claw/feather keratins. It is suggested that different small glycine-rich proteins evolved from
progenitor proteins present in basic (reptilian) amniotes. The evolution of these proteins originated glycine-rich proteins in
scales, claws, beak of reptiles and birds, and in feathers. Some evidence suggests that at least some proteins contained within
␤-keratin filaments are rich in glycine and resemble some keratin-associated proteins present in mammalian corneous
derivatives. It is suggested that glycine-rich proteins with the chemical composition, immunological characteristics, and
molecular weight of ␤-keratins may represent the reptilian counterpart of keratin-associated proteins present in hairs, nails,
hooves, and horns of mammals. These small proteins produce a hard type of corneous material due to their dense packing
among cytokeratin filaments. Anat Rec Part A, 288A:734 –752, 2006. © 2006 Wiley-Liss, Inc.
Key words: lizard; scales; ␤-keratins; mRNA; amino acid sequence; keratin-associated proteins;
Reptilian epidermis produces two major groups of keratins, indicated as ␣- and ␤ (␾)-keratins. These proteins
have different solubility, molecular weight, amino acid
composition, secondary conformation, degree of elasticity
properties, packing modality, X-ray diffraction pattern,
and biosynthesis (Baden and Maderson, 1970; Fraser et
al., 1972; Wyld and Brush, 1979, 1983; Maderson, 1985;
Landmann, 1986; Fraser and Parry, 1996; Sawyer et al.,
2000; Alibardi and Sawyer, 2002; Alibardi et al., 2004a).
␣-keratins represent a scaffold for the formation of the
soft and pliable stratum corneum, termed ␣-layer. The
latter allows scale mobility and houses the barrier against
water loss. Instead, ␤-keratins form a hard and inflexible
layer that protects scales from mechanical stress.
As opposed to ␣-keratins, ␤-keratins of birds and reptiles are not extensible and are small proteins with a
␤-pleated secondary conformation. These proteins aggre©
gate into a densely packed lattice that produces very resistant microfibrils with a typical X-ray and ultrastructural pattern of 3– 4 nm electron pale filaments in a denser
matrix. The mechanism of aggregation of ␤-keratins to
form the very resistant cables is poorly known (Gregg and
Grant sponsor: 60% Grants from the Universities of Bologna
and Padova.
*Correspondence to: Lorenzo Alibardi, Dipartimento di Biologia Evoluzionistica Sperimentale, via Selmi 3, University of Bologna, 40126, Bologna, Italy. Fax: 39-051-2094286.
Received 30 November 2005; Accepted 30 March 2006
DOI 10.1002/ar.a.20342
Published online 7 June 2006 in Wiley InterScience
TABLE 1. List of compared sequences with their NCBI accession number
hypothetical protein [Dictyostelium]
Accession number
(Inglis et al., 1987)
(Dalla Valle et al., 2005)
KAP 5-1[Homo]
K-g3 [Lampetra]
K-gamma2 [Lampetra]
Lfl-K 1 [Lampetra]
Rogers, 1986; Fraser and Parry, 1996). In reptilian epidermis, the ␤-layer mainly determines mechanical resistance but is also the depositary of most epidermal pigmentation: both resilience and pattern ornamentation are the
first positive characteristics exploited in the leather industry using reptilian skin. The above studies have shown
that ␤-keratins in reptiles, as those of birds, have a
smaller molecular mass (10 –22 kDa) than ␣-keratins
(40 – 68 kDa), but very little is known about their amino
acid sequences and spatial conformation.
␤-keratin is believed to be phylogenetically more recent
than ␣-keratin but no information is presently available
on the evolution of ␤-keratin from the likely more primitive ␣-keratin (Sawyer and Knapp, 2003). Previous studies have shown that in lizard epidermis, acidic and basic
cytokeratins similar to those of mammalian epidermis are
present (Fuchs and Marchuk, 1983; Alibardi et al., 2000,
2001; Alibardi and Toni, 2006a).
␣- (soft) keratins are composed of 8 –10 nm thick keratin
filaments (intermediate filaments) and in epidermal cells
are referred to as cytokeratins. The filaments require the
association of one acidic and one basic keratin chain to
form a dimeric coil-coiled molecule that has a central
␣-helical rod and two variable N- and C-heads (Fuchs and
Marchuk, 1983; Klinge et al., 1987; O’Guin et al., 1987;
Steinert and Freedberg, 1991; Fuchs and Weber, 1994;
Coulombe and Omary, 2002).
In the mammalian epidermis, the keratin pair is K5/
K14 in basal and suprabasal layers and K1/K10 pair in
upper spinosus and cornifying layers. ␤-keratin filaments
are thinner (3– 4 nm) than cytokeratin filaments.
The epidermis of lepidosaurian reptiles (snakes and lizards) is produced from cyclically alternating epidermal
Accession number
generations (shedding cycle) (Maderson, 1985; Landmann,
1986; Maderson et al., 1998). ␤-keratin is produced in
specific layers, termed oberhautchen and ␤-layer, but is
replaced by ␣-keratin in the following layers (mesos, ␣-,
lacunar, and clear layer). These keratins are synthesized
during the renewal phase of the shedding cycle of lizard
epidermis, which terminates with the molt of the external
(outer) epidermal generation. After shedding, the epidermis enters in a resting phase in which proliferation is
halted almost completely: this phase generally occupies
most of the time of the shedding cycle, until the next
renewal phase.
The renewal phase of the shedding cycle in the epidermis of lizards can be induced after tail amputation, a
process that stimulates the regeneration of the tail. The
regenerating tail epidermis forms a proximodistal sequence of scale neogenesis that reproduces the renewal
phase of both embryonic and normal epidermis (Maderson
et al., 1998; Alibardi, 2000, 2001, 2003). Thus, the differentiation of a new ␤-layer can be induced for studying the
expression of ␤-keratins. The differentiation of the first
␤-layer can also be studied in embryonic scales in late
embryonic stages of lizards (stages 38 –39) (Alibardi,
In a previous study (Dalla Valle et al., 2005), we have
selected and sequenced by RT-PCR/rapid amplification of
cDNA ends (RACE) analysis a lizard scale keratin cDNA
and deduced the amino acid sequence (Genebank accession number AJ890445).
