Original research or treatment paper
Advances in identifying archaeological traces
of horn and other keratinous hard tissues
Sonia O’Connor1, Caroline Solazzo2,3, Matthew Collins2
1Archaeological Sciences, Division of AGES, University of Bradford, Bradford, UK, 2BioArCh, Department of
Archaeology, University of York, York, UK, 3Museum Conservation Institute, Smithsonian Institution, Suitland,
MD, USA
Despite being widely utilized in the production of cultural objects, keratinous hard tissues, such as horn,
baleen, and tortoiseshell, rarely survive in archaeological contexts unless factors combine to inhibit
biodeterioration. Even when these materials do survive, working, use, and diagenetic changes combine to
make identification difficult. This paper reviews the chemistry and deterioration of keratin and past
approaches to the identification of keratinous archaeological remains. It describes the formation of horn,
hoof, baleen, and tortoiseshell and demonstrates how identification can be achieved by combining visual
observation under low-power magnification with an understanding of the structure and characteristic
deterioration of these materials. It also demonstrates how peptide mass fingerprinting of the keratin can be
used to identify keratinous tissues, often to species, even when recognizable structural information has not
survived.
Keywords: Horn, Hoof, Baleen, Tortoiseshell, Mineral preservation, Keratin, Peptide mass fingerprint, Mass spectrometry
Introduction
Horn, hoof, baleen, and tortoiseshell are all animal
hard tissues that have been exploited as raw materials
for functional or decorative applications. They are
tough, flexible, mouldable, and fusible natural plastics;
bio-composites of the protein keratin and a calcium
mineral, the latter being a minor component and prob-
ably mostly bioapatite.
Horn has always been widely available and was
commonly used in the past as a raw material for the
production of cultural objects as diverse as composite
bows, drinking vessels, powder horns, and inkwells
(MacGregor, 1985, 1991; O’Connor & Duncan,
2002; Richards & Gardiner, 2005), sword handles
(Stead, 2006; O’Connor, 2013), helmet panels
(Doppelfeld, 1964; Bruce-Mitford & Luscombe,
1974), longbow knocks (Jones, 2003), lantern panes
(Richards, 2005), reliquaries (Ryan, 1988) hammer
heads, knife and fork handles, spoons, and spectacle
frames. Compared to horn, the use of hoof has been
more limited; its utility restricted by its size and
shape. Whole cattle or horse hooves can be lidded to
form boxes or cut for buttons, but smaller sheep or
goat hooves have little use other than as toggles or
pendant ornaments.
Baleen, the fiber-fringed horny sheets from the
mouths of filter-feeding whales (Mysticeti), was prob-
ably always available to coastal communities in small
quantities through the opportunistic scavenging of
stranded whales and later as a by-product of the
hunting of various species of baleen whales for pro-
ducts such as oil and meat. There is evidence of
exploitation of cetacean material from the
Mesolithic (Clark, 1947; Whittle, 2000; Mulville,
2002). Whaling on a commercial scale was under-
taken by the Basques as early as the twelfth
century (Aguilar, 1986), and as whaling developed
so did the demand for baleen. From the late thir-
teenth century, it was used in the construction of
crossbow staves, for armour and tourney equipment
(Credland, 1981; Moffat et al., 2008). Strips of
baleen were used for shaping and supporting items
of dress, such as the farthingale, stomachers, and
stay busks. Baleen is the ‘whalebone’ of corsetry
(Sorge-English, 2011). By the eighteenth century,
baleen was a notable import into the UK from the
Dutch whalers (Beck, 1882) but was most widely
available at the height of the nineteenth-century
whaling industry when additionally it was used in
the production of brushes, cricket bat handles,
Correspondence to: Sonia O’Connor, Archaeological Sciences, Division of
AGES, University of Bradford, Bradford, West Yorkshire BD7 1DP, UK.
Email: s.oconnor@bradford.ac.uk
© The International Institute for Conservation of Historic and Artistic Works 2014
MORE OpenChoice articles are open access and distributed under the terms of the Creative Commons Attribution License 3.0
Received December 2013; revised paper accepted March 2014
DOI 10.1179/2047058414Y.0000000134 Studies in Conservation 2014 VOL. 0 NO. 0 1
umbrella stays, furniture inlays (Stevenson, 1907;
West & Credland, 1995; Wilkinson, 2003), and
almost any application where sheets, strips, or rods
of a tough but flexible material was required. Only
two archaeological finds of baleen are known to
the authors from the whole of the UK, one from
Perth (Moffat et al., 2008) and one from Westward
Ho (O’Connor & O’Connor, in preparation). The
picture is similar in the Netherlands where just
three finds of baleen have been published (Bartels,
2005; Rijkelijkhuizen, 2009).
Tortoiseshell is acquired from the hard protective
shell of large marine turtles and has always been an
expensive import in Europe. Used for luxury items,
it is occasionally recovered from Roman and later con-
texts (O’Connor, 1987; Rijkelijkhuizen, 2009) and is
characteristically used for inlays, combs, and other
items that show off the mottled figuring of the turtle
shell to best advantage. It could be argued that it
would be more correct to call this material ‘turtle
shell’ but in the biological literature this is used to
refer to the entire shell of the turtle, both the external
keratinous shield and the underlying boney plates
(Zangerl, 1969; Gilbert et al., 2001). In historic texts
relating to the acquisition and working of the kerati-
nous scutes (scales) themselves (e.g. Smith, 1770;
Siddons, 1837), the single word ‘tortoiseshell’ is
employed. This term remains in wide use today in
the curatorial and conservation literature and so will
be used here.
When compared to other animal hard tissues, such
as bone, antler, and ivory, these keratinous materials
are greatly underrepresented in archaeological assem-
blages, even when taking into account issues of avail-
ability. The reasons for this are poor preservation
and the problems of identification. Most archaeologi-
cal evidence of horn working has been in the form of
caches of the boney horn cores, either whole or
sliced (Wenham, 1964; MacGregor, 1985; Armitage,
1990; MacGregor, 1991) but rarely the horn itself.
The extensive excavations at York, UK, for instance,
have over the years only recovered a few tens of horn
fragments but many hundreds of horn cores.
The perspective of this paper is that of two archae-
ological scientists and a chemist whose collective
experience is largely of European species and burial
conditions. It draws together current understanding
of the chemistry, structure, and decay of keratin, and
reviews and evaluates techniques used in the identifi-
cation of keratinous hard tissues. It presents the cri-
teria for visual identification from structural features
and patterns of deterioration and explores the poten-
tial of peptide mass fingerprinting for the identifi-
cation of very poorly preserved material. By
presenting these two quite different investigative
methods in the same paper, we aim to show their
respective strengths and weaknesses, and the benefits
of an integrated methodology.
Keratin biochemistry and structure
Keratinous tissues arise from the dermis and epidermis
of the skin, but as the secretion of keratin into the cells
(cornification) causes them to die, these are not living
tissues. Keratins are fibrous proteins, of which there
are two major classes; helically organized alpha-kera-
tins, formed by mammals, birds, and reptiles, and
beta-keratins, which have a pleated sheet structure
and are only produced by birds and reptiles
(McKittrick et al., 2012).
Hard (trichocyte) alpha-keratin-made tissues, such
as hair, claw, horn, hoof, and baleen, and soft (cyto-
skeletal) alpha-keratin-made tissues, such as the
stratum corneum of the skin, are formed of dimers
of type I (acidic) and type II (neutral-basic) proteins
aligned into intermediate filaments (Plowman, 2007),
embedded in a non-filamentous protein matrix
(keratin-associated proteins). Keratin proteins have
low sulfur content in comparison to some of the
matrix proteins, which are rich in the sulfur amino
acid cysteine. The sulfur–sulfur cross-linking between
cysteines in the keratin’s non-helical end and tail
domains and the cysteine-rich keratin-associated pro-
teins contribute to the strength and rigidity of the
tissues. ‘Hard’ keratins have, however, a higher sulfur
content than ‘soft’ keratins, which are also present as
minor components in horn and hoof (Solazzo et al.,
2013b). The structural organization and orientation
of the intermediate filaments vary from tissue to
tissue and influence their mechanical properties.
These filaments are randomly oriented in skin cells,
but lie at an angle of 20–30° to the direction of
growth in horn cells (Marshall et al., 1991), while in
the tubules of horse hoof they are aligned in the direc-
tion of the tubules and perpendicular to the tubules in
the intertubular matrix, helping to resist crack propa-
gation (McKittrick et al., 2012).
Beta-keratins are the structural proteins of birds’
beak, claw, and feather and reptile skin. They do not
occur in mammals and instead of forming intermedi-
ate filaments they stack together to produce ß-
pleated sheet structures. They intermingle with ‘soft’
alpha-keratins in the shelled epidermis (carapace and
plastron) of turtles (Toni et al., 2007; Alibardi et al.,
2009; Dalla Valle et al., 2009; Dalla Valle et al.,
2013) and the ratio of beta- to alpha-keratins deter-
mine the degree of hardness of the scutes (Dalla
Valle et al., 2009). For instance, the carapace of the
hard-shelled Florida redbelly turtle Pseudemys
nelsoni, has a higher ratio of beta-keratins to alpha-
keratins compared to that of the soft-shelled turtle
Apalone spinifer (Dalla Valle et al., 2013). The
turtle’s beta-keratin proteins are at least half to one-
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 02
third the molecular weight of their alpha-keratin
counterparts. The central region involved in the beta-
sheet structure (4.8–8.3% in turtle’s beta-keratins) is
highly conserved and rich in proline (P) and valine
(V) (Dalla Valle et al., 2009).
Hard keratinous tissues also generally contain some
amount of calcium phosphate salts, from 0.1% to
nearly 15% (dry weight), depending on their form
and function. Zones of calcification which help main-
tain the stiffness or the shape or sharpness of a struc-
ture through differential wear, have been noted in
cat, lion, and rabbit claw (Pautard, 1964; Halstead,
1974), horn (Hashiguchi & Hashimoto, 1995;
Hashiguchi et al., 2001) baleen (Pautard, 1970;
Szewciw et al., 2010), rhinoceros horn (Hieronymus
et al., 2006) hoof, porcupine quill, and bird beak
(Pautard, 1964). Baleen is the most highly calcified
of these tissues and the level varies from 1% in
minke whale to 14.5% in Sei whale of bioapatite in
the non-filamentous matrix (Szewciw et al., 2010).
Decay and preservation
The decay of keratin, like that of bone (a composite of
protein (collagen and osteocalcin) and mineral (bioa-
patite)), occurs along three pathways: the chemical
deterioration of the organic and the inorganic com-
ponents, and through biodeterioration. Well-preserved
bone, however, is readily recovered from burial
environments where horn does not survive at all.
This is not just because bone has a much higher
mineral content (ca. 70 wt%) but because of the inti-
mate way in which the organic and inorganic com-
ponents are distributed and rigidly packed, affording
protection to each other (Collins et al., 2002). In
even the most mineralized of the keratinous tissues
the protein is completely exposed to both chemical
and microbial attack. The rate of deterioration due
to chemical changes is largely determined by the temp-
erature, moisture content, and pH of the burial
environment, although this is complicated by the con-
dition of the material prior to burial and potentially by
the processing it has undergone in its transformation
from raw material to cultural object. Loss of the
mineral component will occur in low pH conditions
but it is high pH that will have the most deleterious
effect on keratin. However, the major cause of
keratin deterioration is rapid biodegradation. In the
soil, keratins are degraded by enzymes produced by
specialized keratinolytic microorganisms, primarily
fungi. At least 300 fungi are known to use keratins
as a source of nutrients (Błyskal, 2009), but the mech-
anisms of attack have only been studied in detail with
relationship to hair and wool (Wilson et al., 2007,
2010; Solazzo et al., 2013a). The fungal enzymes
denature the constituent proteins and break the disul-
phide bridges. Different elements of the hair structure
are more or less resistant to this attack but surface
erosion, radial penetration, and lateral tunneling by
fungal hyphae are accompanied by separation of the
hair’s structural components, loss of the protective
cuticle, the destruction of the matrix protein, then
the fibrous protein and, finally, the total collapse of
the fiber.
As keratinolytic fungi present in the soil become
active upon the addition of keratin, preservation is
largely dependent on the inhibition of biodeterioration
by environments unfavorable to aerobic fungal activity
(e.g. lack of oxygen, low temperatures, limiting pH,
very low relative humidity), the presence of biocides,
or a combination of the two. Examples include the sur-
vival of finger nails and hair in the deeply frozen
bodies of the Franklin expedition (Beattie & Geiger,
1987) and the Tyrolean Ice Man (Spindler, 1993),
the desiccated sand burials (Dzierzykray-Rogalski,
1986) and embalmed mummies of Ancient Egypt
and the acid bog burials of Northern Europe
(Brothwell & Gill-Robinson, 2002) with their preser-
ving combination of low pH, dark, anoxic, water-
logged conditions and the antiseptic nature of some
of the component vegetation. It is no surprise that
woollen textiles and objects in horn and other kerati-
nous tissues are also preserved with many of these
inhumations. However, through a combination of
sub-aerial weathering and biodeterioration, material
left on the ground surface for any length of time will
not survive to be incorporated into archaeological
deposits in a recognizable form. This is probably
why so little keratinous material survives, for instance,
in the deep, anoxic, waterlogged deposits of urban
sites in York, UK (O’Connor, 1987) or similar sites
across Amsterdam, Netherlands (Rijkelijkhuizen,
2009) despite otherwise favorable conditions for pres-
ervation. That this absence of evidence is not just evi-
dence of absence but largely an issue of preservation is
corroborated by the many examples of traces of horn
discovered as mineral preserved organic (MPO)
remains during the conservation of metal objects
from such sites.
