ThesisPDF Available

The taphonomy of keratin in archosaurs

Authors:

Abstract and Figures

Feather evolution in archosaurs and the preservation potential of proteins in fossils has received much scientific and popular attention in recent years, yet requires further investigation into the taphonomy of keratin, a diverse and common protein. Claims of original keratin proteins in fossils likely used inconclusive methods. This project aims to examine two major questions. 1) Do different keratin types and integumentary structures have different taphonomic patterns which might provide novel information in regards to the evolution of archosaur integument? 2) Does keratin leave any sort of signature in the fossil record? Here, decay and maturation experiments on a variety of keratin types and integumentary structures were carried out to characterize taphonomic changes in archosaur keratin. Morphological, ultrastructural, and chemical changes were observed using scanning electron microscopy and pyrolysis gas chromatography mass spectrometry. Experiments revealed, in contradiction to previous studies, that the rate of bacterial decay in feathers is likely more dependent on the structure of the feather rather than the degree of melanization. Additionally, compression experiments demonstrate that the interpretation of proto-feather morphology in dinosaurs is likely accurate despite criticism. Some variation in taphonomic patterns was observed between different types of keratin resulting in different textures. However, none of these textures have been confidently described in fossils. Feathers matured at 200°C/250 bars and 250°C/250 bars for 24 hours become a viscous, foul-smelling fluid suggesting that over time, proteins fragment and become volatile, meaning that keratins likely do not fossilize. Researchers wishing to study archosaur integument evolution must take into consideration the loss of keratin through taphonomic processes.
Content may be subject to copyright.
The taphonomy of keratin in archosaurs
by EVAN T. SAITTA1
1School of Earth Sciences, University of Bristol, Bristol, UK; e-mail: es14985@bristol.ac.uk
!
!
II!
Abstract: Feather evolution in archosaurs and the preservation potential of proteins in fossils
has received much scientific and popular attention in recent years, yet requires further
investigation into the taphonomy of keratin, a diverse and common protein. Claims of
original keratin proteins in fossils likely used inconclusive methods. This project aims to
examine two major questions. 1) Do different keratin types and integumentary structures
have different taphonomic patterns which might provide novel information in regards to the
evolution of archosaur integument? 2) Does keratin leave any sort of signature in the fossil
record? Here, decay and maturation experiments on a variety of keratin types and
integumentary structures were carried out to characterize taphonomic changes in archosaur
keratin. Morphological, ultrastructural, and chemical changes were observed using scanning
electron microscopy and pyrolysis gas chromatography mass spectrometry. Experiments
revealed, in contradiction to previous studies, that the rate of bacterial decay in feathers is
likely more dependent on the structure of the feather rather than the degree of melanization.
Additionally, compression experiments demonstrate that the interpretation of proto-feather
morphology in dinosaurs is likely accurate despite criticism. Some variation in taphonomic
patterns was observed between different types of keratin resulting in different textures.
However, none of these textures have been confidently described in fossils. Feathers matured
at 200°C/250 bars and 250°C/250 bars for 24 hours become a viscous, foul-smelling fluid
suggesting that over time, proteins fragment and become volatile, meaning that keratins
likely do not fossilize. Researchers wishing to study archosaur integument evolution must
take into consideration the loss of keratin through taphonomic processes.
Key Words: decay, maturation, SEM, pyrolysis GC/MS, feathers, scales
!
!
III!
Acknowledgements. Many thanks to Charles Clapham (University of Bristol) for his
assistance in designing, constructing, and operating the compression rig, Richard Brooker
(University of Bristol) for his assistance in autoclaving, Jakob Vinther (University of Bristol)
and Chris Rogers (University of Bristol) for their assistance in experimentation throughout
the project as well as their guidance and feedback, Antoine Soler (La Ferme aux Crocodiles)
for dissecting and shipping the crocodile skin, Paul Donohoe (Newcastle University) and
Geoff Abbott (Newcastle University) for running the Py/GC/MS and helping to interpret the
data, and Erzsebet!Thornberry (University of Bristol) and Ian Bull (University of Bristol) for
assisting in Py/GC/MS data interpretation and providing useful feedback.
Declaration: I declare that the work contained in this thesis is the author’s own, except
where stated or referenced. The views expressed in this thesis are those of the author and do
not represent those of the University of Bristol.
Date: 9th November 2015
!
!
1!