In the present study, we have utilized the ␤-keratin
cDNA probe, generated in the above study, to analyze
further the expression of the mRNAs by light microscopy
and ultrastructural in situ hybridization in both regener-
ating and embryonic epidermis. This was done in conjunction with an immunolocalization study with an antiserum
directed against a 15–16 kDa protein (␤-keratin) isolated
from lizard epidermis.
Moreover, a comparative analysis of the amino acid
sequence of the lizard protein with mammalian and avian
keratins or with keratin-associated proteins (KAPs) provided some clues on the molecular evolution of hard proteins in amniote skin appendages such as scales, hairs,
and feathers.
Experimental Procedures
Tail amputation was made as previously indicated (Alibardi, 2001) by inducing tail autotomy, a natural process
of tail loss obtained by pulling and twisting the two-third
proximal of the tail in adult males and females of the
lizard Podarcis sicula and P. muralis. Tail regeneration
took place at a temperature ranging from 24 to 28°C. Six
specimens of P. sicula and five specimens of P. muralis
with regenerating tails of 3–5 weeks postamputation were
used. The specimens of P. muralis were studied only for
the histological study on the process of tail regeneration.
After this period, new scales were forming in proximal
regions near the tail stump of the 0.8 –2.0 cm long regenerating tail. The regenerating tail was collected by induced autotomy in the proximal tail stump and skin was
sectioned off and immediately fixed. Eight specimens of P.
sicula were used for the microscopic study.
Nine lizard embryos of P. sicula at embryonic stage 37
(n ⫽ 3), 38 (n ⫽ 3), and 39 (n ⫽ 3) were collected and fixed
for the histological study. In the last two stages, a ␤-layer
is differentiating in embryonic scales (Alibardi, 1996). Sections from embryos at stage 39 were selected for the in situ
hybridization study for ␤-keratin messenger expression
and for immunocytochemistry.
␤-Keratin Isolation, Antibody Production, and
Normal and regenerating skin from the lizard Podarcis
sicula were collected, immediately frozen in liquid nitrogen, and stored at ⫺80°C. The skin was incubated in 5 mM
EDTA in phosphate-buffered saline (PBS) for 3–5 min at
50°C and 2– 4 min in cold PBS. The epidermis was separated from the dermis by dissection under the stereomicroscope and the dermis discarded. Then, the epidermis
was homogenized in lysis buffer (8 M urea, 50 mM TrisHCl, pH 7.6, 0.1 M 2-mercaptoethanol, 1 mM dithiothreithol, 1 mM phenylmethylsulphonyl fluoride) to extract
keratins according to the method of Sybert et al. (1985).
The protein concentration was determined by the Lowry
For antiserum production, lizard epidermis proteins
were extracted (as described above) and separated by
SDS-PAGE. Following electrophoresis, the gel was fixed
and stained with Coomassie blue and two close protein
bands with a molecular weight of 15–16 kDa, corresponding to a ␤-keratin (as confirmed by immunocytochemistry), were detected. The bands were sliced off with a razor
blade and directly emulsified in 2 ml of PBS-CFA (complete Freund’s adjuvant; Sigma) solution (1:1 vol/vol). The
adjuvant/antigen solution containing approximately 150
␮g of the selected proteins was injected subcutaneously in
a rat that was previously bled to collect blood sample
Fig. 1. Gel electrophoretic separation of epidermal proteins in P.
sicula with the chosen bands (square) used for immunization (lane A).
The preimmune sera (lanes B and D) show no labeled bands in the 14 –16
kDa range, which instead appear in their respective immune sera (A612
serum, lane C; A68B serum, lane E).
(preserum). One booster immunization, prepared as
above, was performed 2.5 week after the first injection.
The rat was sacrificed after about 3 weeks after the boost
injection and blood was collected. Serum was then separated by centrifugation from clot and debris and stored at
⫺20°C. Following the above protocol, two different antisera (A612 and A68B) against lizard ␤-keratin were produced.
For Western blot analysis, epidermis proteins separated
by SDS-PAGE (50 ␮g per lane) were transferred to nitrocellulose paper (Hybond C Extra; Amersham, U.K.) and
incubated with rat antisera (A612 or A68B; dilution 1:25–
100) as primary antibodies. The filters were then washed
and further incubated for 1 hr at room temperature with
antirat HRP-conjugated secondary antibody (dilution
1:1,000) in TBS-Tween ⫹ 5% nonfat milk powder. Detection was performed by using the enhanced chemiluminescence’s procedure developed by Amersham (ECL).