Mineral preserved organic remains
Even in apparently unfavorable burial conditions, ker-
atinous tissues do survive as MPO remains where they
have lain in close contact with corroding metals. The
metal corrosion products pervade the organic
material, coloring, or coating, and/or invading its
structures and affording some level of protection to
the organic matrix. The survival of plant- and
animal-derived MPO remains have long been recog-
nized (Biek, 1963) and this knowledge has both influ-
enced the direction of development of conservation
techniques and greatly increased our understanding
of the organic components of archaeological
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 3
assemblages. Even in anoxic, waterlogged sites such as
Coppergate, York, many more instances of MPO horn
are found than waterlogged examples, providing a
much clearer picture of its widespread use.
The mechanisms of mineral preservation have been
explored mostly in relation to wood and textiles in
contact with iron and copper alloy in the context of
inhumations where these materials are often intimately
combined. The extent (or absence) of MPO remains,
the form the mineralization takes (positive or negative
cast, or both) and the quality of detail preserved
depends on factors such as the relative rates of deterio-
ration of the organic and metal components, which
organic and metallic materials are involved, the con-
centration of metal ions in solution, the rate of ion
take-up by the organic material, and the constituents
of the ground water (Keepax, 1975; Janaway, 1983,
1989; Turgoose, 1989; Gillard et al., 1994). Positive
casts, where the metal ions penetrated the cells and
fibers of the organic material, produce the best preser-
vation of detail. The corrosion products of several
metals have biocidal properties that also help preserve
the organic matrix of the material while mineralization
proceeds. Foremost among these is copper, although
MPO remains on silver and lead have been observed
(Edwards, 1974; Stock, 1976). The metal ions released
by the oxidation of iron, however, are not toxic to the
microbiota and the cascade of iron oxidation products
does not passivate the surface, unlike copper. Taken
together this means that biodeterioration is not
directly affected by iron corrosion, and indeed in
some cases can be accelerated by it, but rapidly corrod-
ing iron can engulf organic material demonstrably
conferring indirect protection.
Fig. 1 is a most spectacular example of the MPO
preservation of keratinous material. This is one of
two coins that had been placed over the eyelids of an
eighteenth-century corpse buried near St John’s
harbor Newfoundland, Canada, that were recovered
with a patch of woollen shroud and eyelashes on
either side (Hett, 1980). A slow rate of corrosion has
kept the copper ion concentration low and produced
positive casts, with minimal green staining, while inhi-
biting fungal attack of the keratin. The results are
MPO remains that have retained much of the appear-
ance, properties, and micro structure of the original
materials, including the pigment granules in the eye-
lashes. This contrasts sharply with the preservation
of MPO keratin by corroding iron. Without the protec-
tion of a biocide, decay can be so fast that before min-
eralization has penetrated any distance, the keratinous
material is reduced to a formless mush. Recognizable
MPO remains may be found, if at all, only as an
impression in the corrosion at the original surface of
the metal object. For instance, evidence of joins and
grain direction of the horn handle components of
one of the South Cave Iron Age swords were clearly
preserved against the iron tang but the shape and
thickness of the guard, grip, and pommel were entirely
lost (O’Connor, 2013).
Visual identification
Even when these keratinous materials do survive in the
archaeological record they may not be recognized.
Horn, for instance, is a material with which we are
unfamiliar today, its place having almost entirely
been supplanted by synthetic plastics. The shape of a
complete horn is perhaps unmistakable but once frag-
mented, worked, or decayed its identification can be
problematic.
Most guides to the identification of worked animal-
derived hard tissues are centered on osseous materials
with a preoccupation with ivory and its substitutes
(Penniman, 1952; Krzyszkowska, 1990). Campbell
Pedersen’s (2004) book includes a range of keratinous
materials and is a very useful resource for the identifi-
cation of historic objects but is of limited use when
faced with degraded archaeological finds. Only a few
publications address the identification of archaeologi-
cal keratin. O’Connor (1987) is mostly based on water-
logged finds recovered from sites in York and
Rijkelijkhuizen (2008) on finds from Amsterdam.
Florian (1987) briefly describes the features of these
materials in the context of their decay in marine
environments and Lauffenburger (1993) covers the
identification, historic uses, and past conservation
treatments of baleen. The work presented here builds
on these papers and the results of O’Connor’s recent
study into the identification of animal hard tissues.
While the identification of MPO remains through
scanning electron microscopy is well established
(Keepax, 1975; Janaway & Scott, 1983; Watson,
Figure 1 MPO remains of keratinous materials first
published in Hett (1980). © Government of Canada, Canadian
Conservation Institute, CCI 2003063-0001.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 04
1988), it is not usually necessary for the visual identi-
fication of keratinous tissues, however preserved. SEM
offers high magnification, excellent depth of field and
resolution, and environmental chambers now allow
objects to be examined without the need for sampling
and coating, but cannot reveal variations in color or
translucency. Many of the diagnostic features ident-
ified in this paper were visible, in good light, to the
unaided eye and have been captured using nothing
more sophisticated than a Panasonic Lumix DMC
TZ6, a compact digital camera with a good macro
facility. The more detailed characterization and
recording of these features were undertaken using a
Wild Heerbrugg M8 stereomicroscope or a Dino-
Lite (AM-7013MZT) 5 megapixel digital microscope
with polarizer, at magnifications typically between
20× and 50×, and occasionally at 200×. At higher
magnification, these keratinous materials can look
very similar, all being formed in part from compressed
layers of keratinized epithelial cells. At lower magnifi-
cations it is easier to see how these cells are organized
in relationship to other more specialized structures and
their orientation to any surviving natural surfaces.
Horn
True horns are found only on the skulls of ruminant
artiodactyls such as cattle, sheep, goat, and antelope.
The bone horn cores grow out from the frontal bone
of the skull and the horn develops from its epithelial
covering, forming a sheath with a finely layered
cone-within-cone structure. Horn has a longitudinal,
fibrous grain and the surface may have thin, uneven,
circumferential ridges marking the edges of sequential
cones, particularly toward the junction with the skull
(Fig. 2). The cross section of the horn is determined
by that of the horn core and in general is circular to
oval in cattle, elliptical in goat, and triangular in
sheep although there is much variation in the color,
texture, torsion, and external sculpting between
different domestic breeds today. Sheep and goat horn
can be translucent and virtually colorless while cow
horn may have a pale brown or greenish tinge, or be
layered and longitudinally streaked with colors
ranging from white to reddish brown or black, while
buffalo horn is black and opaque.
The horn accumulates throughout the life of the
animal (except that of the pronghorn antelope
(Antilocapra americana), which sheds its horn sheath
annually but retains the horn core) increasing the
thickness of the horn over the sheath and developing
a solid mass beyond the distal tip of the horn core.
The flattened cornified epithelial cells form growth
layers over the surface of the horn core. These layers
are periodically punctuated by tubules that run the
length of the horn and appear as oval or elliptical por-
osities between the layers in a transverse section. The
morphology of the layers and the distribution and
size of the pores vary between cattle, sheep, and goat
horn and can aid identification even when the shape
and color of the horn are altered by working and
decay.
Cow and ox horn have very few, relatively inconspic-
uous pores but the growth layers are wavy with a
compact boundary that give the unworked surface a
longitudinally corrugated appearance. In worn or
worked surfaces these corrugations may still be appar-
ent from the distribution of the pigmentation in the
truncated layers (Fig. 2), but pressed and polished
translucent horn can look like a homogeneous sheet
of plastic. With time, starting at the edges of the
horn, deterioration allows air to infiltrate between
the layers, making the corrugations visible once
again in either reflected or transmitted light. In trans-
verse section, the layers describe a series of intersecting
arcs of similar size. These can be particularly obvious
toward the centee of the solid part of the horn tip
where each layer follows the path of the layer below
around a central point and the synchronous intersec-
tions of the arcs produce a pattern of radially orga-
nized lines between the columns of arcs (Fig. 3A).
Moving out from the center, the arcs of each successive
layer are minutely longer as the circumference of the
horn increases. Eventually a critical length of arc is
reached and the next layer has additional arcs.
Occasionally two columns of arcs also merge but the
overall size and shape of the arcs is maintained
throughout the radius of the horn. Buffalo horn has
flattened, almost triangular, cross sections and the
layers of the horn in the solid tip form around a line
of small anomalies, instead of a single point. The
layers are finer and the wave produced by the arcs is
less distinct and sometimes absent.
Sheep and goat horn is often unpigmented and
transparent. The solid tip in transverse section has
an elongated or discontinuous central anomaly and
Figure 2 Detail of the naturally worn surface of a cattle horn.
© Sonia O’Connor.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 5
in some sheep horn the triangular path of the layering
around this can distinguish it from goat. The layering
does not have the pattern of intersecting arcs and
radial lines seen in cattle horn (Fig. 3B). Sheep horn
in particular contains many larger pores, that flatten
as the horn dries, and the layers follow an undulating
path over them. Hanausek (1907, p. 433) records
pores of 30–160 μm in sheep horn, while McKittrick
et al. (2010) record pores of 40 by 100 μm and a
mid-range tubule density of 22 mm−2. The horn of
the bighorn sheep (Ovis canadensis) has evolved to
withstand extreme loading impacts during fighting
and has been studied in detail (Tombolato et al.,
2010). The horn is particularly densely packed with
large tubules, up to 200 μm at their maximum diam-
eter, interleaved with layers of horn only 2–5 μm thick.
In dry conditions horn shrinks and concentric and
longitudinal cracks open revealing the lamella struc-
ture. The compact boundaries of the corrugated
layers of cattle horn can be very distinctive at this
stage. Deterioration in the soil leads to a loss of
color and transparency as the cells of the layers
begin to separate and disintegrate. If deterioration
continues eventually these sheets degrade to a fibrous
longitudinal mass and are lost. In alkaline conditions
horn will follow a different diagenetic pathway.
Chemical hydrolysis rapidly destroys higher order
structure producing a structureless gel that eventually
dissolves. Even if the further deterioration of this gel
was arrested, it is unlikely to be recovered during exca-
vation. In the case of the cattle horn tip fragment in
Fig. 4, the corrugated sheet structure is still recogniz-
able but some layers have decayed to a gel that has
dried, presumably after excavation, to a hard, resinous
material. It is not understood why this has occurred
only in certain layers nor are the precise details of
the original burial conditions knowable.
Where mineral preservation occurs, this fibrous
mass may survive as the metal object and its corrosion
provide some physical protection. Stained brown with
iron corrosion, MPO horn is often mistaken for wood
but examination under low magnification will indicate
a complete absence of the cellular organization of
wood, particularly ray cells (Fig. 5). With very
Figure 4 Fragment of horn tip (YAT 1977.7.16022) from
waterlogged deposits, (A) side view and (B) interior view (York
Archaeological Trust specimen).
Figure 3 Worked surfaces approximating to the transverse
section of the horn tip (A) of cattle (Hawley Collection
specimen) and (B) of sheep horn. © Sonia O’Connor.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 06
decayed horn a positive identification will require the
application of other, more destructive analytical tech-
niques (see below).
Hoof
Hoof forms as a capsule over the distal phalanx of the
feet of ungulates and is very similar to horn in color
and structure. In horse this is a single structure
(Fig. 6) but in cloven-footed animals, such as cattle,
the foot ends in two digits and each is covered by a sep-
arate sheath or claw. The fine detail of the hoof struc-
ture varies depending on the location within the hoof,
e.g. the wall or the sole.
Like finger nail, the hoof wall is formed from its
proximal edge, the junction between the hoof and
the skin of the foot. Here cells are produced by the cor-
onary corium, become cornified and die, and are
moved forward over the dermis as new cells are
formed behind them. Variations in growth rate can
produce slight textural features that ring the hoof hori-
zontally (Fig. 6). The surface of the corium has long
papillae from the ends of which tubules form within
the intertubular keratin matrix (Fig. 7). The tubules
lie parallel to the surface of the hoof and extend
from the corium to the base of the wall. This gives
the external surface of the hoof wall a finely striated
appearance (Fig. 8), the paired strae being the walls
of the hollow tubules. Variations in color and translu-
cency of the intertubular matrix may form stripes or
streaks in this direction. Toward the proximal edge
of dry hoof specimens, successive edges of thin layers
of the hoof may be visible and in polished surfaces
variation in the color relates both to the tubule direc-
tion and changes through the depth of the wall. The
sole of the hoof has its own corium that produces a
softer more pliable horn material.
Although primarily a tubular structure, the outer
portion of the wall has some layering. The fully kera-
tinized intertubular matrix consists of plate-like flat-
tened cells, the longitudinal axes of which lie parallel
to the direction of tubule growth (Marshall et al.,
1991). The inner surface of the wall is sculptured
Figure 5 Detail of MPO horn remains on an iron knife handle
with copper alloy rivets (York Archaeological Trust
specimen).