1. INTRODUCTION
Soft tissue preservation in the fossil record is relatively rare (Allison & Briggs 1993)
meaning that exceptionally preserved specimens receive much scientific interest. Discoveries
of avian and non-avian dinosaur fossils with preserved feathers and filaments from Early
Cretaceous deposits of northeastern China have transformed our understanding of dinosaur
and bird evolution and lead to radical reinterpretations of dinosaur biology and appearance
(Xu & Guo 2009). Despite the abundance of fossil material, the point at which feathers
appear in the evolutionary tree of archosaurs (the group that includes the most recent
common ancestor of all modern birds and crocodilians and all of its descendants) is unknown.
Crucial evidence might come from a better understanding of soft tissues in fossil archosaurs –
in this case, keratin.
1.1 Keratin characteristics
Keratins are a highly diverse group of fibrous structural proteins found in hair,
feathers, scales, horns, hooves, nails, claws, beaks, and skin and are produced by epidermal
cells. They are a type of intermediate filament protein and the resulting polymer is insoluble,
flexible, and hard. Two main groups of keratin exist: α-keratin and β-keratin (Fraser et al.
1972). Disulphide bonds add stability to the protein, making keratin insoluble and resistant to
enzymatic digestion from proteases (Korniłłowicz-Kowalska & Bohacz 2011). Glycine-rich
sequences are also thought to make keratins more insoluble (Steinert & Freedberg 1991). The
α- and β-keratins possess α-helical and (more rigid) β-pleated sheet secondary structure,
respectively (Pauling et al. 1951). The epidermis of all amniotes, including reptile and bird
interscale regions and skin as well as mammal skin and integumentary appendages, contain
!
2!
α-keratin. The β-keratins are only found in reptile and bird epidermis and integumentary
appendages (which also contain α-keratin). The β-keratins are smaller proteins than α-
keratins, but have similar basic chemistries and abilities to form filaments (Prum & Brush
2002). The β-keratins typically contain high amounts of histidine, lysine, and tryptophan,
while having low amounts of cystine (Hill et al. 2010). They consist of a tightly packed
lattice of microfibrils with an ultrastructure of 3–4 nm filaments (Fraser et al. 1972).
1.2 Extant archosaur keratinous integumentary appendages
Crocodilian scales. Crocodilian scales possess a thin, flexible layer called the hinge
region made of α-keratin (40–68 kDa) and a hard, thick, corneous outer surface made of
mostly β-keratin (10–22 kDa) (Wyld and Brush 1979, 1983, Sawyer et al. 2000, Alibardi &
Toni 2006). Crocodilian β-keratin sequences have high levels of tyrosine, glycine, serine, and
proline and contain cysteines towards the terminal regions (Dalla Valle et al. 2009).
Feathers. Feathers have been defined as tubular integumentary structures made of
feather keratin originating from a feather follicle (Prum & Brush 2002). Modern feathers are
made up of a subgroup of β-keratins called φ-keratins (Brush 1978). They have four repeating
units of two β-pleated sheets, creating a helical structure that is surrounded by a matrix
(Greenwold & Sawyer 2010). X-ray diffraction has found a 32 amino acid sequence that
makes up the 2–3 nm filament. The remaining 65 amino acid residues form the matrix (Fraser
& Parry 2008). The amino acid composition of φ-keratins has been described as mostly
proline, serine, and glutamic acid with little methionine, lysine, and histidine. This differs
from other β-keratins (Korniłłowicz-Kowalska & Bohacz 2011).
!
3!
Avian scales. Avian scutate scales are the large, rectangular, overlapping scales on the
anterior metatarsus and dorsal digit region (Shames et al. 1989). Avian reticulate scales are
the scales on the plantar foot surface. Avian scutate scales, claws, and beaks consist of
slightly longer φ-keratins than do feathers – 13.5 kDa versus 10.4 kDa (Prum & Brush 2002).
Crocodilian claw keratin is 14 kDa and is highly related in sequence to the φ-keratins in avian
scutate scales, beaks, and claws, making φ-keratins widely distributed in archosaurs (Sawyer
et al. 2000). This φ-keratin has also been observed in embryonic alligator scales (Alibardi et
al. 2005). Some have proposed that φ-keratins appeared in an Archosaurian ancestor to birds
and crocodilians, diversifying into shorter filaments in feathers and longer filaments in avian
scutate scales, claws, and beaks, and that scutate scales evolved via modification of leg
feathers. Avian reticulate scales are made up of non-featherlike β-keratins similar to those
from other reptiles (Prum & Brush 2002).