Sequence Analysis
The amino acid sequences of lizard scale (Dalla Valle et
al., 2005) and lizard claw (Inglis et al., 1987) were compared with some avian and mammalian keratin-like proteins. These proteins represented a keratin-associated
protein, an ␣-keratin, a scale ␤-keratin, and a feather
␤-keratin. Furthermore, a broader phylogenetic analysis
was also performed to correlate the amino acid sequences
of ␤-keratins of scales, feathers, and claws, the sequences
of cytokeratins of different vertebrates, and the sequences
of keratin-associated proteins deposited in the NCBI protein database (protein accession numbers are reported in
Table 1). These sequences were aligned using ClustalX
(1.81) software (Thompson et al., 1997). The phylogenetic
tree was constructed by the neighbor-joining method (Saitou and Nei, 1987) and was visualized using the NJPLOT
software. ClustalX (1.81) and NJPLOT software are available for free download (
Two to 5 mm long skin pieces collected from three regenerating tails and from three embryos at stage 39 were
Fig. 2. Light microscopic view of normal (A), regenerating (B–E), and
embryonic (F–H) epidermis of lizard. A: Beneath the ␣-layer, precorneous cells (arrows) are differentiating. Scale bar ⫽ 10 ␮m. B: Epidermal
peg penetrating the dermis in regenerating tail at 3 weeks postamputation. Beneath the lacunar epidermis, a granulated (clear) layer is forming
(arrows), followed by early differentiating ␤-cells above a stratified epidermis. Scale bar ⫽ 10 ␮m. C: Cross-sections of proximal regenerating
scale at 3 weeks postamputation showing the formation of large granules in the clear layer (arrows) with the thick differentiating ␤-layer. Scale
bar ⫽ 10 ␮m. D: Detail of epidermal differentiation in elongating pegs at
4 weeks of tail regeneration. Beneath the granulated layer (arrows, the
future shedding line), 3– 4 layers of fusiform ␤-cells are forming the
␤-layer. Scale bar ⫽ 10 ␮m. E: Regenerating scales of 4 weeks postamputation. The figure shows the passage from a cellularized ␤-layer
near the tip of the regenerating scale to the dark compact ␤-layer. Cells
of the granulated layer (arrow) also become dense and form the compact
clear layer (arrowhead). The latter produces denticles that interdigitate
with the specular oberhautchen serration. Scale bar ⫽ 20 ␮m. F: Symmetric ventral scale at stage 37. Scale bar ⫽ 10 ␮m. G: Tail scale at
stage 39 with a 4 –5 dark layers of flat ␤-cells in the outer surface (arrow)
beneath the embryonic epidermis (arrowhead). The dark layer remains
monocellular in the inner surface and hinge region. Scale bar ⫽ 20 ␮m.
H: Part of mature dorsal scale at stage 39 showing the packed ␤-layer
(arrow) and dense detaching embryonic epidermis (arrowhead). Scale
bar ⫽ 20 ␮m. ␤-, ␤-layer; ␤C, compacted ␤-layer; D, dermis; E, living
epidermis; H, hinge region; I, inner scale surface; L, lacunar layer; W,
wound epidermis.
fixed for 4 –5 hr in 4% paraformaldehyde in 0.1 M phosphate buffer at pH 7.4. Tissues were dehydrated and embedded in the hydrophilic resin bioacryl (Scala et al.,
1992). Thick (2– 4 ␮m) and thin (40 –70 nm) sections were
collected respectively for light and ultrastructural cytochemistry. For histological study, sections were stained in
0.5% toluidine blue or by hematoxylin and eosin stain.
For immunocytochemistry, the above-described rat immune serum against a lizard ␤-keratin of 15–16 kDa was
utilized on sections of lizard scales and embryos.
Nonspecific binding components were blocked with 5%
normal goat serum and 1% BSA in PBS containing 0.3%
Triton X-100 for 30 min at room temperature. The primary antiserum (1:50 –100 in 0.5 M Tris buffer at pH 7.6
containing 2% BSA) was overlaid on each section for
14 –16 hr at 4°C. Sections were subsequently incubated
with a goat antirat FITC-conjugated secondary antibody
diluted 1:50 (Sigma) for 30 min at room temperature.
Immunoreactivity was detected under a UV epifluorescence microscope. The specificity of the immunolabeling
was tested by omitting the primary antibody.
Thin sections of skin (40 –90 nm thick) were collected on
nickel grids using an ultramicrotome. Sections were incubated overnight with the primary rat antibody as for light
microscopy immunocytochemistry, rinsed in buffer, and
incubated for 1 hr at room temperature with goat antirat
IgG conjugated with 10 nm gold particles (Sigma; diluted
1: 50). Detection was done using a secondary antirat antibody conjugated with 10 nm gold particles (Sigma; diluted 1: 50). After rinsing, sections were stained for 7 min
with 2% aqueous uranyl acetate, and grids were observed
under the electron microscope.
In Situ Hybridization
Tails were collected from six lizards with regenerating
tail showing scale neogenesis (3–5 weeks) and fixed in 4%
paraformaldehyde as above for immunocytochemistry.
The other three embryos (stage 39) were fixed in 4% paraformaldehyde. Tissues were dehydrated in ethanol at increasing concentration, infiltrated in xylene, and embedded in wax.
Sections of 6 – 8 ␮m thicknesses were collected using a
rotary microtome, dewaxed with xylene, and hybridized
using incomplete digoxigenin-labeled antisense probes
(corresponding to the last part of the coding region and the
3⬘-UTR sequence of lizard keratin, nucleotides ⫹382/
⫹835), or complete digoxigenin-labeled antisense cDNA
probes (corresponding to the complete coding region). The
plasmids containing these fragments [prepared in previous work; see details in Dalla Valle et al. (2005)] were
amplified by PCR with the M13 forward and reverse primers. The two fragments obtained were then used to produce, by asymmetric PCR with digoxigenin-labeled nucleotides (DIG, Roche), digoxigenin-labeled sense and
digoxigenin-labeled antisense incomplete and complete
cDNA probes.
For the hybridization, the probe concentration was 0.5–
1.5 ng/␮l of digoxigenin-labeled antisense probe in hybridization solution (50% formammide, 4 ⫻ SSC, 0.1% Tween20, 50 ␮g/ml tRNA, 10 mM EDTA, 50 ␮m/ml heparin,
0.5% blocking reagent; Roche, Applied Science, Milan, Italy). Hybridization was carried out overnight at 40 – 42°C.
Posthybridization treatment included a washing procedure with 2 ⫻ SSC (3 ⫻ 10 min, at 50°C), 0.5 ⫻ SSC (1 ⫻
15 min, at 60°C), 0.2 ⫻ SSC (1 ⫻ 15 min, at 60°C), and
0.1 ⫻ SSC (1 ⫻ 15 min, at 60°C). Afterward, the incubation with the anti-DIG-fluorescein Fab fragment antibodies (Roche) diluted 1:15 in Tris buffer was performed for 2
hr. The labeling was detected under fluorescence microscopy. Other sections were incubated with anti-DIG alkaline phosphatase-conjugated antibody (Roche) diluted
1:500 in PBS and color reactions were started with NBT/
BCIP substrate.