Figure 7 Detail of the proximal edge of the horse hoof wall
showing the upper ends of the tubules and the white ridges
on the interior surface of the wall. © Sonia O’Connor.Figure 6 Hoof sheath from a horse. © Sonia O’Connor.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 7
with unpigmented, vertical ridges (Fig. 9) that interdi-
gitate with the fine leaf-like laminae of the dermis
(some 550–600 in horse hoof wall; Pollitt, 2004).
Removed from the bone and cleaned of the dermis,
this inner surface appears white against the pigmented
intertubular matrix of the bulk of the wall and is
visible on the exterior of the hoof as a white line
between its wall and sole. The lower edge of the hoof
wall is worn away by contact with hard surfaces but
if the rate of growth outstrips the rate of wear this
can cause lameness and other health problems. To
avoid this, it is often necessary to trim the hooves of
domestic livestock (Fig. 10).
In young, healthy horses the outer surface of the
wall is covered in a distinctly smooth and shiny layer
(the periople) which flakes away in adult hooves. The
papillae of the coronary corium are 4–5 mm long
and taper to a point. The holes they leave behind are
easily visible to the eye in the concave, upper edge of
the wall of the detached hoof (Fig. 7). The tubules
these form in the bulk of the hoof wall are oval in
cross section and have a well-defined cortex of flat-
tened, unpigmented, keratinized cells arranged con-
centrically around the medulla (Figs. 8 and 9).
Where the cells of the intertubular matrix are darkly
pigmented with melanin, the tubules appear particu-
larly prominent (Pollitt, 2004). The tubules increase
in diameter but reduce in number from the outer
toward the inner region of the wall (Bragulla &
Hirschberg, 2003), up to 220 × 140 μm with a
medulla of 50 μm (McKittrick et al., 2012). The
Figure 10 Dried clippings from a horse hoof. © Sonia
O’Connor.
Figure 8 Details of the exterior of the horse hoof wall
showing the vertical striations formed by the tubules. © Sonia
O’Connor.
Figure 9 Photomicrograph of the worn lower edge of the
horse hoof wall with the white ridges of the interior surface
uppermost. © Sonia O’Connor.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 08
tubules toward the exterior of the hoof become quite
flattened parallel to the surface in a dried specimen.
In the region of the largest tubules there is proportion-
ately very little intertubular matrix between them.
McKittrick et al. (2010) records a mid-range tubule
density of 24 tubules per mm−2 while Pollitt (2004)
reports a decline from 22 to 11 mm−2 step-wise
toward the inner surface, with an overall mean
density of 16 mm−2.
In modern cattle, the hoof wall thickness is c.
4–4.5 mm and is formed from between 50 and 60
tubules per mm−2, embedded in the intertubular
matrix. In sections of fresh hoof, the tubules nearest
the claw surface are flattened parallel to that surface.
Deeper in, the tubules are more oval (of the order of
50 μm maximum diameter) but the layer nearest the
underlying laminar dermis, is virtually free of
tubules, in contrast to horse hoof (Franck et al.,
2006). The walls of the tubules are thinner than in
horse hoof (McKittrick et al., 2012)
Once removed from the living hoof the keratin
capsule loses moisture and shrinks. Hoof clippings
trimmed from the base of the wall crack and curl lat-
erally into tight coils with the exterior of the hoof
wall to the outside (Fig. 10). For this paper, it was
not possible to examine sufficient archaeological
examples of known hoof to detail the deterioration
but the tubular structure is persistent and can be
made more obvious by these changes. Cracks can pro-
pagate through the thickness of the hoof between
tubules and also between the flattened cells of the
intertubular matrix producing delamination roughly
parallel to the outer surface. Sections of hoof wall
can easily be mistaken for horn but should be dis-
tinguishable on the detail of edges perpendicular to
the tubules (Figs. 3 and 9). Gelatinization in patches
producing irregular holes through the wall of an
unworked cattle claw has also been observed from
the material at York.
Baleen
Baleen plates are thin, roughly triangular sheets
(Fig. 11) of keratinous tissue that hang in racks,
like the closely stacked vanes of vertical window
blinds, down from the upper jaw on either side of the
mouths of the baleen whales. The main plates,
the sheets at the outer (labial) edge of the mouth, are
the longest and are usually flanked on their inner
lingual edge by smaller accessory plates. The fibers,
or bristles, of the inner edge of each sheet mesh
together with those of adjacent sheets to form a sieve
against which the food of the whale is trapped.
Sections through a baleen plate reveal a tripartite
structure consisting of a covering layer of finely lamel-
lar, horn-like material surrounding a filling of tightly
packed, longitudinal tubules, with well-defined
cortex and medulla, cemented together by intertubular
horn (Halstead, 1974). The horny covering wraps
smoothly over the longitudinal edges except where
the tongue has worn it away and the tubules are
exposed as the bristly fringe. This inner (lingual)
edge tapers to the lower (distal) end of the plate,
which terminates with a tuft of bristles.
Baleen is formed through the cornification of epi-
thelial cells of the gum and grows continuously. Each
main plate corresponds to a transverse dermal ridge
in the gum that has numerous extremely long dermal
papillae along its edge from which the tubules
develop. The keratinized cells of the tubules are flat-
tened parallel to the tubule’s surface and oriented con-
centrically around the medulla. The outer horny
covering of the baleen plate consists of flattened, cor-
nified cells arranged in fine layers parallel to the
surface of the plates and is formed by the epithelium
on the sides of the dermal ridges. The surfaces of the
Figure 11 A sheet of baleen (KINCMW7-7-76) lit to show the
natural surface texturing (Hull Museums specimen).
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 9
plates are often striated longitudinally but also show
shallow undulations across the width of the plate
that may relate to variations in growth rate (Fig. 11).
The development and structure of baleen in the fetal
and adult whale is covered in detail by Fudge et al.
(2009), who present an annotated translation of the
seminal 1883 paper by Tycho Tullber.
The tubules of a single plate have a range of diam-
eters but the average tubule diameter depends on the
species (Figs. 12 and 13). The tubule stiffness also
varies between species and is related to the level of cal-
cification (Szewciw et al., 2010). Melanin pigmenta-
tion is associated both with the tubules and the
covering layers. The size, shape, and thickness of the
baleen plates vary with species and their color can be
bluish black, brown, or pale honey colored, with longi-
tudinal, darker streaks (Watson, 1981). The color,
shape, and size of the plates and the relative bristle
density and diameters of different whale species are
detailed by Young (2012).
In worked objects where the tripartite structure is
retained, the material is easily identified as this is a
feature unique to baleen. Stripped away from the
fibrous core, the outer horny covering layers of the
baleen can be difficult to distinguish from horn.
The size of the sheet may rule out horn and the
combination of fine vertical striations and horizontal
undulations on the exterior may help confirm identifi-
cation, if these have not been lost through working
(Fig. 11). The layers of horny baleen are finer,
flatter, and less fibrous in appearance than cattle or
sheep horn. On edges that approximate to a transverse
section the layers, although wavy, are not organized
into columns of nested corrugations like cattle horn
nor are they interspersed with flattened tubules as in
sheep horn. On their own, the tubules can be mistaken
for a range of mammal hair, depending on their diam-
eter, but under high magnification the surface is seen
to be formed from a mosaic of epithelial cells, which
vary between species (Pfeiffer, 1992), rather than
cuticular scales.
Long exposure to light produces a graying of dark
baleen. As baleen dries it shrinks and becomes increas-
ingly stiff and brittle and the tubules become rigid and
easily fracture. Thin layers of the horny covering
exfoliate along incised designs and cut edges
(Fig. 14). On the surface of the plates, oval-shaped
blisters can form where the outer layers detach
around fine longitudinal splits. In archaeological
examples the horny covering layer and tubules
usually prove more persistent than the intertubular
matrix. The many examples preserved in Arctic per-
mafrost may appear to be in a reasonable condition
until they thaw when they are found to have delami-
nated and become detached from the tubular layer
(Wardlaw & Grant, 1994). This was also the case
with the two examples found in waterlogged deposits
in the UK; a worked object from Westward Ho!
Figure 12 The edge of a baleen stomacher (KINCM 2005-
2401) providing a longitudinal section through the thickness
of the baleen plate. The tubular layer lies horizontally between
the layers of the horny covering (Hull Museums specimen).
Figure 13 The end of a baleen stomacher (KINCMW5-90-76)
providing a transverse section of the tubular layer (Hull
Museums specimen).
Figure 14 Surface delamination around the incised
decoration of a baleen stomacher (46.1983.246 Leeds
Museums and Galleries specimen).
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 010
(O’Connor, in preparation) and unworked plate frag-
ments (Fig. 15) from Perth (Moffat et al., 2008).
Tortoiseshell
The bodies of tortoises, turtles, and terrapins are
encased in protective shells formed from a shield of
keratinous scutes (scales) over a boney mosaics of geo-
metric plates. Large marine turtles, most particularly
hawksbill turtle (Eretmochelys imbricate) but also
green turtle (Chelonia mydas), produce thick kerati-
nous shields with excellent working properties. These
turtles have large thick scutes over the top (vertebral
scutes) and sides (pleural scutes) of the carapace
(dorsal plate). Although the scutes of this shield can
be dark, matt, and algal covered in life, when separ-
ated from the bone and worked, the material trans-
forms into glossy, vibrant, translucent, mottled
tortoiseshell (Fig. 16). The tortoiseshell of the plain
colored scutes of the plastron (ventral plate) is com-
paratively thin but the scutes are large and this is
also utilized for inlays and coverings.
The boney mosaic is covered by a germinal layer
that forms the scutes but the edges of the scales do
not align with the junctions of the boney plates. The
polygonal cornified cells of tortoiseshell are flattened
parallel to the surface of the germinal layer and
coalesce into a compact layer about 20 cells deep.
Each layer is separated from the next by a distinct
boundary layer that appears to form in response to
seasonal changes in metabolic activity. Despite this,
the keratinized cells form a quite transparent, pale
colored ground. Melanin granules within the cells
produce the dark mottling of the tortoiseshell, which
is seen throughout the thickness of the material. In
many species of turtle the outer (older) layers are
periodically shed but in the hawksbill and green
turtle the scutes get appreciably thicker over time. As
the turtle maintains the same number of scutes
throughout its life, the area of successive layers
increases as the turtle grows, the discontinuity
between successive layers producing the appearance
of contour lines on the map of a hill. However, the
growth in area of the scute may not be the same in
all directions so the thickest part of the scute may be
toward one particular edge in some species, rather
than centrally located (Hanausek, 1907; Zangerl,
1969; Solomon et al., 1986). The distinctive white
pattern of curved, diffuse-edged lines on the underside
of the scutes (Fig. 17) is not described in the biological
literature but may indicate bands of transitional cells
(Dalla Valle et al., 2009) in the process of changing
from epithelium to fully cornified shield.
The tortoiseshell of the plastron of hawksbill turtle
is plain yellow, although occasionally blotched with
black, while that of the green turtle is thinner and
varies from yellowish white to dark gray or bluish
green in different individuals. The scutes of the cara-
pace are more distinctive and their colors and pattern-
ing are central to the identification of each species.
Figure 17 Detail of the natural patterning on the underside of
a tortoiseshell scute from a hawksbill turtle. © Sonia
O’Connor.
Figure 15 Fragment of waterlogged baleen from the Horse
Cross site, Perth, Scotland, excavated by the Scottish Urban
Archaeological Trust (SUAT) in 2004. © Sonia O’Connor.
Figure 16 Tortoiseshell from (A) hawksbill turtle and (B)
green turtle. © Sonia O’Connor.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 11
Those of the hawksbill vary in color between scutes
and between different oceanic populations
(Hanausek, 1907, pp. 433–435). The translucent back-
ground color of this tortoiseshell can be a blonde or
pale fawn, sometimes with a hint of green, or golden
yellow through to reddish amber, and densely but
unevenly mottled with blotches of dark brown or
almost black spots (Fig. 16A). These spots can seem
randomly spaced or merge to form irregular stripes
covering the majority of the surface of the scute.
Green turtle carapace scutes can be plain but usually
they are patterned with radiating brown stripes as if
painted with broad brush strokes (Fig. 16B) from the
highest point at the edge of the vertebral and pleural
scutes. The background color can vary from pale
yellow to dark brown, often with a greenish tinge.
Although these patterns of mottling are seemingly
very different, it is surprisingly difficult to distinguish
the species in a finished object. Not only is this
because the size and form of the object may provide
only a very limited view of the pattern of mottling
but as tortoiseshell can be readily and invisibly fused
to itself, the object may be cut from laminated sheets
or reformed, randomly oriented scraps. Hanausek,
writing in the early twentieth century, notes that the
plain colored tortoiseshell of the green turtle is artifi-
cially colored to imitate that of hawksbill, which is
deemed to be ‘true’ tortoiseshell (Hanausek, 1907,
p. 434).
The light colored ground of tortoiseshell is relatively
transparent and featureless. Where this pale ground
meets the darker mottling there are no hard bound-
aries. At its densest the mottling may be quite
Figure 18 Photomicrograph of the pigment distribution in
tortoiseshell from a hawksbill turtle (Horniman Museum
specimen).