Turkey beards. Adult wild turkeys (Meleagris gallopavo) have bristles on the ventral
side of their neck, which are outgrowths that grow continuously from a region of elevated
skin. Bristles are generated from finger-like epidermal outgrowths that, unlike a feather
follicle, do not show invagination at their base. Distal ends of dermal cores eventually retract,
leaving a corneous tip and hollow bristle (Schorger 1957). Beards are known to consist of
both feather-type and avian scale-type φ-keratins. Bristles can show simple branching,
irregular cross-sectional shape, and hollow tips, although some have cellular elements in
central canals that are possibly connective tissue. Some doubt the homology of turkey beards
to other feathers (Sawyer et al. 2003).
Non-archosaur keratinous integumentary appendages. Mammalian hair is made
primarily of α-keratins (~60 kDa, low in sulphur, with right-handed α-helixes) with an
!
4!
increasingly complex structural hierarchy (e.g., α-helixes , coiled-coils, and microfibrils).
Sulphur rich, globular γ-keratins (~15 kDa) form the matrix around microfibrils (Hill et al.
2010).
1.3 Ancient biomolecules
The taphonomy of keratin ultimately ties into the question of why some biomarkers
are preserved in the fossil record and why others are not. Biomolecules vary in preservation
potential; from least to most robust, these are: nucleic acids, proteins, carbohydrates, and
lipids. Thus, most biomarkers are derived from lipids (Briggs & Summons 2014). Many
examples of long chain aliphatic compounds have been reported in fossils of graptolites
(whose periderm consists of collagen in life, but whose fossils lack protein), arthropod
cuticles, leaves, and fish scales, and maturation and decay experiments, in combination with
pyrolysis gas chromatography mass spectrometry (Py/GC/MS), have revealed that these
aliphatic compounds derive from lipids via in situ polymerization (Briggs 1999, Gupta et al.
2006a, 2006b, 2007a, 2007b, 2007c, 2008, 2009). These examples are not too dissimilar from
the diagenetic changes that occur during the formation of fossil fuels from the lipids of
photosynthetic organisms (Brooks & Smith 1967, 1969). The above suggest that
biomolecules can be preserved but are often altered. Recently, porphyrins derived from heme,
a cofactor of the protein hemoglobin, were detected in the gut of a 46 million-year-old, blood
engorged mosquito fossil in oil shale. This suggests that certain complex organic molecules
can extend deep into to fossil record, and the researchers suggest that the structure of heme
promotes its longevity, specifically the fact that it is stabilized by iron (Greenwalt et al.
2013).
!
5!
The peptide bonds that form between amino acids to form polypeptides and proteins
are thermodynamically unstable in aqueous solutions. However, under certain conditions
(neutral pH and ambient temperatures) the rate of hydrolysis is slow, providing stability to
peptide bonds (Nakai & Modler 1996), resulting in their preservation in the fossil record.
Type I collagen has been reported from Pleistocene aged bone (Orlando et al. 2013). It has
been proposed that the crystalline matrix in which the collagen resides in, its triple helix
structure with intra- and inter-molecular cross-links, and abundance of thermally stable amino
acids confers stability to the protein. The six most stable amino acids are alanine, glycine,
valine, leucine, phenylalanine, and proline (with glycine and alanine being the most stable).
These amino acids comprise 62% of type I collagen, placing them in the top 0.09–0.31% of
their proteomes in a ranking of the percent abundance of these thermally stable amino acids
(Wang et al. 2012). As discussed in 1.2, the amino acid concentrations of different keratins
can vary, but can often be relatively rich in thermally stable glycine and proline. However,
they can also be rich in less thermally stable amino acids such as serine or cysteine. One
study of feather plumage found that serine, proline, systeine, and glycine (in that order) were
the most abundant amino acids, comprising 46 mol% of the protein (Murphy & King 1982).
It seems that keratin is likely less thermally stable than collagen.
1.4 Keratin taphonomy
Despite the importance of keratin, relatively little is known about how it degrades and
fossilizes. Poultry farming has led to an interest in biodegradation of feather waste (Haddar et
al. 2009), but relatively little work has been done on the degradation of keratins as it might
relate to the fossil record. There have been many hypothesized mechanisms by which
organisms such as bacteria or fungi degrade keratin, though the process is not entirely
!
6!
understood (Korniłłowicz-Kowalska & Bohacz 2011). Bacteria and fungi are known to
secrete metalloenzymes utilizing various metal ions that deposit onto the keratin surface
(Gupta & Ramnani 2006). Another study found melanized feathers to be more resistant to
feather degrading bacteria than non-melanized feathers (Gunderson et al. 2008).