For ultrastructural in situ hybridization, 40 – 80 nm
thick sections of regenerating scales containing differentiating ␤-cells were collected on nickel grids and incubated
with the same hybridization solution as described above
for 4 –5 hr. Rinsing steps were the same as described
above. Afterward, the sections were incubated at room
temperature for 20 min in Tris 0.05 M, pH 7.6, with 2%
bovine serum albumin and for 3 hr with the mouse antiDIG antibody (Roche) diluted at 1:100. After repeated
rinsing in the buffer, grids were reacted for 1 hr at room
temperature with the secondary antimouse antibody (conjugated with gold particles of 5 nm in diameter). After
rinsing, grids were stained for 5–7 min in 2% uranyl
acetate, dried, and observed under a CM-100 Philips electron microscope. In situ hybridization controls were performed on sections consecutive to positive reactions and
included omission of the probes and hybridization with a
labeled sense probe.
The injection of epidermal protein bands at 15–16 kDa
of P. sicula epidermis (Fig. 1, lane A) into rats produced
immune sera (Fig. 1, lanes B and C for serum A612, and D
and E for serum A68B). Immune sera generally reacted
with lizard epidermal proteins, producing a specific immunolabeled band at 15–16 kDa, sometimes with a weak
band at 30 –32 kDa. These bands were absent in presera.
In negative controls, omitting the primary antiserum, no
immunolabeled bands appeared.
The distribution of both the antiserum and the probe for
the ␤-keratin mRNA was studied in lizard epidermis. No
difference in the histology of regenerating tail was observed between the two species used (P. muralis and P.
sicula). Normal epidermis consisted of a corneous layer
where an outer ␤-layer was followed by the ␣-layer and
the remaining living epidermal layers (Fig. 2A). After 2–3
weeks postamputation of the tail in both species, a wound
epidermis made of spinosus-like keratinocytes covered the
regenerating tail (data not shown). In proximal regions,
near the skin of the tail stump, the wound epidermis
formed pegs penetrating the dermis (Fig. 2B). The pegs
progressively elongated in distal-proximal direction, and
in the following 1–-2 weeks their axial part started to
differentiate fusiform cells of the ␤-layer (Fig. 2C and D).
The following maturation and compaction of ␤-cells allowed the new formation of a ␤-layer (Fig. 2E). The detachment from the upper cornified wound epidermis occurred by the differentiation of a shedding complex,
comprising a granulated layer (termed clear layer, with
down-pointing serration) and an oberhautchen layer (with
up-pointing serration). Therefore, beneath the cornified
wound epidermis, the sequence of layers was clear layer
with its granules, serrated oberhautchen, and ␤-layer.
Fig. 3. Immunofluorescence of lizard scales after application of the
lizard ␤-antiserum (A68B) in normal scale (A), regenerating scales (B and
C), and negative control (D). Immunostaining with the lizard keratin
antiserum on embryonic scales (E and F). In situ hybridization immunofluorescence with antisense DNA incomplete probe (aDi) on regenerating
scales (G and H), sense DNA incomplete (sDi) probe (I), and negative
control (co) probe (J). H, hinge region; T, scale tip. Dashes underline the
basal layer; arrows indicate the more or less compact ␤-layer. Scale
bars ⫽ 10 ␮m.
The ␣-keratogenic cells differentiated underneath the
␤-layer to reconstitute later a new ␣-keratin layer (data
not shown).
In the embryo, at stages 37 in some areas of the body
(e.g., midtrunk lateral and limbs), the formation of bumps
represented the symmetric stage of scale development
(Fig. 2F). Bumps in other regions of the body (e.g., lumbar
and tail) at stage 37 and in the remaining body areas at
stage 38 became asymmetric and rapidly formed the outer
scale surface (Fig. 2G). At this stage, the epidermis was
made of 3– 4 layers of cells above the germinal layer.
During elongation of the outer scale surface, fusiform
␤-cells started to accumulate beneath the embryonic layers and the epidermis became thicker (Fig. 2G). In the
inner side and hinge region of the developing scales, no
fusiform ␤-cells were present and the epidermis remained
thin. The three more external, very flat layers formed the
outer and inner periderm and the third layer represented
the embryonic oberhautchen (data not shown). Beneath
the latter, fusiform and dark cells started to form the
embryonic ␤-layer that was very reduced in the inner scale
surface and hinge regions (Fig. 2G). In the outer scale
surface, the ␤-layer was formed by 3–5 layers of fusiform
␤-cells and eventually formed a compact ␤-layer at stage
39 (Fig. 2H). In some scales (e.g., in the proximal tail and
ventral scale), both the embryonic epidermis and the
␤-layer packed into two dense layers, and the embryonic
epidermis started to detach from the ␤-layer (Fig. 2H).
However, scales in different parts of the body were at
slightly different stages, and the ␤-layer was still immature (unpacked).
Light Microscopy Immunocytochemistry and In
Situ Hybridization
The ␤-layer of normal, regenerating, and embryonic
scales was immunofluorescent using our rat antiserum
A68B against the 15–16 kDa lizard ␤-keratin (Fig. 3A–C).
In normal epidermis, the ␤-layer corresponded to the external compact corneous layer (Fig. 3A). In the regenerating epidermis, only the forming ␤-layer (Fig. 3B) and the
mature compact ␤-layer of regenerated (neogenic) scales
were immunofluorescent (Fig. 3C). Negative controls were
not immunolabeled (Fig. 3D).
In the embryonic epidermis at stage 39, the ␤-layer was
also immunostained and the labeling tended to disappear
in the hinge region (Fig. 3E and F). Other layers of the
epidermis or other tissues of the skin were immunonegative. Negative controls were not immunolabeled (data not
The in situ hybridization study by immunofluorescence
was done using the antisense DNA incomplete probe (aDi;
digoxigenin-labeled corresponding to the last part of the
coding region and the 3⬘-UTR sequence of lizard keratin;
nucleotides ⫹382/⫹835). The antisense DNA complete
probe (aDc; digoxigenin-labeled whole 485 nucleotidic coding region probe; see sequence in Fig. 9A) was also used.