Figure 19 Successive stages in the deterioration of worked tortoiseshell, (A) loss of cohesion at the edges of the layers (Hawley
collection specimen), (B) surface delamination and degradation (WCM 100879 sf 60), (C) degraded layers throughout the
thickness of the material, viewed from the edge (Hawley collection specimen), (D) loss of degraded cells leaving only the
boundary layers (2006.5201.923 kind permission of York Archaeological Trust).
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 012
opaque but at the edges the two gradually diffuse into
each other throughout the thickness of the material. If
the piece is relatively thin, it is possible to see clusters
of melanin-pigmented cells under low magnification
and transmitted light. These clusters are spaced
increasingly further apart as the edge of the mottling
merges with the transparent ground (Fig. 18).
Freshly polished, tortoiseshell is entirely smooth but
in time, probably through a combination of dehy-
dration and long exposure to light, the edges of the
fine layers become visible giving an appearance like
watermarked silk where they are cut through by
surface working (Fig. 19A).
The condition of archaeological finds of tortoise-
shell varies greatly. As deterioration proceeds, the
surface layers begin to separate, the cells of the layers
begin to degrade, and the bonds between them are dis-
rupted. Groups of cells are lost and the surface devel-
ops awhitish, opaque, granular appearance (Fig. 19B).
This deterioration proceeds in from the edges of the
object along the layers. Eventually the layers are com-
pletely reduced to a granular state but the junction
between each layer is still marked by the boundary
layer (Fig. 19C). This thin film-like layer can persist
after the granular material disintegrates, leaving tor-
toiseshell comb teeth, for instance, with the appear-
ance of the vanes of a miniature fan (Fig. 19D). All
of these stages in the deterioration of tortoiseshell
were observed together in a single cache of comb-
making debris from the site of the nineteenth-century
Rougier family comb factory in York.
Of all the keratinous tissues described, tortoiseshell
is the most likely to be imitated in other materials
because of the high price of this exotic import and
these imitations may be recovered from archaeological
deposits. Low magnification microscopy can be used
to distinguish these from tortoiseshell. Mottling with
sharply defined edges, swirls in the pigmentation,
and irregular shaped, colored, or reflective particles
in the pigmented areas or the pale ground are indica-
tive of a plastic imitation tortoiseshell. Fine, random
crazing of the surface also indicates a decayed
plastic. If traces of striae are seen within the material
this suggests that the material is pressed blond horn
used in imitation of tortoiseshell. In this case, any
mottled coloration will be limited to the surfaces of
the material. Surface-only coloration may also be
observed where plainer material from plastron or car-
apace has been enhanced by painting to imitate more
highly prized and priced ‘true’ tortoiseshell. In
addition, thin veneers of tortoiseshell can be bonded
to either side of a sheet of pressed, blond horn, as
O’Connor has observed on the backs and handles of
Victorian hair brushes. This can look very like solid
tortoiseshell unless the striae of the horn become
visible as the materials deteriorate.
Other approaches to identification
Where researchers have been unfamiliar with the fea-
tures of these keratinous materials, the evidence is
ambiguous or the material badly degraded, several
chemical identification techniques have been
employed. Wellman (1997) investigated the use of the
iodine azide spot test to detect high sulfur protein to
help distinguish horn from leather. The test could
not distinguish between different sources of keratins
or protein and non-protein sources of sulfur.
Anheuser & Roumeliotou (2003) were successful in
distinguishing microscopic samples of highly degraded
animal and plant derived MPO textiles using chemical
staining techniques. These stains could separate
animal tissue from wood, but could not distinguish
collagen from keratin.
Vibrational spectroscopy using infrared light has
proven more useful in the identification of keratinous
tissues and requires little or no preparation of the
specimen. Edwards et al. (1998) used Fourier trans-
form Raman spectroscopy (FT-Raman) to distinguish
between modern specimens of five different keratinous
tissues: bovine hoof and horn, antelope horn, marine
turtle shell, and human finger nail, and compared
these with commercial bovine keratin powder. The
predominance of beta-keratin in the tortoiseshell
made it readily distinguishable from the mammalian
tissues but the differences between the mammalian
tissue spectra were altogether more subtle and a
greater number of samples might have shown more
variation, reducing the apparent differences between
these tissues. Espinoza et al. (2007) used diffuse reflec-
tance infrared Fourier transform spectroscopy
(DRIFTS) to analyze bovid and sea turtle keratinous
tissues and differentiate them from casein, a protein-
based plastic historically used in imitation of tortoise-
shell. They acquired spectra from 35 bovid horn or
hoof specimens representing 24 different species and
24 sea turtle shells or mandibular beaks from four
different species. Although DRIFTS has a lower resol-
ution than FTIR, using discriminant analysis it was
possible to separate the horn and turtle shell keratin
with a performance index of 95.7% and all 59 of the
samples were correctly assigned.
Archaeological materials present unique problems
for vibrational spectroscopy. Deterioration of the
organic components produces spreading of spectral
features. At the same time there may be loss of the
inorganic component and increased noise from
mineral and organic contamination from the burial
environment. Together these factors combine to
degrade the spectra. An extreme example of this was
a poorly preserved cattle claw from waterlogged
deposits at York from which FT-Raman failed to
gain recognizable keratin spectra. With well-preserved
finds it may be possible to separate alpha-keratin from
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 13
beta-keratin but only through visual examination of
the structural evidence can the identification of the
tissue be refined. An example of this combined
approach is the use of attenuated total reflectance
infrared spectroscopy (ATR) and visual identification
criteria to identify a find from the 1718 ship Queen
Anne’s Revenge (Welsh et al., 2012). Originally ident-
ified as leather, ATR produced a spectrum that,
when compared to a range of known samples, strongly
resembled known horn. These results only confirmed
the object to be a keratinous material but linear corru-
gations observed upon cleaning supported the identifi-
cation as horn.
None of these techniques can take the identification
of keratinous tissues further than visual identification
techniques. aDNA can provide an answer although,
depending on the context, this may involve a curato-
rially unacceptable level of destructive sampling. The
DNA primarily derives from the mitochondria that
power the high levels of keratin production in the epi-
thelial cells, which ultimately leads to the death of the
cells. When the cells die, the DNA is plasticized but
remains entrapped within the keratin from where it is
very simple to decontaminate (Gilbert et al., 2006).
Under conditions in which the keratin is well pre-
served, it is a valuable source of DNA (e.g. Gilbert
et al., 2007; Clack et al., 2012), and has been used in
the reconstruction of the first ancient human genome
(Rasmussen et al., 2010). Biodeterioration of the
keratin, however, destroys the protection it affords
aDNA, but in such circumstances analysis of surviving
keratin protein remnants might still provide an identi-
fication. Here we present the results of a pilot study of
keratin peptide analysis applied to the problem of dif-
ferentiating keratinous tissues when diagnostic mor-
phological features cannot be seen.
Species identification of keratins using peptide
mass fingerprinting
Species identification by mass spectrometry is based
on the fact that proteins have variable amino acid
sequences across taxa, that these amino acids have
different chemical compositions and, therefore, differ-
ent masses. The more two species diverge in evolution-
ary terms, the more sequence variations are to be
expected for a given protein. Enzymatically digested
proteins produce a mixture of peptides (short strings
of amino acids) that are identified by their mass-to-
charge ratio (m/z) using matrix-assisted laser deso-
rption ionization-time of flight-mass spectrometry
(MALDI-ToF-MS). Due to the conserved nature of
keratins, most of the cleaved peptides will have the
same string of amino acids and therefore the same
m/z and same peak position. However, if there is
one difference in the original protein sequence
between two species, the peptide cleaved from this
region will have a different m/z in each of the
species, and hence peak positions will differ, resulting
in a diagnostic peptide marker.
The profile or peptide mass fingerprint (PMF)
obtained (Fig. 20) is representative of the analyzed
species (Buckley et al., 2009, 2010; Collins et al.,
2010; van Doorn et al., 2011). An unknown sample
can then be identified by matching its PMF against
the PMF of reference samples, usually using several
diagnostic peptide markers to validate the identifi-
cation. PMF has been widely used on collagen from
bone and soft tissues and specimens can generally be
identified at the genus level but recent studies have
shown that some markers were diagnostic to the
species level in certain marine mammals (Kirby et al.,
2013). PMFs of keratin in animal fibers have also
been successfully used to identify ancient textiles and
cloths (Solazzo et al., 2011; Hollemeyer et al., 2012).
Prior to digestion, cysteine residues of the extracted
keratins are chemically reduced to break the disulfide
bridges and alkylated to prevent their reformation.
The enzyme trypsin is used to digest the alpha-keratin
proteins as it cuts specifically after the basic amino
acids arginine (R) and lysine (K), in which alpha-kera-
tins are particularly rich, producing peptides with mass
ranges suitable for PMF (typically 800–4000 Da)
(Fig. 20). Trypsin is less efficient on the matrix proteins
Figure 20 Schematic representation of the preparation and
analytical steps for the identification of proteins by mass
spectrometry.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 014
and therefore the alpha-keratin proteins are expressed
to a greater degree in the PMF and peptide markers
belonging to the most abundant alpha-keratins can
provide reliable identification up to the genus level
(Solazzo et al., 2013b). The wool (or hair), horn, and
hoof of an animal produce the same keratin sequences
but, due to variations in protein expression, differences
can be identified between some species markers in ‘soft’
and ‘hard’ tissues (Solazzo et al., 2013b). Trypsin is less
well adapted to beta-keratin digestion as these keratins
contain less of the basic residues. However, the
proteolytic action of trypsin on beta-keratins is suffi-
cient to identify major markers and when working
with unknown keratinous materials a methodology
that works indiscriminately on alpha- and beta-keratin
is necessary.
Experimental
To explore the potential of PMF for identifying archae-
ological horn and other keratinous hard tissue remains,
samples were collected from five archaeological finds in
various states of preservation, previously identified on
Table 1 Archaeological keratinous materials sampled for PMF
Origin/site code and
finds number (sf) Object Material Condition
Sample size
taken for
analysis
York Archaeological Trust
1977.7 sf 16022 Offcut Horn (a) Opaque and pale fawn/grey with surviving
corrugated lamellar structure, (b) Gelatinized –
brittle, brown/black resinous appearance with no
surviving structure (Fig. 4)
(a) 13 mg;
(b) 5 mg;
1876.13 sf2930 Offcut Horn Opaque and pale fawn/gray with surviving corrugated
lamellar structure
7 mg
1981.7 sf 11937 Knife handle Horn Iron stained MPO remains with surviving corrugated
lamellar structure (Fig. 5)
5 mg
1983.32 sf 1349 Comb
making
debris
Tortoiseshell Brown, opaque and delaminating into fine layers 10 mg
Worcester archaeology
WCM100879 sf 60 Comb
fragment
Tortoiseshell Surface layers becoming granular and detached but
natural pigmentation and areas of translucency
surviving in the core
5 mg
Figure 21 Zoomed area (m/z 1800–2800 Da) of the PMF of modern samples of sheep horn, cow horn and horse hoof with
markers indicated as S (sheep), cow (C) and horse (H).
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 15
their visible structures by O’Connor (Table 1 and
Figs. 4, 5, and 19B). These were compared with the
PMFs gained from modern cow horn, sheep horn,
and horse hoof, and well-preserved historic tortoise-
shell of hawksbill and green turtle, from the teaching
collection of the University of Bradford. Sampling of
the finds was kept to a minimum, between 5 and
13 mg, and done as discreetly as possible to reduce
visual impact. The samples of comparandum speci-
mens were of a similar size, between 9 and 15 mg.
Details of the sample preparation and analytical pro-
cedure are given in the Appendix.
Table 2 Main markers for differentiating sheep, cow, and horse. [M+H]+ calculated with carboxymethylation of cysteine
(+58 Da on C)
[M+H]+ Peptide sequence
Sheep
(Ovis aries)
Cow
(Bos taurus)
Horse
(Equus caballus)
952.50 LQFFQNR S
968.49 LQFYQNR S C H
1041.49 WQFYQNR H
1109.53 DVEEWYIR S C
1896.86 DVEEWFTTQTEELNR H
1834.98 TVNALEVELQAQHNLR S C
1848.99 TVNALEIELQAQHNLR H
2063.03/2191.12 SDLEANSEALIQEIDFLR/KSDLEANSEALIQEIDFLR S C
2075.06/2203.16 SDLEANVEALIQEIDFLR/KSDLEANVEALIQEIDFLR C
2086.04 LEAAVSQAEQQGEVALTDAR H
2114.07 LEVAVSQAEQQGEVALTDAR H
2144.97 SQQQEPLVCPNYQSYFR S C
2234.91 CQQQEPEVCPNYQSYFR H
2563.34 YSSQLSQVQGLITNVESQLAEIR H
2577.30 YSCQLAQVQGLIGNVESQLAEIR C
2665.35 YSCQLSQVQSLIVNVESQLAEIR S
2154.07 GDLEANAHSLVEEVCVLGLK C
In bold, markers indicated on Fig. 21.
Figure 22 Zoomed area (m/z 1800–2800 Da) of the PMF of the archaeological samples 1977.7 sf16022 a (flake) and b
(gelatinized), 1976.13 sf2930 and 1981.7 sf11937 with cow markers indicated as C.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 016
Results
Horn and hoof: alpha-keratins
While it is preferable to identify a combination of
markers for reliable identification, some unique
markers have been found to be diagnostic to the
genus level (Solazzo et al., 2013b) and therefore can
be used to discriminate the most common taxa.