Studies on archaeological keratin have demonstrated that a high pH results in rapid
decay and that the action of microorganisms (biodeterioration) plays a larger role in keratin
decay than changes in moisture, pH, or temperature (chemical deterioration). Fungi are
particularly detrimental to the ordered structure of keratin; a variety of fungi in soil
enzymatically break disulphide bridges and denature the proteins. Keratinous materials rarely
survive in archaeological sites unless fungal biodeterioration is inhibited through low
temperatures, low humidity, anoxia, and limiting pH (O’Connor et al. 2014). Despite
prevalent biodeterioration, keratins can still survive in the archaeological record as mineral
preserved organic remains when in contact with corroding metals, the products of which coat
and protect the organic material. (Solazzo et al. 2013a, O’Connor et al. 2014, Solazzo et al.
2014). Deamidation of asparagine and glutamine can act as markers for protein decay and
this technique has been used on archaeological wool (Solazzo et al. 2013b).
Keratinous structures with relatively high levels of biomineralization of calcium
phosphate (e.g., rachises and claws) are known from fossils since the inorganic component
can be preserved (Benton et al. 2008), but can the keratin protein itself survive? One study
claimed to have identified components of β-keratin protein in the filaments of the theropod
Shuvuuia from the Late Cretaceous using modern duck feathers as a control. In combination
with scanning and transmission electron microscopy (SEM and TEM, respectively) and time
of flight secondary ion mass spectrometry (TOF-SIMS) they found structural and chemical
signatures consistent with β-keratin. More surprisingly, when exposed to avian anti-β-keratin
antisera, an immunoreactive response was observed in the filaments (Schweitzer et al. 1999).
!
7!
Might these results represent a false positive – a risk that must be accounted for when doing
immunohistochemistry (True 2008)? Fourier transform infrared spectroscopy (FTIR) was
used by one group of researchers to demonstrate the presence of amide groups in fossil reptile
skin, including a Late Cretaceous “mummy” hadrosaur, which they took as evidence for the
breakdown products of (or even the original) β-keratin protein. Py/GC/MS data from the
hadrosaur was interpreted to show endogenous, partly aliphatic organic material in the skin
distinct from the silicate matrix. Py/GC/MS of Eocene fossil reptile skin was interpreted
similarly as the organic material was distinct from the data obtained from the sediment matrix
as well as a fossil plant from the same formation (Manning et al. 2009, Edwards et al. 2011,
Bergmann et al. 2012). However, keratins are extremely rare even in Quaternary fossils
(Hofreiter et al. 2012). Conclusive keratin protein identification in fossils, particularly of
Mesozoic age, is currently lacking.
While keratin protein identification in fossils is questionable, many have studied
melanin pigment through examination of fossil melanosomes that were contained within the
keratin of feathers and scales during life. However, these studies do not report the presence of
keratin (Vinther et al. 2008, Vinther 2015). The taphonomy of melanin has also drawn some
interest as a result of this work. As the amount of surviving fossil eumelanin decreases
through maturation, the relative amount of kerogen in these specimens increases (Glass et al.
2013). It is unknown how keratin decay and maturation affects melanosome arrangement.
Py/GC/MS and FTIR of a fossil tadpole with melanosomes has shown that its organic
remains do not represent contamination because chemical composition differs between fossil
and matrix. Plant and bacterial biomarkers are present in the matrix and not the fossil,
indicating that these melanosomes are not mistakenly identified bacteria (Barden et al. 2015).
!
8!
1.5 Project aims
The aims of this project are to determine 1) if different keratin types and
integumentary structures show distinct patterns of decay and/or maturation that might provide
insight into their evolution and 2) what signatures keratin structures might leave in the fossil
record or if keratin can survive in fossils at all. These two questions will be addressed over a
range of levels (chemical, ultrastructural, and morphological). This means that observations
of the chemical nature of keratin all the way up to taphonomic effects on whole body feather
plumage will be made. Depending on what sorts of signatures keratin leaves in fossils,
potential taphonomic differences between evolutionarily distinct integumentary structures
might allow for a better understanding of integument evolution in archosaurs. Ideally,
different keratin types might be distinguishable in fossils. Given the relatively limited
sampling of soft tissues across Archosaurian phylogeny, understanding keratin taphonomy
may tell us to what extent fossils can be informative in understanding the evolution of
keratinous appendages.