The probes were added to sections of regenerating tail (4
weeks postamputation) at a concentration of 0.5–1.5 ng/␮l
(see details in Dalla Valle et al., 2005), and the formed
hybrid produced a specific immunofluoresecence in differentiating fusiform ␤-cells present in the regenerating
outer scale surface (Fig. 3G and H). Both the sense DNA
incomplete probes (sDi) and the negative control showed a
very weak or absent labeling (Fig. 3I and J).
The in situ hybridization using the detection system
based on alkaline phosphatase produced a reddish-purple
positive reaction exclusively in the forming ␤-layer of regenerating scales (4 weeks postamputation) or embryonic
scales (stage 39; Fig. 4A–J). In apical regions of the regenerating tail, where the epidermis was still undifferentiated (stratified wound epidermal cells), no detection of
labeled cells was seen in epidermal pegs (Fig. 4A and B).
In more proximal regions where scales were differentiating new epidermal layers underneath the wound epidermis, the first fusiform ␤-cells appeared labeled with the
probes. Differentiating fusiform cells of the ␤-layer appeared distinctively reactive for the probes and produced a
red-purple stain. The stratification of the forming ␤-layer
increased to 3–5 cell layers in fully differentiating scales
(Fig. 4C–E). No other skin tissues were stained. Negative
and sense controls were not stained, and only a brown
pigmentation was sometimes seen (Fig. 4F). In embryonic
scales, 2–3 layers of ␤-cells appeared specifically stained
with the probe, while negative and sense controls showed
no labeling or very low, mainly due to the pigmentation
(Fig. 4K and L).
Ultrastructural Immunocytochemistry and In
Situ Hybridization
The ultrastructural examination, using immunogold
of 10 nm, showed that the lizard ␤-antibody (A68B,
directed against a 15–16 kDa lizard epidermal protein)
decorated only electron-pale bundles of keratin among
ribosomes of ␤-cells (Fig. 5A). The pale keratin bundles
of the oberhautchen layer were less labeled or unlabeled
with this antibody. High-magnification observation on
early differentiating ␤-cells showed that small ␤-packets were labeled with the antibody (Fig. 5B). The small
keratin packets formed a typical reticulate pattern or a
network of matrix material of low electrondensity,
mainly around more densely packed filaments of ␤-keratin material. The latter appeared at a following stage
of compaction of the small ␤-packets forming the reticulate pattern.
Dense granules in oberhautchen and ␤-keratin cells
were less labeled or completely unlabeled. Keratohyalin
granules of the clear (granulated) layers and ␣-keratin bundles were unlabeled. The large ␤-keratin filaments were
merging into a denser mass of corneous material in more
mature compacting ␤-cells. These tangled filaments were
intensely labeled with the lizard ␤-keratin antibody. The
labeling disappeared in the differentiating mesos and ␣-cells
that were formed underneath the ␤-layer (data not shown).
Ultrastructural in situ analysis over regenerating
scales using the antisense complete DNA probe (aDc)
showed clusters of gold particles sparse in the cytoplasm
of differentiating ␤-keratin cells (Fig. 6A). The clusters
were localized among ribosomes or associated with small
␤-packets: the latter formed the pale ␤-keratin reticulate
pattern previously described (in Fig. 6B) among the larger
␤-keratin filaments (Fig. 6). This finer material was associated with the periphery of larger and compact ␤-keratin
filaments, while their central part was generally devoid of
clusters of gold particles (Fig. 6B). Gold particles often
formed clusters among the reticulate network of material
of low electrondensity: the latter represented early stages
of ␤-keratin accumulation (Fig. 6C).
Although less frequent than in the cytoplasm, some gold
clusters were also seen inside the nucleus of ␤-cells, espe-
Fig. 4. In situ hybridization with alkaline phosphatase detection of
antisense DNA complete probe of early regenerating stages (epidermal
pegs; A and B), incomplete probe (aDi; C) on elongated pegs, complete
probe (aDc) of later stages of elongated pegs (D and E), sense DNA
incomplete probe (sDi; F) in regenerating scales. aDc in dorsal scales of
embryo at stage 39 (G) and its control (co; H). Detail of labeled (aDc)
fusiform ␤-cells in dorsal (I) and tail (J) scales of embryo at stage 39.
Negative control (K) and sense control (L) of tail scales of embryo at
stage 39. E, living epidermis; H, hinge region; T, scale tips; W, wound
epidermis. Arrows indicate the differentiating the ␤-layer; arrowheads
indicate pigments; dashes underline the basal layer; asterisks indicate
non specific reaction. Scale bars ⫽ 10 ␮m.
Fig. 5. Ultrastructural immunolabeling of ␤-keratin filaments with the
lizard keratin antiserum. A: Region of passage from the granulated (clear)
layer to the underlying oberhautchen and ␤-layer. While no labeling is
present over keratohyalin-like granules and keratin bundles (arrowheads)
of clear cells, gold particles cover some oberhautchen ␤-keratin packets
and intensely the pale keratin filaments of ␤-cells (arrow). Scale bar ⫽
500 nm. B: Detail of labeling (arrowheads) of ␤-keratin packets in early
differentiating ␤-cell. Scale bar ⫽ 100 nm. G, dense granule; KH, keratohyalin-like granules; O, oberhautchen serration.
cially among the finer euchromatine (Fig. 7). Clusters
were sometimes seen associated with the nuclear membrane or localized in the cytoplasm near the nucleus: in this
way, the sparse probe labeling extended from the internal
part of the nucleus to the cytoplasm. The immunogold labeling was completely absent or randomly diffused and did not
form clusters in other cells of the epidermis and dermis (data
not shown) or in sense controls (Fig. 8).
with a keratin-associated protein, an ␣-keratin, a scale
␤-keratin, and a feather ␤-keratin is presented (Figs. 9 –12).
They showed numerous region homologies as it is reported in
the Discussion section. A phylogram of different cytokeratins, ␤-keratins, and KAPs is reported in Figure 13.