Fig. 21 shows the main markers used for species identi-
fication in horse, cow, and sheep gained from the com-
parandum specimens and Table 2 gives the peptide
sequences for these peaks. A complete list of markers
can be found in Solazzo et al. (2013b) where sheep
horn, cow horn, and horse hoof were compared to
their wool or hair equivalents.
The same diagnostic area is shown in Fig. 22 for the
archaeological samples. All these samples match cattle
horn, confirming the visual identifications. All but one
sample displays three cow diagnostic peaks at m/z
2075, 2154, and 2577. In 1976.13 sf2930, peak m/z
2154 is missing but there is a peak at m/z 2203,
which is also present in 1981.7 sf1937.
Turtle: beta-keratins
As limited sequence information was available for
green and hawksbill turtles, nanoLC-ESI-MS-MS
(liquid chromatographic separation of peptides and
fragmentation of the entities in their amino acid
sequences) data were obtained and peaks from their
respective PMFs matched to a sequence (when poss-
ible) by comparison with available sequences from
other turtle species (Tables 3 and 4). The green turtle
(Chelonia mydas) and the hawksbill turtle
(Eretmochelys imbricate) belong to the Cheloniidae
family of marine turtles but to two different genera.
Table 3 Proteins identified in green turtle by nanoLC-ESI-MS-MS, for which more than two peptides were matched to a score>
30 (some peptides are present in more than one proteins)*
Accession
number† Mr
Error
(ppm) Score Sequence
Reported
modifications
with error
tolerant
Error tolerant
sequence
Peaks
present
on PMF
[M+H]+
gi/215541341 905.3735 −1.82 33 T.CNEPCVR.Q E3→ V T.CNVPCVR.Q
1133.494 6.56 62 G.LCGSGVSCHR.Y
1185.536 2.84 55 R.QCPDSEVVIR.P Gln-> pyro-Glu
(N-term Q)
1186.52
1190.4808 −0.82 60 G.GLCGSGVSCHR.Y
1202.5601 0.36 69 R.QCPDSEVVIR.P 1203.57
1247.5023 1.15 62 Y.GGLCGSGVSCHR.Y
1399.5748 1.56 51 F.SSLCYPECGVAR.P
1402.6630 3.6 46 Y.GGLYGLGGYGGYGGR.Y
1467.5871 4.13 57 Y.GYGGLCGSGVSCHR.Y
1560.6549 10.4 90 R.PSPVTGTCNEPCVR.Q T7→ S R.PSPVTGSCNEPCVR.Q 1561.7
1617.6803 3.56 61 M.TFSSLCYPECGVAR.P T1→A M.AFSSLCYPECGVAR.P
1647.6909 3.79 65 M.TFSSLCYPECGVAR.P
1659.6909 −4.78 74 M.TFSSLCYPECGVAR.P Acetyl
(Nterm);
T1→A
M.AFSSLCYPECGVAR.P 1660.71
1687.6719 −10.18 48 G.GYGYGGLCGSGVSCHR.Y
1744.6934 −8.79 46 L.GGYGYGGLCGSGVSCHR.Y
1781.7462 6.94 102 Y.LGGYGYGGLCGSGVSCHR.Y Y6→ S Y.LGGYGSGGLCGSGVSCH
gi|215541525 1524.6086 6.13 72 Y.GGYGGLCGSGVSCHR.Y
1530.6620 9.37 65 R.PSPVTGSSNEPCVR.Q Carboxy E10 1531.7
gi|215541337 713.3497 14.4 40 G.YGGYGR.R G3→ V G.YGVYGR.R
728.3242 2.66 30 W.GYGGYGR.R
928.4403 −1.92 36 L.YGGLYGLGR.L L4→ S L.YGGSYGLGR.L
954.4923 −6.98 65 L.YGGLYGLGR.L 955.51
1651.799 −6.06 50 G.GYGYGGLYGGLYGLGR.L G3→ S G.GYSYGGLYGGLYGLGR.L 1652.81
gi|215541529 1199.5492 −10.1 60 R.QCPDSEVIIR.P Gln→ pyro-Glu
(N-term Q)
1216.5758 11.3 55 R.QCPDSEVIIR.P 1217.6
1546.6392 9.62 69 R.PSPVSGSCNEPCVR.Q 1547.66
*Are shown: accession number from NCBI (National Center for Biotechnology Information, USA), molecular weight Mr, error in ppm of
the identified peptides, their score (Mascot), sequence, and modifications and adjusted sequence obtained after running error tolerant.
†All proteins identified are ‘beta-keratin-like protein [Pseudemys nelsoni]’ from the Florida redbelly turtle. Finally peaks identified on
PMFs are indicated and in bold are markers found in both green and hawksbill turtles.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 17
Ta
b
le
4
P
ro
te
in
s
id
en
tif
ie
d
in
ha
w
ks
b
ill
tu
rt
le
b
y
na
no
-L
C
-E
S
I-
M
S
-M
S
.
A
cc
es
si
o
n
nu
m
b
er
†
M
r
E
rr
o
r
(p
p
m
)
S
co
re
S
eq
ue
nc
e
R
ep
o
rt
ed
m
o
d
ifi
ca
tio
ns
w
ith
er
ro
r
to
le
ra
nt
E
rr
o
r
to
le
ra
nt
se
q
ue
nc
e
P
ea
ks
p
re
se
nt
o
n
P
M
F
[M
+
H
]+
gi
|2
15
54
15
25
11
90
.4
80
8
−
1.
88
42
G
.G
LC
G
SG
VS
C
H
R
.Y
12
47
.5
02
3
−
2.
77
73
Y.
G
G
LC
G
SG
VS
C
H
R
.Y
14
67
.5
87
1
−
2.
41
71
G
.G
YG
G
LC
G
SG
VS
C
H
R
.Y
15
24
.6
08
6
8.
40
43
Y.
G
G
YG
G
LC
G
SG
VS
C
H
R
.Y
15
30
.6
62
0
4.
37
63
R
.P
SP
VT
G
SS
N
EP
C
VR
.Q
C
ar
bo
xy
E1
0
15
31
.7
3
16
47
.6
90
9
1.
77
58
M
.T
FS
SL
C
YP
EC
G
VA
R
.P
16
70
.7
10
8
−
3.
63
73
G
.G
YG
G
YG
G
LC
G
SG
VS
C
H
R
.Y
C
9
→
S
G
.G
YG
G
YG
G
LS
G
SG
VS
C
H
R
.Y
17
44
.6
93
4
3.
62
71
G
.G
YG
G
YG
G
LC
G
SG
VS
C
H
R
.Y
18
01
.7
14
8
3.
50
53
L.
G
G
YG
G
YG
G
LC
G
SG
VS
C
H
R
.Y
19
14
.7
98
9
−
2.
24
87
G
.L
G
G
YG
G
YG
G
LC
G
SG
VS
C
H
R
.Y
28
00
.1
71
9
−
1.
61
57
G
.S
G
G
H
YG
G
LS
G
LG
G
YG
G
YG
G
LC
G
SG
VS
C
H
R
.Y
C
ar
ba
m
yl
(N
-te
rm
);
S9
→
G
G
.S
G
G
H
YG
G
LG
G
L
G
G
YG
G
YG
G
LC
G
S
G
VS
C
H
R
.Y
28
01
.3
3
gi
|2
15
54
13
41
12
02
.5
60
1
8.
99
75
R
.Q
C
PD
SE
VV
IR
.P
15
79
.7
42
0
2.
80
71
L.
YG
G
LY
G
LG
G
YG
G
YG
G
R
.Y
G
11
→
A
L.
YG
G
LY
G
LG
G
YA
G
YG
G
R
.Y
15
80
.7
1
gi
|2
15
54
13
39
12
33
.5
65
9
−
0.
91
79
R
.Q
C
Q
D
SE
VV
IR
.P
gi
|2
15
54
13
37
72
8.
32
42
−
1.
95
45
W
.G
YG
G
YG
R
.R
79
1.
42
90
0.
21
47
Y.
G
G
LY
G
LG
R
.Y
95
4.
49
23
0.
19
65
L.
YG
G
LY
G
LG
R
.L
95
5.
51
99
7.
49
81
10
.8
64
L.
YG
G
LY
G
LG
R
.L
C
ar
ba
m
yl
(N
-te
rm
)
A
ce
ty
l
(N
te
rm
);
O
xi
da
tio
n
Y1
10
13
.5
3
10
12
.4
97
8
2.
23
50
L.
YG
G
LY
G
LG
R
.L
gi
|2
15
54
15
29
10
68
.5
35
3
3.
17
49
F.
G
LG
G
LY
G
YG
G
H
.Y
H
11
→
R
F.
G
LG
G
LY
G
YG
G
R
.Y
11
89
.5
51
6
−
1.
45
72
R
.Y
G
YG
G
LS
G
YG
G
R
.Y
Y3
→
F
R
.Y
G
FG
G
LS
G
YG
G
R
.Y
11
90
.6
1
12
15
.6
03
7
−
3.
8
52
S.
FG
LG
G
LY
G
YG
G
H
.Y
H
12
→
R
S.
FG
LG
G
LY
G
YG
G
R
.Y
12
16
.5
75
8
−
0.
06
59
R
.Q
C
PD
SE
VI
IR
.P
12
35
.5
57
1
12
.1
86
R
.Y
G
YG
G
LS
G
YG
G
R
.Y
G
8
→
S
R
.Y
G
YG
G
LS
SY
G
G
12
36
.6
12
78
.5
62
9
−
4.
66
56
R
.Y
G
YG
G
LS
G
YG
G
R
.Y
C
ar
ba
m
yl
(N
-te
rm
);
G
8
→
S
R
.Y
G
YG
G
LS
SY
G
G
R
.Y
13
02
.6
35
7
−
10
.2
1
42
G
.S
FG
LG
G
LY
G
YG
G
H
.Y
H
13
→
R
G
.S
FG
LG
G
LY
G
YG
G
R
.Y
13
59
.6
57
2
−
4.
70
68
G
.G
SF
G
LG
G
LY
G
YG
G
H
.Y
H
14
→
R
G
.G
SF
G
LG
G
LY
G
YG
G
R
.Y
14
16
.6
78
7
−
2.
94
82
Y.
G
G
SF
G
LG
G
LY
G
YG
G
H
.Y
H
15
→
R
Y.
G
G
SF
G
LG
G
LY
G
YG
G
R
.Y
15
32
.6
92
9
6.
08
74
R
.P
SP
VS
G
SC
N
EP
C
VR
.Q
C
8
→
F
R
.P
SP
VS
G
SF
N
EP
C
VR
.Q
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 018
It is therefore expected to see a substantial difference in
the peptide profiles between genera as is seen between
alpha-keratin-made tissues. Fig. 23 indicates the
unique markers (unique, that is, in this two-taxon com-
parison) identified for green turtle and Fig. 24 those
for hawksbill turtle (see Discussion).
The profiles of the archaeological tortoiseshell
samples are included in Fig. 24 as they matched the
hawksbill turtle better than the green turtle. In particu-
lar, sample WCM100879 sf60 has all m/z 1190, 1236,
1580, and 2200 markers but showed none of the green
turtle markers. Sample 1983.32 sf1349 showed more
signs of degradation with less peaks and only m/z
2200 as a major peak and m/z 1630 as a minor
peak, still matching Hawksbill turtle best of the
species available as comparanda.
Discussion
Strategies of identification by mass
spectrometry
Cow, sheep, and horse are well-sequenced species, for
which alpha-keratin sequences are now available
through public databases. Markers to separate all
three species have been described elsewhere (Solazzo
et al., 2013b) and are summarized in Table 2, but we
focused here on the most diagnostic region of the
peptide profiles, between m/z 1800 and 2800 Da.
One marker in particular is diagnostic of sheep at
m/z 2665, its equivalent in goat is at m/z 2692. The
equivalent marker in horse is found at m/z 2563 and
in cow at m/z 2577. Cow has two more reliable
markers at m/z 2154 and at m/z 2075/2203 (m/z
2203 is one residue longer than m/z 2075), and horse
three at m/z 2086, 2114 and 2235 (Table 2).
For turtle beta-keratins, available sequences are
limited in the public database to incomplete sets of
sequences from the Florida redbelly turtle (18
sequences) and green turtle (two sequences). MS-MS
data searched against all animals logically matched
beta-keratins from Florida redbelly turtle (Tables 3
and 4). In addition, a few alpha-keratin-like proteins
were observed in hawksbill turtle (data not shown)
but only beta-keratin peptides were identified on
PMF (which allows only a snapshot of the total
protein content), most likely due to the alpha-keratins
being in the minority.
To increase the chances of matching peptides, the
MS-MS dataset was searched against ‘error tolerant’,
a search parameter which uses all possible post-trans-
lational modifications as well as possible substitutions
of amino acids to match sequences in the proteins
identified (here the different beta-keratin-like protein
from the Florida redbelly turtle). A high number of
peptides were successfully identified through substi-
tutions of amino acids; however, these identifications
have to be taken with caution until more sequencesgi
|2
15
54
13
31
94
5.
39
02
3.
89
36
G
.Y
G
G
LC
G
SG
V.
S
S7
→
Y
G
.Y
G
G
LC
G
YG
V.