2. MATERIAL AND METHODS
Samples of various archosaur integumentary structures, with mammalian hair as an
outgroup, underwent decay and maturation treatments (Table 1). Samples were either (a)
fresh, (b) bacterially decayed (using cultured, naturally occurring feather bacteria in a salt
broth at ~37°C for 50 days), (c) matured and non-decayed, or (d) matured after decay. These
were imaged using light microscopy (Leica M205 C stereomicroscope with a Leica DFC425
C camera) and SEM (Hitachi S3500N variable-pressure scanning electron microscope under
vacuum, scanning electron mode) and chemically analyzed using Py/GC/MS. Most samples
!
9!
were matured at a moderate temperature (100°C/250 bars for 24 hours) to allow for structural
changes to be observed. However, two samples of feathers, one dark and one white
(Supporting Information), were matured at higher temperature (250°C/250 bars for 24 hours)
to produce a foul-smelling, viscous fluid (referred to here as a ‘goo’) for analysis. The
solubility of these highly matured samples was investigated using a binocular microscope
(Nikon Eclipse LV100D–U stereomicroscope). Maturation of white feathers at 200°C/250
bars for 24 hours also produced a ‘goo’, but the sample leaked out of the autoclave tube and
could not be analyzed.
Sample (Abbreviation)
250
°
C, 250 bars, 24 hours
White feathers – matured into ‘goo’ (GW)
Dark feathers – matured into ‘goo’ (GD)
100
°
C, 250 bars, 24 hours
Feathers – iridescent (FI)
Feathers – white (FW)
Feathers – black (FB)
Avian scutate scales (ST)
Avian reticulate scales (RT)
Turkey beard – adult (BA)
Turkey beard – juvenile (BJ)
Crocodilian scale – black (CB)
Crocodilian scale – white (CW)
Mammalian hair (H)
Decayed iridescent feathers (DFI)
Decayed white feathers (DFW)
Decayed black feathers (DFB)
Table 1. List of maturation runs. Samples that are not listed as decayed are fresh samples.
!
10!
Additionally, compression experiments (25 tonnes for approximately one minute)
were carried out in a specially designed rig on entire zebra finch (Teaniopygia guttata)
carcasses to mimic sub-aqueous burial and compression. Low energy sediment deposition
was mimicked and pressure was applied while carcasses were buried in waterlogged mud. To
study the taphonomy of whole body feather plumage, experiments were run on one fresh bird
and one bird bacterially decayed while sub-aqueously buried, prior to compaction, (using
naturally occurring soil bacteria at ~37°C for 46 days).
See Supporting Information for full methods.
3. RESULTS
3.1 Structural analysis
Fresh samples. The adult turkey beard was purchased as a dried specimen and the
epidermis has a flaky appearance under microscopy as a result (Fig. 1A, F). The bristles
themselves also show a flaky appearance on their keratin surface when viewed in SEM under
high magnification (Fig. 1B–C). Their cross sections show a solid, homogeneous structure
and when cut, the bristles show fraying, splitting, and flaking at their ends (Fig. 1D–E).
The juvenile turkey beard shows similar features to the adult beard with a flaky or
‘scaly’ texture to the epidermis (Fig. 2A, F) and a flaky texture to the keratin of the bristles
under higher magnification (Fig. 2B–C). The bristle cross sections also are homogenous and
prone to splitting or fraying at their ends when cut (Fig. 2D–E). The variation in bristle cross-
sectional shape is more apparent in the juvenile than the adult, with some bristles flattened
compared to more rounded bristles.
!
11!
The predominantly black crocodile scale shows a non-uniformly pigmented outer
surface with a light colored, soft internal layer (Fig. 3A–B). The outer surface shows
prominent flaking or cracking resulting in sub-polygonal flakes (Fig. 3C). At greater
magnification, a rippled texture is apparent on the surface of the outer layer (Fig. 3D). A
sharp contact can be observed between the outer and inner layers in cross section (Fig. 3E).
The inner layer shows a more fibrous composition than the outer layer (Fig. 3F).
The predominantly white crocodile scale shows the same structure as the
predominantly black scale. The outer surface is not uniformly pigmented (Fig. 4A), shows
prominent flaking or cracking resulting in sub-polygonal flakes (Fig. 4B), and has a rippled
pattern at higher magnification (Fig. 4C). There is a sharp contact between the outer and
inner layers (Fig. 4D–E). The outer layer’s cross section reveals that it is composed of
multiple, overlapping keratinous layers and even the deeper layers show the rippled
patterning on their surface (Fig. 4F).