In Figure 9, we have compared the lizard scale protein
with a keratin-associated protein (16-1) of mouse hair and
with a cytokeratin (␣-2) of lamprey epidermis. The homology between the lizard glycine-rich protein and the keratin-associated protein is high, while glycine-rich regions of
the lamprey cytokeratin toward the N- or the C-terminal
present relatively high homology with the lizard scale pro-
Comparative Analysis of Protein Sequences
The comparison between amino acid sequences of lizard
scale (Dalla Valle et al., 2005) and claw (Inglis et al., 1987)
Fig. 6. Ultrastructural in situ hybridization in the cytoplasm of differentiating ␤-cells. A: Gold clusters
(arrowheads) in the cytoplasm among ␤-packets. B: Detail of clusters of gold particles associated with the
periphery of larger ␤-keratin filaments (arrowhead). C: Details of cluster of gold particles among alveolate
␤-packets (arrowhead) near a compact mass of ␤-keratin filament (arrow). Scale bars ⫽ 100 nm.
tein. In Figure 10, we have compared the lizard scale protein
with a scale keratin of chick and with a feather keratin of
chick. The homology is also high in most region of the scale
keratin and in some regions of feather keratin. In Figure 11,
we have compared the lizard claw protein with a human
keratin-associated protein (5-1) and with a human keratin of
type II. There is some homology in various glycine-rich regions between the lizard claw and a human KAP and less
homology with the cytokeratins. In Figure 12, we have compared the lizard claw protein with scale and feather keratin
of the chick, where the homology is high in various regions.
Finally, a phylogenetic analysis using the ClustalX
(1.81) software has shown that lizard scale and claw glycine-rich proteins are more related to mammalian KAPs
than avian ␤-keratins and less related to several vertebrate cytokeratins (Fig. 13). As a result of this analysis on
Fig. 7. Ultrastructural in situ hybridization in nuclei of differentiating ␤-cells. A: Clusters of gold particles
(arrow) within the nucleus (N). Arrowheads indicate unlabeled ␤-keratin filaments in the cytoplasm. B: Details
of cluster of gold particles among reticulate euchromatin material. C: Details of a cluster of gold particle
(arrow) near the nuclear membrane. N, nucleous; Cy, cytoplasm. Scale bars ⫽ 100 nm.
Fig. 8. Ultrastructural view of a sense control section of a differentiating ␤-cell. Both the cytoplasm and
␤-keratin filaments (double arrowheads) show completely absent or very sparse gold particles. Scale bar ⫽
100 nm.
the only two known primary sequences of reptilian keratins (lizard scale and claw), it appears that lizard keratins
are a mosaic of sequences present in KAPs and in ␤-keratins of mammals and birds.
The conformational prevision, using the program “consensus secondary structure prediction,” for the lizard scale
protein shows that no ␤-pleated sheets are present. In
particular, 104 amino acids (64.2%) have a random coil
Fig. 9. Sequence comparison of the lizard scale protein (B), deduced
from its complete cDNA (A) (Dalla Valle et al., 2005), with KAP 16-1 of
mouse (C), and ␣-2 keratin of lamprey (D). Stars indicate identities; colon
indicates conserved substitutions; period indicates semiconserved substitutions detected by the ClustalW software. Glycine residues are evi-
denced in gray shadows. Underlines indicate the glycine-rich regions
(mammalian-like) in N- and C-terminal ends of the lizard protein, while
the boxed sequence indicates the central feather-claw-like sequence.
Accession numbers of sequences analyzed are reported in Table 1.
conformation, 50 amino acids (30.7%) an extended strand
conformation, and only 9 amino acids (5.5%) an ␣-helical
conformation. The specific X-ray pattern and tertiary
structure of this protein remain to be analyzed. It is possible that the strand conformation or interchain interactions are responsible for the ␤ X-ray pattern ascribed to
these proteins (Fraser et al., 1972; Fraser and Parry,
Three amino acid regions of the lizard glycine-prolinerich protein are interesting, one in the N-region, one in the
C-region, and one in the central region. They are here
compared using the BLAST or ClustalX (1.81) software
with sequences present in cited publication or deposited in
Genbank database. In the present study, only some examples of sequence homology between reptilian and avian
proteins are indicated (Figs. 9 –12). The homology is indicated by the percentage of identities (⫽ same amino acids)
and positivities (⫽ same amino acids or amino acids that
preserve the physicochemical properties).
From amino acid residues 19 – 60 near the N-end, the
lizard protein contains glycine-X or glycine-glycine-X sequences with high homology with mouse glycine-tyrosine-
Fig. 10. Sequence comparison of the amino acids of the lizard scale
protein (A), with chicken scale keratin (B) and chicken feather keratin (C).
Stars indicate identities; colon indicates conserved substitutions; period
indicates semiconserved substitutions detected by ClustalW software.
Glycine residues are evidenced in gray shadows. Underlines indicate the
glycine-rich regions (mammalian-like) in N- and C-terminal ends of the
lizard protein, while the boxed sequence indicates the central featherclaw-like, sequence Accession numbers of sequences analyzed are
reported in Table 1.
rich type I proteins (Genbank accession number
AAKO7673.1; 62–78% identities and 65– 81% positivities).
Other sequence homology is present comparing lizard protein with goat keratin-associated protein (Genbank accession number AAR89458.1; 70% identity, 77% positivity),
with human keratin-associated protein (Genbank accession number NP853640.1; 65% identity, 73% positivity), or
with glycine-tyrosine rich proteins (Genbank accession
number AAP97270.1, 56% identities, 61% positivities).
Near the C-terminal region, the lizard protein also presents homologies with mammalian glycine-tyrosine-rich
hair-associated proteins. For example, it resembles a keratin-associated protein of mice hairs (Genbank accession
number NP570927.1; 61% identities, 64% positivities), a
glycine-tyrosine-rich mouse protein (Genbank accession
number AAKO7673.1; 59% identities, 70% positivities),
and also a human keratin-associated protein of hairs
(Genbank accession number NP853638.1; 62% identities,
66% positivities).