S
15
60
.6
54
9
4.
38
70
R
.P
SP
VT
G
TC
N
EP
C
VR
.Q
T7
→
S
R
.P
SP
VT
G
SC
N
EP
C
VR
.Q
15
61
.6
1
16
17
.6
80
3
−
1.
05
74
M
.T
FS
SL
C
YP
EC
G
VA
R
.P
S4
→
G
M
.T
FS
G
LC
YP
EC
G
VA
R
.P
16
59
.6
90
9
−
8.
09
48
M
.T
FS
SL
C
YP
EC
G
VA
R
.P
A
ce
ty
l(
N
te
rm
);
S4
→
G
M
.T
FS
G
LC
YP
EC
G
VA
R
.P
16
60
.7
6
21
91
.9
05
2
0.
71
51
L.
G
G
YL
G
G
YG
YG
G
LC
G
SG
VS
C
H
R
.Y
L.
G
G
YL
G
G
YG
YG
G
G
LC
G
SG
VS
C
H
R
.Y
24
19
.3
22
−
4.
78
51
Y.
G
G
LG
G
YL
G
G
YG
YG
G
LC
G
SG
VS
C
H
R
.Y
YG
G
G
LC
G
SG
VS
C
H
R
.Y
26
39
.1
17
0
−
8.
35
74
Y.
G
G
LG
G
YL
G
G
YG
YG
G
LC
G
SG
VS
C
H
R
.Y
+
57
(G
)→
G
16
Y.
G
YG
G
LG
G
YL
G
G
YG
YG
G
G
LC
G
SG
VS
C
H
R
.Y
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 19
are made available. Fortunately, the recent sequencing
of the soft-shell turtle Pelodiscus sinensis, the green
turtle, and the western painted turtle Chrysemys picta
bellii (Li et al., 2013; Shaffer et al., 2013; Wang
et al., 2013) should increase the number of keratin
sequences available. Using error tolerant, however,
many peptides could be related to beta-keratin
sequences, and major peaks observed on the PMFs
of both species of turtles could also be associated to
a sequence. On the PMFs, four peaks were found in
common between green and hawksbill turtles but
this number increased to 14 after LC-MS-MS analysis
Figure 23 PMF m/z 800–3000 Da of the green turtle with markers indicated as GT.
Figure 24 PMFm/z 800–3000 Da of the hawksbill turtle and archaeological samples WCM100879 sf60 and 1983.32 sf1349 with
markers indicated as HT.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 020
(in bold, Tables 3 and 4). Two such peptides, common
to both species in LC-MS-MS runs, were observed on
the green turtle PMF but not the hawksbill’s. Using
these complementary methods, diagnostic markers
could be established for both turtles: m/z 1547 and
m/z 1652 for green turtle, and m/z 1190, mz 1236,
and m/z 1580, for hawksbill turtle. Three major
peaks were not identified to a sequence but were
found in both LC-MS-MS and PMFs: m/z 2088 for
green turtle and m/z 1630 and m/z 2200 for hawksbill
turtle (Figs. 23 and 24).
Degradation of hard keratinous tissues
Identification to cow is possible thanks to several
markers; however, loss of some peptides and variable
proportions of these markers show variable degra-
dation of the archaeological samples (Table 5). In
modern cattle horn, m/z 1835, 2154, and 2577 are the
most intense peaks with much lower m/z 2075 and
2203. In 16022a the proportion is maintained except
for m/z 1835 that loses a lot of intensity and in the
gel all peaks become very low compared to m/z 2154.
In 11937, m/z 2154 is also the most intense peak fol-
lowed by m/z 2577, while in 2930 m/z 2154 is absent
but the intensity of m/z 2075 and 2203 are higher.
Generally, during degradation there is a loss of some
of themost intense peaks (m/z 1835 and 2577) and con-
sequently a relative increase of the low intensity peaks
at m/z 2075 and 2203 with m/z 2075 consistently
more intense than m/z 2203.
Conclusions
On the vast majority of archaeological sites, the har-
vesting and working of keratinous hard tissues is evi-
denced largely by secondary indicators, such as
caches of boney horn cores. Where these materials
do survive it is generally in circumstances where the
macro or micro environment has resulted in the inhi-
bition of biodeterioration. As intimate contact with
corroding metals is the most frequent preservation
mechanism (MPO remains) for these materials in tem-
perate zone, free-draining sites, the evidence for their
exploitation prior to the Bronze Age is particularly
poor. Their ‘invisibility’ in the archaeological record
is compounded by a lack of familiarity with their struc-
tural features and working properties. Today horn and
hoof have largely been superseded as raw materials by
modern metal alloys and plastics whilst conservation
concerns have made the use of baleen and tortoiseshell
unavailable and unacceptable.
This paper illustrates how low power microscopy in
combinationwith high quality, directional lighting pro-
vides a powerful tool enabling the differentiation and
identification of all these keratinous tissues on the
basis of structural criteria and characteristic patterns
of deterioration without the need for destructive
sampling or surface preparation. Horn, hoof, baleen,
and tortoiseshell can all be differentiated from each
other but identification to species is not always feasible.
While cattle horn can be separated from that of sheep/
goat horn with some confidence, even when quite
decayed, tortoiseshell cannot normally be attributed
to the hawksbill or green turtle by these techniques
unless the material has retained its translucency and a
sufficiently large area has survived to allow the charac-
teristics of the mottled patterning to be observed.
The results presented here demonstrate that peptide
mass fingerprinting has the capability to identify
degraded keratinous remains to species even when
preservation is largely due to mineral replacement or
when the fibrous protein has been reduced to a gel
by diagenetic changes. This analytical technique does
require destructive sampling but the small sample
size, typically no bigger than half a grain of rice,
should make it curatorially acceptable in the majority
of cases where species identification is critical. When
proteins break down, the chances of detecting species
markers decrease. Identification therefore often
depends more on the state of the material than on
the size of the sample; there is no analytical gain in
increasing the sample size. If PMF fails, an alternative
method would be to use nano-LC-MS-MS that not
only allows the identification of a higher number of
peptides through chromatographic separation and
concentration of the samples; it also allows the
amino acid sequence of peptides to be ‘read’, guaran-
teeing a correct identification of peptides.
What PMF currently cannot do is distinguish
between different keratinous tissues of the same
species, such as cattle horn from cattle hoof. Used in
conjunction with visual identification criteria,
however, PMF has the potential to obtain identifi-
cations from even highly worked and degraded
materials with a level of confidence not previously
available.
Acknowledgements
We thank Cluny Chapman, the Hawley Collection, the
Horniman Museum, Hull Museums, Leeds Museum
Table 5 Peak intensity ratios as compared tom/z 2577 (= 1)
1835/
2577
2075/
2577
2154/
2577
2203/
2577 2577
Modern cow 0.65 0.13 0.78 0.07 1
1981.7
sf11937
0.16 0.54 4.76 0.21 1
1976.13
sf2930
0.49 0.41 0 0.29 1
1977.7
sf16022a
0.17 0.14 0.64 0 1
1977.7
sf16022b
0.54 0.83 7.34 0.67 1
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 21
and Galleries, and the York Archaeological Trust for
access to the specimens used in this paper. Other
than Fig. 1, for which we thank the Canadian
Conservation Institute, all photographs are by
O’Connor and all analytical graphics are by Solazzo.
The research by O’Connor to evaluate and develop
techniques for the identification of keratinous hard
tissues was undertaken as part of her post-doctoral fel-
lowship, Cultural Objects Worked in Skeletal Hard
Tissues, funded by the Arts and Humanities
Research Council and the Engineering and Physical
Sciences Research Council, through their Science
and Heritage Programme. Solazzo’s work was sup-
ported by a Marie Curie International Outgoing
Fellowship (FP7-PEOPLE-IOF-GA-2009-236425,
THREADs) jointly between University of York,
UK, and AgResearch, NZ (2009–2012), and a
Smithsonian’s Museum Conservation Institute (MCI)
post-doctoral grant. Analyses were carried out at
The Centre of Excellence in Mass Spectrometry,
University of York, supported by Science City York
and Yorkshire Forward, using funds from the
Northern Way Initiative. We particularly thank
Dr. David Ashford for carrying out analyses on the
maXis instrument.
Appendix
Chemicals
Urea, tris(2-carboxyethyl)phosphine (TCEP), iodoa-
cetic acid (IAA), ammonium bicarbonate (AB),
formic acid, and trifluoroacetic acid (TFA) were pro-
vided by Sigma-Aldrich (St Louis, Missouri, USA),
acetonitrile (ACN) and acetone by Fisher Scientific
(Loughborough, Leicestershire, UK), ethanol
(EtOH) by AnalaR NORMAPUR (Lutterworth,
Leicestershire, UK), trypsin by Promega (Madison,
Wisconsin, USA), and alpha-cyano-4-hydroxycin-
namic acid (CHCA) by Bruker Daltonics (Bremen,
Germany). The 3500 MWCO Slide A Lyzer® Mini
Dialysis units were obtained from Thermo Scientific
(Rockford, Illinois, USA).
Sample preparation
Samples were ground to a fine powder in a mortar with
the help of liquid nitrogen and allowed to dry over-
night, then solubilized by shaking overnight in a sol-
ution of 8 M urea, 50 mM Tris, and 50 mM TCEP
at pH 8.3. An aliquot of the supernatant was alkylated
with 150 mM IAA and vortexed for four hours in the
dark. This was followed by 24 hours dialysis with
100 mM AB on 3500 MWCO dialysis units (two
changes). The dialyzed aliquot was then digested over-
night with 0.5 μg of trypsin at 37°C. Samples were then
dried down and re-solubilized in 10 μl of 0.1% TFA.
Peptide mass fingerprinting by MALDI-TOF-MS
A matrix solution was prepared by diluting 0.1 mg of
CHCA in 97/3 (acetone/0.1% TFA) and 1 μl was
applied onto an AnchorChipTM target (Bruker) and
allowed to dry. A 1 μl aliquot of analytical solution
was applied and removed after one minute followed
by 1 μl of washing buffer (0.1% TFA). The residual
droplet was removed and 1 μl of recrystallization sol-
ution (0.1 mg of CHCA in 6/3/1 (EtOH/acetone/
0.1% TFA)) applied. The plate was loaded in a
UltraflexTM III mass spectrometer (Bruker), and ana-
lyses were carried out in positive reflector mode using
a Nd:YAG laser operating at 355 nm. Spectra were
acquired using flexControl 3.0 (Bruker) on a mass
range of m/z 800–4000 Da with an accumulation of
500 shots on the standards and 1000 shots on the
samples. The calibration standard (Bruker) was pre-
pared according to the manufacturer’s instructions
for instrument calibration and consisted of angiotensin
I, ACTH clip(1–17), ACTH clip(18–39), and ACTH
clip(7–38) peptides.
Peptide analysis by nano-LC-ESI-MS-MS
Analyses of the reference samples were performed on a
UHR Q-TOF. HPLC was performed using a
nanoAcquity UPLC system (Waters) equipped with a
nanoAcquity Symmetry C18, 5 μm trap (180 μm ×
20 mm; Waters) and a nanoAcquity BEH130 1.7 μm
C18 capillary column (75 m × 250 mm; Waters). The
trap wash solvent was 0.1% aqueous formic acid and
the trapping flow rate was 10 μl/minute. The trap
was washed for five minutes after sample loading
before switching flow to the capillary column. The sep-
aration used a gradient elution of two solvents: solvent
A (0.1% formic acid) and solvent B (acetonitrile con-
taining 0.1% formic acid). The flow rate for the capil-
lary column was 300 nl/minute, column temperature
was 60°C and the gradient profile was as follows:
initial conditions 5% solvent B (2 minutes), followed
by a linear gradient to 35% solvent B over 20
minutes and then a wash with 95% solvent B for 2.5
minutes. The column was returned to initial conditions
and re-equilibrated for 25 minutes before subsequent
injections. The nano-LC system was interfaced to a
maXis UHR Q-TOF (ultrahigh-resolution quadru-
pole-time-of-flight) mass spectrometer (Bruker) with
a nano-electrospray source fitted with a steel emitter
needle (180 μm outside diameter × 30 μm inside diam-
eter; Thermo). Positive-ion MS and CID (collision-
induced dissociation) – MS-MS spectra were acquired
using AutoMSMS mode. Instrument control and data
acquisition were performed using Compass 1.3 SP1
software (micrOTOF control and Hystar, Bruker).
Instrument settings were: ion spray voltage, 1400 V;
dry gas, 4 l/minute; dry gas temperature, 160°C; and
ion acquisition range m/z, 50–2200. AutoMSMS
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 022
settings were for MS 0.5 seconds (acquisition of survey
spectrum) and for MS-MS (CID with N2 as collision
gas) ion acquisition range m/z 350–1400; 0.1 second
acquisition for precursor intensities above 100,000
counts; for signals of lower intensities down to 1000
counts acquisition time increased linearly to 1.5
seconds; the collision energy and isolation width set-
tings were automatically calculated using the
AutoMSMS fragmentation table; three precursor
ions; absolute threshold 1000 counts; preferred
charge states 2–4; singly charged ions excluded. Two
MS-MS spectra were acquired for each precursor and
previously selected target ions were excluded for 60
seconds. Spectra were calibrated using a lock mass
signal (m/z 1221.99064) prior to compound detection
and peak list creation using DataAnalysis (Bruker
Daltonics).