From the above results, it can be seen that the N- and
C-termini of the scale lizard glycine-proline-rich protein
have mammalian-like homologies that suggest a common
ancestry with those of mammals (Gillespie et al., 1982;
Marshall and Gillespie, 1982).
Finally, the central region between amino acids 77 and
118 resembles (40% identities, 61% positivities) various
sequences of avian feather, scale, and claw keratins [see
sequences in Table 2 of Sawyer et al. (2003)]. In particular, the analysis of amino acids 77–111 of the lizard protein shows homology with chick claw keratin (Genbank
accession number XP 428214.1; 75% identities, 90% positivities), chick feather keratin I (Genbank accession number XP424547.1; 73% identities, 94% positivities), and
another feather keratin (Genbank accession number
426556.1; 70% identities, 88% positivities).
The analysis of amino acids in position 26 – 60 show
homology with chicken claw keratin (Genbank accession
number P25692; 54% identities, 71% positivities), and
analysis of amino acids 23– 64 show homology with chick
feather keratin (Genbank accession number P04458; 45%
identities, 64% positivities). Therefore, a large part of the
central region of the lizard glycine-proline-rich protein
shows homologies with claw, scale, and feather proteins,
which also may indicate some common ancestry with
those of archosaurian progenitors. The possibility that
these sequences derive from convergence of molecular evolution remains another possibility that requires further
analysis of much more reptilian primary structure.
The lizard claw glycine-cystein-proline-rich protein also
appears as a chimeric protein and presents homologous
regions with both mammalian (high-sulfur- and high-tyrosine-rich proteins) and avian ␤-keratins (Gillespie et al.,
1982; Inglis et al., 1987). Amino acid residues 1–36 have
homology with human cytokeratins. Also, residues 1–36,
63–98, and 132–142 have homologies with high-cystein
wool keratins. Finally, residue 32– 67 shows homologies
with feather keratin. Conformational analysis of the 142
amino acid lizard claw ␤-keratin sequence showed that
the amino acid residues 36 – 67 and 94 –125 determine the
␤-sheet component, as amino acids 24 –55 in emu ␤-keratin of feathers also determines most of the ␤-sheet conformation (Fraser and Parry, 1996). ␤-keratin sequences of
birds, alligator, and lizard claw have a very similar amino
acidic sequence from amino acids 39 –53 (Fraser and
Parry, 1996; Sawyer et al., 2000).
Using the “consensus secondary structure prediction”
program on the claw protein, it results that 105 amino
acids (73.9%) have a random coiled conformation and 37
amino acids (26.1%) an extended strand conformation,
while no ␤-sheet or ␣-helical conformation is present.
Fig. 11. Sequence comparison of the lizard claw (A), with human
KAP 5-1 (B) and a human cytokeratin type II (C). Stars indicate identities;
colon indicates conserved substitutions; period indicates semiconserved substitutions detected by ClustalW software. Glycine residues
are evidenced in gray shadows. Underlines indicate the glycine-rich
regions (mammalian-like) in N- and C-terminal ends of the lizard protein,
while the boxed sequence indicates the central feather-claw-like sequence. Accession numbers of sequences analyzed are reported in
Table 1.
Cellular Synthesis and Distribution of Keratins
keratin packets and bundles contain this small protein.
The results confirm and overlap with the immunolocalization previously seen using antibodies directed toward
avian ␤-keratins (Alibardi and Sawyer, 2002; Alibardi et
al., 2004a, 2004b; Alibardi and Toni, 2005). The labeling is
uniform over ␤-keratin filaments and no fibrous structure
(reminiscent of cytokeratin filaments) can be detected at
high magnification. The study also confirms a previous
study that showed the presence of a low immunolabelig for
cytokeratin in ␤-keratin cells (Alibardi, 2000). The latter
observation indicates that the antiserum labels the amorphous matrix contained in ␤-filaments that show a thickness of 3– 4 nm (the ␤-keratin pattern) (Maderson, 1985;
Landmann, 1986). Both in situ hybridization and immunological results indicate that these basic proteins (pI
7.4 – 8.2) (Dalla Valle et al., 2005; Alibardi and Toni,
2006a, 2006b) belong to ␤-keratins.
The present observations extend a previous study on
mRNA isolation and localization for a lizard ␤-keratin
(Dalla Valle et al., 2005). The study showed that the
messenger codes for one small protein produced in ␤-cells
of regenerating epidermis, and that ␤-keratin mRNA was
highly expressed in differentiating ␤-cells. This indication
suggested that the utilization of a cDNA probe, which is
easier to handle and less susceptible to artifact degradations
than cRNA probes, was sensitive enough for our hybridization study. The present study also shows that the mRNA for
this protein is expressed also in embryonic epidermis and
therefore suggests that the protein is ubiquitous to all ␤-layers of the epidermis of P. sicula (and P. muralis).
Using our rat antisera against a 15–16 kDa lizard
␤-keratin, we have for the first time shown that the thick
Fig. 12. Sequence comparison of the lizard claw sequence (A), with
chicken scale ␤-keratin (B) and a chicken feather ␤-keratin (C). Stars
indicate identities; colon indicates conserved substitutions; period indicates semiconserved substitutions detected by ClustalW software. Glycine residues are evidenced in gray shadows. Underlines indicate the
glycine-rich regions (mammalian-like) in N- and C-terminal ends of the
lizard protein, while the boxed sequence indicates the central featherclaw-like sequence. Accession numbers of sequences analyzed are
reported in Table 1.
The nuclear localization and the structure of the gene
for this proteins (no introns appear to be present inside
the coding region; Dalla Valle, personal communication)
suggest that the probe can directly label the primary transcript inside the nucleus (Dalla Valle et al., 2005). The
presence of sparse-labeled clusters from the inner part of
the nucleus to the nuclear membrane and then mainly in
the cytoplasm may be associated with the movement of
the messengers toward the cytoplasm. The mRNAs are
present not only in the “free” cytoplasm, but often also
among the small ␤-keratin packets or associated in the
periphery of larger ␤-keratin filaments. This confirms previous studies using tritiated proline that indicated that
synthesis and polymerization of this protein occurs on the
surface of ␤-keratins filaments (Alibardi, 2004). This process is also typical for the synthesis of feather (␤-)keratin
in barb and barbule cells (Kemp et al., 1974).