The peak list files obtained were submitted for data-
base searching to a locally running copy of Mascot
(Matrix Science Ltd., version 2.3.02). Mascot
(Matrix Science) was used to search for matches
against the database of publicly available sequences
NCBI (National Center for Biotechnology
Information, US). Searches were carried out using
semi-trypsin as enzyme (horn and hoof ) or no
enzyme (turtles), two missed cleavages, peptide mass
tolerance (MS) of 20 ppm, and fragment mass toler-
ance (MS-MS) of 0.1 Da, carboxymethylation as a
fixed modification (alkylation using IAA adds a
+58 Da to the mass of the cysteine residues) and
acetyl (N-term), carbamyl (N-term), deamidated
(NQ), Gln-> pyro-Glu (N-term Q), Glu-> pyro-Glu
(N-term E), and oxidation (M) as variable modifi-
cations. Horn and hoof were searched against other
mammals while green turtle and hawksbill turtle
were searched against all animals.
References
Aguilar, A. 1986. A Review of OldBasque Whaling and Its Effect on
the Right Whales (Eubalaena glacialis) of the North Atlantic.
Report of the International Whaling Commission, Special
Issue, 10: 191–9.
Alibardi, L., Dalla Valle, L., Nardi, A. & Toni, M. 2009. Evolution
of Hard Proteins in the Sauropsid Integument in Relation to the
Cornification of Skin Derivatives in Amniotes. Journal of
Anatomy, 214: 560–86.
Anheuser, K. & Roumeliotou, M. 2003. Characterisation of
Mineralised Archaeological Textile Fibres Through Chemical
Staining. The Conservator, 27: 23–33.
Armitage, P. 1990. Post-medieval Cattle Horn Cores from the
Greyfriars Site, Chichester, West Sussex, England. Circaea,
7(2): 81–90.
Bartels, M.H. 2005. The Van Lidth de Jeude family and the Waste
from their Privy: Material Culture of a Wealthy Family in
18th-century Tiel, the Netherlands. Northeast Historical
Archaeology, 34: 5–62.
Beattie, O. & Geiger, J. 1987. Frozen in Time: The Fate of the
Franklin Expedition. London: Bloomsbury.
Beck, S.W. 1882. The Drapers’ Dictionary. A Manual of Textile
Fabrics: Their History and Applications. London: The
Warehousemen and Drapers’ Journal Office.
Biek, L. 1963. Archaeology and the Microscope. London:
Butterworth.
Błyskal, B. 2009. Fungi Utilizing Keratinous Substrates.
International Biodeterioration & Biodegradation, 63: 631–53.
Bragulla, H. & Hirschberg, R.M. 2003. Horse Hooves and Bird
Feathers: Two Model Systems for Studying the Structure and
Development of Highly Adapted Integumentary Accessory
Organs—The Role of the Dermo-epidermal Interface for the
Micro-architecture of Complex Epidermal tructures. Journal
of Experimental Zoology Part B: Molecular and
Developmental Evolution, 298B: 140–51.
Brothwell, D. & Gill-Robinson, H. 2002. Taphonomic and Forensic
Aspects of Bog Bodies. In: W.D. Haglund &H.S. Marcella, eds.
Advances in Forensic Taphonomy: Method, Theory, and
Archaeological Perspectives. Boca Raton: CRC Press, pp.
119–33.
Bruce-Mitford, R. & Luscombe, M.R. 1974. The Benty Grange
Helmet. In: R. Bruce-Mitford, ed. Aspects of Anglo-Saxon
Archaeology. Sutton Hoo and Other Discoveries. London:
Victor Gollancz Limited, pp. 223–42.
Buckley, M., Collins, M., Thomas-Oates, J. & Wilson, J.C. 2009.
Species Identification by Analysis of Bone Collagen Using
Matrix-assisted Laser Desorption/Ionisation Time-of-flight
Mass Spectrometry. Rapid Communications in Mass
Spectrometry, 23: 3843–54.
Buckley, M., Whitcher Kansa, S., Howard, S., Campbell, S.,
Thomas-Oates, J. & Collins, M. 2010. Distinguishing Between
Archaeological Sheep and Goat Bones Using a Single
Collagen Peptide. Journal of Archaeological Science, 37: 13–20.
Campbell Pedersen, M. 2004. Gem and Ornamental Materials of
Organic Origin. Oxford: Elsevier Butterworth-Heinemann.
Clack, A.A., MacPhee, R.D.E. & Poinar, H.N. 2012. Mylodon
Darwinii DNA Sequences from Ancient Fecal Hair Shafts.
Annals of Anatomy=Anatomischer Anzeiger: Official Organ of
The Anatomische Gesellschaft, 194(1): 26–30.
Clark, G. 1947. Whales as an Economic Factor in Prehistoric
Europe. Antiquity, 21: 84–104.
Collins, M., Buckley, M., Thomas-Oates, J., Wilson, J. & Van
Doorn, N., 2010. ZooMS: The Collagen Barcode and
Fingerprints. Spectroscopy Europe, 11–3.
Collins, M.J., Nielsen-Marsh, C.M., Hiller, J., Smith, C.I., Roberts,
J.P., Prigodich, R.V., Wess, T.J., Csapò, J., Millard, A.R. &
Turner-Walker, G. 2002. The Survival of Organic Natter in
Bone: A Review. Archaeometry, 44: 383–94.
Credland, A.G. 1981. Crossbow Remains (Part 2). Journal of the
Society of Archer Antiquaries, 24: 9–16.
Dalla Valle, L., Michieli, F., Benato, F., Skobo, T. & Alibardi, L.
2013. Molecular Characterization of Alpha-keratins in
Comparison to Associated Beta-proteins in Soft-shelled and
Hard-shelled Turtles Produced During the Process of
Epidermal Differentiation. Journal of Experimental Zoology
Part B: Molecular and Developmental Evolution, 320(7): 1–14.
Dalla Valle, L., Nardi, A., Toni, M., Emera, D. & Alibardi, L. 2009.
Beta-keratins of Turtle Shell are Glycine-proline-tyrosine Rich
Proteins Similar to Those of Crocodilians and Birds. Journal
of Anatomy, 214: 284–300.
Doppelfeld, O. 1964. Das fränkische Knabengrab unter dem Chor
des Kölner Domes. Germania, 42: 156–88.
Dzierzykray-Rogalski, T. 1986. Natural Mummification in Egypt.
In: A.R. David, ed. Science in Egyptology. Manchester:
Manchester University Press, pp. 101–12.
Edwards, G.M. 1974. The Preservation of Textiles in Archaeological
Contexts. PhD dissertation, University of London Institute of
Archaeology.
Edwards, H.G.M., Hunt, D.E. & Sibley, M.G. 1998. FT-Raman
Spectroscopic Study of Keratotic Materials: Horn, Hoof and
Tortoiseshell. Spectrochimica Acta Part A: Molecular and
Biomolecular Spectroscopy, 54: 745–57.
Espinoza, E.O., Baker, B.W. & Berry, C.A. 2007. The Analysis of Sea
Turtle and Bovid Keratin Artefacts Using Drift Spectroscopy
and Discriminant Analysis. Archaeometry, 49: 685–98.
Florian, M.L.E. 1987. Deterioration of Organic Materials Other
than Wood. In: C. Pearson, ed. Conservation of Marine
Archaeological Objects. London: University of Michigan,
Butterworths, pp. 21–54.
Franck, A., Cocquyt, G., Simoens, P. & De Belie, N. 2006.
Biomechanical Properties of Bovine Claw Horn. Biosystems
Engineering, 93: 459–67.
Fudge, D.S., Szewciw, L.J. & Schwalb, A.N. 2009. Morphology and
Development of Blue Whale Baleen: An Annotated Translation
of Tycho Tullberg’s Classic 1883 Paper. Aquatic Mammals, 35:
226–52.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 23
Gilbert, M.T.P., Djurhuus, D., Melchior, L., Lynnerup, N.,
Worobey, M., Wilson, A.S., Andreasen, C. & Dissing, J.
2007. DNA from Hair and Nail Clarifies the Genetic
Relationship of the 15th Century Qilakitsoq Inuit Mummies.
American Journal of Physical Anthropology, 133(2): 847–53.
Gilbert, M.T.P., Menez, L., Janaway, R.C., Tobin, D.J., Cooper, A.
& Wilson, A.S. 2006. Resistance of Degraded Hair Shafts to
Contaminant DNA. Forensic Science International, 156(2–3):
208–12.
Gilbert, S.F., Loredo, G.A., Brukman, A. & Burke, A.C. 2001.
Morphogenesis of the Turtle Shell: the Development of a
Novel Structure in Tetrapod Evolution. Evolution &
Development, 3: 47–58.
Gillard, R.D., Hardman, S.M., Thomas, R.G. & Watkinson, D.E.
1994. The Mineralization of Fibres in Burial Environments.
Studies in Conservation, 39: 132–40.
Halstead, L.B. 1974. Vertebrate Hard Tissues. London: Wykeham
Publications.
Hanausek, T.F. 1907. The Microscopy of Technical Products.
New York: J. Wiley.
Hashiguchi, K. & Hashimoto, K. 1995. The Mineralization of
Crystalline Inorganic Components in Japanese Serow Horn.
Okajimas Folia Anatomica Japonica, 72: 235–43.
Hashiguchi, K., Hashimoto, K. & Akao, M. 2001. Morphological
Character of Crystalline Components Present in Saiga Horn.
Okajimas Folia Anatomica Japonica, 78: 43–8.
Hett, C. 1980. Unearthed Coins: A Pennyworth of History. The
Journal of the Canadian Conservation Institute, 4: 21–3.
Hieronymus, T.L., Witmer, L.M. & Ridgely, R.C. 2006. Structure of
White Rhinoceros (Ceratotherium simum) Horn Investigated by
X-ray Computed Tomography and Histology with Implications
for Growth and External Form. Journal of Morphology, 267:
1172–6.
Hollemeyer, K., Altmeyer, W., Heinzle, E. & Pitra, C. 2012. Matrix-
assisted Laser Desorption/Ionization Time-of-flight Mass
Spectrometry Combined with Multidimensional Scaling,
Binary Hierarchical Cluster Tree and Selected Diagnostic
Masses Improves Species Identification of Neolithic Keratin
Sequences from Furs of the Tyrolean Iceman Oetzi. Rapid
Communications in Mass Spectrometry, 26: 1735–45.
Janaway, R. 1983. Textile Fibre Characteristics Preserved by Metal
Corrosion: The Potential of S.E.M. Studies. The Conservator, 7:
48–52.
Janaway, R.C. 1989. Corrosion Preserved Textiles: Mechanisms,
Bias and Interpretation. In: R.C. Janaway & B. Scott, eds.
Evidence Preserved In Corrosion Products: New Fields in
Artifact Studies. London: United Kingdom Institute for
Conservation (Occasional Paper 8), pp. 21–9.
Janaway, R.C. & Scott, B. eds. 1983. Evidence Preserved in Corrosion
Products: New Fields in Artifact Studies. London: United
Kingdom Institute for Conservation, Occasional Paper 8.
Jones, M. 2003. For Future Generations: Conservation of a Tudor
Maritime Collection. Portsmouth: Mary Rose Trust Limited.
Keepax, C. 1975. Scanning Electron Microscopy of Wood Replaced
by Iron Corrosion Products. Journal of Archaeological Science,
2: 145–50.
Kirby, D.P., Buckley, M., Promise, E., Trauger, S.A. & Holdcraft,
T.R. 2013. Identification of Collagen-based Materials in
Cultural Heritage. Analyst, in press., 138: 4849–58.
Krzyszkowska, O. 1990. Ivory and Related Materials: An Illustrated
Guide. London: London Institute of Classical Studies (Classical
Handbook 3, Bulletin Supplement 59).
Lauffenburger, J.A. 1993. Baleen in Museum Collections: Its
Sources, Uses, and Identification. Journal of the American
Institute for Conservation, 32: 213–30.
Li, Y.I., Kong, L., Ponting, C.P. & Haerty, W. 2013. Rapid
Evolution of Beta-Keratin Genes Contribute to Phenotypic
Differences That Distinguish Turtles and Birds from Other
Reptiles. Genome Biology and Evolution, 5: 923–33.
MacGregor, A. 1985. Bone, Antler, Ivory and Horn: The Technology of
SkeletalMaterials Since the Roman Period. London: CroomHelm.
MacGregor, A. 1991. Antler, Bone and Horn. In: J. Blair &
N. Ramsay, eds. English Medieval Industries. Craftsmen,
Techniques, Products. London: The Hambledon Press, pp.
356–78.
Marshall, R.C., Orwin, D.F.G. & Gillespie, J.M. 1991. Structure
and Biochemistry of Mammalian Hard Keratin. Electron
Microscopy Reviews, 4: 47–83.
McKittrick, J., Chen, P.Y., Bodde, S.G., Yang, W., Novitskaya, E.E.
& Meyers, M.A. 2012. The Structure, Functions, and
Mechanical Properties of Keratin. JOM Journal of the
Minerals, Metals and Materials Society, 64: 449–68.
McKittrick, J., Chen, P.Y., Tombolato, L., Novitskaya, E.E., Trim,
M.W., Hirata, G.A., Olevsky, E.A., Horstemeyer, M.F. &
Meyers, M.A. 2010. Energy Absorbent Natural Materials and
Bioinspired Design Strategies: A Review. Materials Science
and Engineering C, 30: 331–42.
Moffat, R., Spriggs, J. & O’Connor, S. 2008. The Use of Baleen for
Arms, Armour and Heraldic Crests in Medieval Britain. The
Antiquaries Journal, 88: 207–15.
Mulville, J. 2002. The Role of Cetacea in Prehistoric and Historic
Atlantic Scotland. International Journal of Osteoarchaeology,
12: 34–48.
O’Connor, S. 1987. The Identification of Osseous and Keratinaceous
Materials at York. In: K. Starling & D. Watkinson, eds.
Archaeological Bone, Antler and Ivory. London: United
Kingdom Institute for Conservation, pp. 9–21.
O’Connor, S. 2013. Exotic Materials used in the Construction of
Iron Age Sword Handles from South Cave, UK. In: A.
Choyke & S. O’Connor, eds. From These Bare Bones: Raw
materials and the Study of Worked Osseous Objects.
Proceedings of the Raw Materials session at the 11th ICAZ
Conference, Paris, 2010. Oxford: Oxbow, pp. 188–200.
O’Connor, S. in press. Exotic Materials Used in the Construction of
Iron Age Sword Handles from South Cave, UK. In: A. Choyke
& S. O’Connor, eds. From These Bare Bones: Raw Materials
and the Study of Worked Osseous Materials. Proceedings of
the Raw Materials Session at the 11th ICAZ Conference,
Paris, 2010. Oxford: Oxbow.
O’Connor, S. & Duncan, H. 2002. The Bone, Antler and Ivory
Artefacts. In: I. Roberts, ed. Pontefract Castle Archaeological
Excavations 1982–86. Leeds: West Yorkshire Archaeology
Service, pp. 300–8.
O’Connor, S. & O’Connor, T. In prep. Reconsideration of the
‘Mesolithic Harpoon’ from Westward Ho!. Devon.
Pautard, F.G.E. 1964. Calcification of Keratin. In: A. Rook &
R.H. Champion, eds. Progress in the Biological Sciences in
Relation to Dermatology. London: Cambridge University
Press, pp. 227–40.
Pautard, F.G.E. 1970. The Mineral Phase of Calcified Cartilage,
Bone and Baleen. Calcified Tissue Research, 4: 34–6.
Penniman, T.K. 1952. Pictures of Ivory and other Animal Teeth, Bone
and Antler, Pitt Rivers Museum Occasional Papers on
Technology 5. Oxford: University of Oxford.
Pfeiffer, C.J. 1992. Cellular Structure of Terminal Baleen in Various
Mysticete Species. Aquatic Mammals, 18: 67–73.
Plowman, J.E. 2007. The Proteomics of Keratin Proteins. Journal of
Chromatography B, 849: 181–9.
Pollitt, C.C. 2004. Anatomy and Physiology of the Inner Hoof Wall.
Clinical Techniques in Equine Practice, 3: 3–21.
Rasmussen, M., Li, Y., Lindgreen, S., Pedersen, J.S., Albrechtsen,
A., Moltke, I., Metspalu, M., Metspalu, E., Kivisild, T.,
Gupta, R., Bertalan, M., Nielsen, K., Gilbert, M.T., Wang,
Y., Raghavan, M., Campos, P.F., Kamp, H.M., Wilson, A.S.,
Gledhill, A., Tridico, S., Bunce, M., Lorenzen, E.D.,
Binladen, J., Guo, X., Zhao, J., Zhang, X., Zhang, H., Li, Z.,
Chen, M., Orlando, L., Kristiansen, K., Bak, M., Tommerup,
N., Bendixen, C., Pierre, T.L., Grønnow, B., Meldgaard, M.,
Andreasen, C., Fedorova, S.A., Osipova, L.P., Higham, T.F.,
Ramsey, C.B., Hansen, T.V., Nielsen, F.C., Crawford, M.H.,
Brunak, S., Sicheritz-Pontén, T., Villems, R., Nielsen, R.,
Krogh, A., Wang, J. & Willerslev, E. 2010. Ancient Human
Genome Sequence of an Extinct Palaeo-Eskimo. Nature,
463(7282): 757–62.
Richards, M. 2005. Lighting Equipment. In: J. Gardiner &
M.J. Allen, eds. Before the Mast. Life and Death Aboard the
Mary Rose. Portsmouth: The Mary Rose Trust, pp. 343–8.
Richards, M. & Gardiner, J. 2005. Books and Writing Equipment.
In: J. Gardiner & M.J. Allen, eds. Before the Mast. Life and
Death Aboard the Mary Rose. Portsmouth: The Mary Rose
Trust, pp. 127–33.
Rijkelijkhuizen, M. 2008. Handleiding voor de determinatie van
harde dierlijke materialen. Amsterdam: Amsterdam University
Press.
Rijkelijkhuizen, M. 2009. Whales, Walruses, and Elephants:
Artisans in Ivory, Baleen, and Other Skeletal Materials in
Seventeenth- and Eighteenth-Century Amsterdam.
International Journal of Historical Archaeology, 13: 409–29.
Ryan, M. 1988. The Irish Horn-reliquary of Tongres/Tongeren,
Belgium. In: G. Mac Niocaill & P.F. Wallace, eds. Keimelia:
O’Connor et al. Advances in identifying archaeological horn and keratinous tissuess
Studies in Conservation 2014 VOL. 0 NO. 024
Studies in Medieval Archaeology and History in Memory of Tom
Delaney. Galway: Galway University Press, pp. 127–42.
Shaffer, B.H., Minx, P., Warren, D., Shedlock, A., Thomson, R.,
Valenzuela, N., Abramyan, J., Amemiya, C., Badenhorst, D.,
Biggar, K., Borchert, G., Botka, C., Bowden, R., Braun, E.,
Bronikowski, A., Bruneau, B., Buck, L., Capel, B., Castoe,
T., Czerwinski, M., Delehaunty, K., Edwards, S., Fronick, C.,
Fujita, M., Fulton, L., Graves, T., Green, R., Haerty, W.,
Hariharan, R. & Hernandez, O. 2013. The Western Painted
Turtle Genome, a Model for the Evolution of Extreme
Physiological Adaptations in a Slowly Evolving Lineage.
Genome Biology, 14: R28.
Siddons, G.A. 1837. The Cabinet Maker’s Guide. London:
Sherwood, Gilbert and Piper.
Smith, G. 1770. The Laboratory, or, School of Arts. London:
Crowder and Collins.
Solazzo, C., Dyer, J.M., Clerens, S., Plowman, J.E., Peacock, E.E. &
Collins, M.J. 2013a. Proteomic Evaluation of the
Biodegradation of Wool Fabrics in Experimental Burials.
International Biodeterioration and Biodegradation, 80: 48–59.
Solazzo, C., Heald, S., Ballard, M.W., Ashford, D.A., DePriest,
P.T., Koestler, R.J. & Collins, M.J. 2011. Proteomics and
Coast Salish Blankets: A Tale of Shaggy Dogs? Antiquity, 85:
1418–32.
Solazzo, C., Wadsley, M., Clerens, S., Dyer, J.M., Collins, M. &
Plowman, J. 2013b. Characterisation of Novel α-keratin
Peptide Markers for Species Identification in Keratinous
Tissues using Mass Spectrometry. Rapid Communications in
Mass Spectrometry, 27: 2685–98.
Solomon, S.E., Hendrickson, J.R. & Hendrickson, L.P. 1986. The
Structure of the Carapace and Plastron of Juvenile Turtles,
Chelonia Mydas (the Green Turtle) and Caretta Caretta (the
Loggerhead Turtle). Journal of Anatomy, 145: 123–31.
Sorge-English, L. 2011. Stays and Body Image in London: The
Staymaking Trade, 1680–1810. London: Pickering and Chatto.
Spindler, K. 1993. The Man in the Ice. London: Weidenfeld and
Nicholson.
Stead, I.M. 2006. British Iron Age Swords and Scabbards. London:
British Museum.
Stevenson, C.H. 1907. Whalebone: Its Production and Utilization.
Department of Commerce and Labor, Bureau of Fisheries
Document #626. Washington: Government Printing Office.
Stock, S. 1976. An Introduction to the Identification of Animal and
Vegetable Fibres used in Antiquity. PhD Dissertation,
University of London Institute of Archaeology.
Szewciw, L.J., de Kerckhove, D.G., Grime, G.W. & Fudge, D.S.
2010. Calcification Provides Mechanical Reinforcement to
Whale Baleen α-keratin. Proceedings of the Royal Society B:
Biological Sciences, 277: 2597–605.
Tombolato, L., Novitskaya, E.E., Chen, P.Y., Sheppard, F.A. &
McKittrick, J. 2010. Microstructure, Elastic Properties and
Deformation Mechanisms of Horn Keratin. Acta
Biomaterialia, 6: 319–30.
Toni, M., Dalla Valle, L. & Alibardi, L. 2007. Hard (Beta-)Keratins in
the Epidermis of Reptiles: Composition, Sequence, andMolecular
Organization. Journal of Proteome Research, 6: 3377–92.
Turgoose, S. 1989. Corrosion and Structure: Modelling the
Preservation Mechanisms. In: R. Janaway & B. Scott, eds.
Evidence Preserved In Corrosion Products: New Fields in
Artifact Studies. London: United Kingdom Institute for
Conservation (Occasional Paper 8), pp. 30–2.
van Doorn, N.L., Hollund, H. & Collins, M.J. 2011. A Novel and
Non-destructive Approach for ZooMS Analysis: Ammonium
Bicarbonate Buffer Extraction. Archaeological and
Anthropological Sciences, 3: 281–9.
Wang, Z., Pascual-Anaya, J., Zadissa, A., Li, W., Niimura, Y.,
Huang, Z., Li, C., White, S., Xiong, Z., Fang, D., Wang, B.,
Ming, Y., Chen, Y., Zheng, Y., Kuraku, S., Pignatelli, M.,
Herrero, J., Beal, K., Nozawa, M., Li, Q., Wang, J., Zhang,
H., Yu, L., Shigenobu, S., Wang, J., Liu, J., Flicek, P., Searle,
S., Wang, J., Kuratani, S., Yin, Y., Aken, B., Zhang, G. &
Irie, N. 2013. The Draft Genomes of Soft-shell Turtle and
Green Sea Turtle Yield Insights into the Development and
Evolution of the Turtle-specific Body Plan. Nature Genetics,
45: 701–6.
Wardlaw, L. & Grant, T. 1994. Treatment of Archaeological Baleen
Artifacts at the Canadian Conservation Institute. Journal of the
International Institute for Conservation – Canadian Group, 19:
31–7.
Watson, J. 1988. Identification of Organic Materials Preserved by
Metal Corrosion Products. In: S.L. Olsen, ed. Scanning
Electron Microscopy in Archaeology. Oxford: Archaeopress.
British Archaeological Reports (B.A.R.), pp. 65–76.
Watson, L. 1981. Sea Guide to Whales of the World. London:
Hutchinson & Co.
Wellman, H. 1997. A Short Note on a Test that Failed. Conservation
News (United Kingdom Institute for Conservation of Historic
and Artistic Works), 62: 30.
Welsh, W., Biscardi, B., Fink, T., Watkins-Kenney, S. &Kennedy, A.
2012. Identification of Suspected Horn from the Queen Anne’s
Revenge (1718), North Carolina, USA. The International
Journal of Nautical Archaeology, 41: 190–3.
Wenham, L.P. 1964. Hornpot Lane and the Horners of York. York:
Annual Report Yorks Philosophical Society.
West, J. & Credland, A.G. 1995. Scrimshaw. The Art of the
Whaler. Hull: City Museums and Art Gallery and Hutton
Press Ltd.
Whittle, A. 2000. ‘Very Like a Whale’: Menhirs, Motifs and Myths
in the Mesolithic-Neolithic Transition of Northwest Europe.
Cambridge Archaeological Journal, 10: 243–59.
Wilkinson, R.S. 2003. Baleen: Its Use as Line Inlay on an 18th
Century Chest of Drawers. Arlington, VA: Wooden Artifacts
Group Postprints.
Wilson, A.S., Dodson, H.I., Janaway, R.C., Pollard, A.M. & Tobin,
D.J. 2007. Selective Biodegradation in Hair Shafts Derived
from Archaeological, Forensic and Experimental Contexts.
British Journal of Dermatology, 157: 450–7.
Wilson, A.S., Dodson, H.I., Janaway, R.C., Pollard, A.M. & Tobin,
D.J. 2010. Evaluating Histological Methods for Assessing Hair
Fibre Degradation. Archaeometry, 52: 467–81.
Young, S.A. 2012. The Comparative Anatomy of Baleen:
Evolutionary and Ecological Implications. MSc Thesis, San
Diego State University.
Zangerl, R. 1969. The Turtle Shell. In: C. Gans, A. d’Bellairs &
T. Parsons, eds. Biology of the Reptilia. London: New York
Academic Press, pp. 311–39.
O’Connor et al. Advances in identifying archaeological horn and keratinous tissues
Studies in Conservation 2014 VOL. 0 NO. 0 25