1982) and histochemical (Alibardi, 2001, 2003) studies.
These putative cystein-rich proteins remain to be isolated
and sequenced in lizard epidermis.
The lizard scale protein has a molecular weight, amino
acid composition, and predicted secondary conformation
very different from that of ␣- or cytokeratins (Fuchs et al.,
1987; Steinert and Freedberg, 1991). The scale proteins
localize in the same sites (␤-keratin filaments) where a
prevalent incorporation of tritiated proline was previously
shown (Alibardi, 2001; Alibardi et al., 2004a). The scale
glycine-proline-rich protein enters in the composition of
␤-keratin filaments, which rapidly replaces cytokeratin
bundles present in early differentiating ␤-cells (Alibardi,
2000). This process resembles the localization of keratinassociated proteins among trichocytic keratins of mammalian hairs (Gillespie, 1991; Powell and Rogers, 1994; Rogers, 2004). In comparison to mammalian glycine-tyrosinerich proteins of 7–12 kDa, the glycine-cystein-proline-rich
(varanus claw) and glycine-proline-rich (lizard scale) proteins are larger proteins with a molecular weight of 13–16
kDa and with a lower content in tyrosine.
The ultrastructural and molecular data suggest that
both the lizard scale and claw glycine-proline-rich proteins have the composition of KAP isolated in mammalian
corneous appendages (hair, nail, etc.) (Gillespie, 1991;
Powell and Roger, 1994). Ongoing studies from this laboratory indicate that at least two more members of this
protein type are present in the epidermis of P. sicula, and
that similar glycine-rich proteins are present in the epidermis of other lizards, snakes, and turtles (data not
Previous studies (Wyld and Brush, 1979, 1983) indicated that different ␤ (␾)-keratin types were present in
low amounts in reptilian ␣-layers and in higher amount in
␤-layers, as defined by histological and ultrastructural
studies (Baden and Maderson, 1970; Maderson, 1985;
Some Proteins of ␤-Keratin Filaments Resemble
Mammalian Keratin-Associated Proteins
The amino acid sequence of the scale epidermal protein
of P. sicula is very different from a proteins isolated from
the claw of a varanus lizard (Inglis et al., 1987). The latter
is an acidic protein of 13.1 kDa proteins (142 amino acids,
pI 5.6), richer in sulfur and glycine, and with an amino
acid composition similar to type II glycine-tyrosine-rich
proteins of mammals.
The in situ hybridization study indicates that the sequenced glycine-proline-rich protein (Fig. 10A) is present
in the ␤-layer of P. sicula scale. It is possible that our
primer selection has isolated mainly the messengers for
one of the minor fractions of hard keratins, relatively poor
in cystein and resembling type I tyrosine-rich proteins
(Marshall and Gillespie, 1982). If this may be the case,
other keratins richer in cystein should be present in
␤-cells, as indicated from biochemical (Gillespie et al.,
Fig. 13. Phylogram among different cytokeratins derived from some
representative of different groups of vertebrates, from fish to mammals
(bottom group in parenthesis), ␤-keratins from avian skin (intermediate
group in parenthesis), and keratin-associated proteins from hairs (KAPs;
upper group in parenthesis) that includes the scale and claw proteins of
lizard (boxed). Numbers indicate distances (percent divergence) between all pairs of sequence. Accession numbers of sequences analyzed
are reported in Table 1.
Landmann, 1986). It is likely that some ␾-keratins of
reptilian skin represent keratin-associated proteins localized in ␤-keratin filaments: the latter are present in high
amount in ␤-layers and in very low amount in ␣-layers of
reptilian epidermis. Therefore, aside from filamentous
proteins, termed ␤-keratin, at least some proteins composing ␤-keratin filaments are represented by glycine-rich
proteins. The filamentous nature of ␤-keratins present in
␤-keratin filaments remains uncertain and only the complete protein analysis of these filaments will clarify this
problem (Gillespie et al., 1982; Marshall and Gillespie,
1982; Wyld and Brush, 1983).
It is likely that ␣-layers of reptilian epidermis contain a
very small fraction amount of ␤ (␾)-keratins among the
prevalent bundles of cytokeratin. This has been detected
by immunological and autoradiographic methods in turtle
and lizard soft epidermis (Alibardi et al., 2004b; Alibardi
and Toni, 2006a, 2006b). Differently from the ␣-layer,
␤-layers contain higher levels of ␤ (␾)-keratins than ␣-layers (Wyld and Brush, 1979, 1983). Therefore, it is possible
that ␣- and ␤-layers of reptilian epidermis simply reflect
the massive activation of genes synthesizing keratin-associated proteins and ␤-keratins in differentiating ␤-cells
or their nearly complete repression in ␣-keratogenic cells.
Reptilian ancestors of both mammals and birds might
have produced glycine-rich proteins. The presence of more
glycine-proline-rich and glycine-cystein-rich proteins in
lizard epidermis indicates that also in reptiles, like in
mammals, more members of KAPs are present. The latter
conclusion has important implications for the general evolution of KAPs in amniote skin.
The two proteins so far sequenced in extant lizards
(with others unpublished but similar sequences; data not
shown) suggest that these proteins represent the prototype of a class of small proteins that might have been
present in basal amniotes (reptiles by definition). Some of
the genes coding for these proteins were later selected in
the diapsid/anapsids (sauropsids) that evolved in modern
reptiles and birds, while others proteins were selected in
synapsid and therapsids that evolved into mammals. The
genes coding for these proteins were inherited and largely
modified to produce avian ␤-keratins, including scale,
claw, beak, and possibly also feather keratins, the smallest (Gregg and Rogers, 1986; Sawyer et al., 2000). In
mammals, KAPs evolved instead to serve as hard material
for keratinized appendages such as hairs, claws, and
horns (Gillespie, 1991; Marshall et al., 1991; Powell and
Rogers, 1994).
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