The black feather has a white fringe (Fig. 5A). The rachis surface is simple but some
topographic variability is apparent (i.e., it is not a completely smooth surface) (Fig. 5C). The
calamus shows more complex keratin layering and folding that the rachis (Fig. 5D–E). The
rachis cross section shows a central pith filled with what appear to be medulloid cells and/or
melanosomes and an outer cortex with a columnar pattern radiating outward (Fig. 5F).
The iridescent feather shows the most prominent iridescence at the distal portion of
the vane (Fig. 6A). In the iridescent region, the barbules show a unique, broadened
morphology (Fig. 6B). The plumulaceous barbules show evenly-spaced nodes (Fig. 6C). The
rachis is relatively smooth but the calamus has more complex surface topology (Fig. 6D). The
cross sections of the barbs show compartmentalization consisting of sub-polygonal medulloid
cells (Fig. 6E). The rachis cross section has a central pith consisting of medulloid cells and/or
melanosomes (Fig. 6F). However, the outer cortex is disorganized and does not show the
!
12!
columnar structure seen in the black feather, and the cross-sectional shape is overall more
irregular.
The white feather (Fig. 7A) has pennaceous barbules that do not show the same
proximal-distal variation seen in the black feather, but instead consist of the thickened ridge
on one side that supports a flattened vane (Fig. 7B). The plumulaceous barbules show evenly-
spaced nodes (Fig. 7C). The rachis surface is not perfectly smooth, but is not relatively
complex topographically (Fig. 7D–E). The rachis cross section shows an outer layer with
columnar structuring and a small central pith consisting of medulloid cells (Fig. 7F).
The horse hair is darkly pigmented and coarse (Fig. 8A). The surface of the hair
shows flaking in a manner reminiscent of the turkey beard bristles (Fig. 8B–D). The cross
section is solid, but disorganized (Fig. 8E–F). The cut end of the strand did not split or fray
like the turkey bristles did. The turkey bristles also show a more organized, almost concentric
structuring of keratin in cross section compared to the more clumped appearance of the
keratin in the hair.
The turkey reticulate scales are sub-polygonal, with distinct boundaries between each
scale (Fig. 9A–B). The surface of the outer layer is rough, reminiscent to the surface of the
outer layer of crocodile scales (Fig. 9C) and at higher magnification is appears
topographically complex (Fig. 9D). The cross section shows a fairly well defined contact
between an inner and outer layer (Fig. 9E–F). The cross section of the outer layer is fairly
homogenous.
The turkey scutate scales are large and sub-rectangular (Fig. 10A). They show flaking
on the outer surface (Fig. 10B–E). However, other than this flaking, the surface is not
topographically complex and is very smooth, unlike the rippled texture of the outer surface of
crocodile or even the turkey reticulate scales. Even at high magnification, the only
topographic variation appears to primarily come from flaking of the outer surface. The cross
!
13!
section shows a fairly well defined contact between the inner and outer layers (Fig. 10F). The
outer layer appears to show splitting or flaking in cross section but is otherwise homogenous.
FIG. 1. Structural features of ‘fresh’ (although dried) adult turkey beard. A, under light
microscopy, and B–F, SEM. B–C, bristle surfaces. D–E, bristle cross section. F, epidermal
surface.
!
14!
FIG. 2. Structural features of fresh juvenile turkey beard. A, under light microscopy, and B–
F, SEM. B–C, bristle surfaces. D–E, bristle cross sections. F, epidermal surface.
!
15!
FIG. 3. Structural features of fresh predominantly black crocodile scale. A–B, under light
microscopy, and C–F, SEM. A, cross section. B–D, outer surface. E, cross section at contact
between outer (above) and inner (below) layers. F, cross section of inner layer.
!
16!
FIG. 4. Structural features of fresh predominantly white crocodile scale. A, under light
microscopy, and B–F, SEM. A, view of outer surface in dorsal view with inner layer also
visible to the right. B–C, outer surface. D–E, cross section at contact between outer (above)
and inner (below) layers. F, cross section of outer layer.
!
17!
FIG. 5. Structural features of fresh black feather. A, under light microscopy, and B–F, SEM.
A, white fringe to barbs is visible. B, pennaceous barbs and barbules. C, rachis surface at
distal end. D–E, calamus surface. F, rachis cross section.
!
18!
FIG. 6. Structural features of fresh iridescent feather. A, under light microscopy, and B–F,
SEM. B, pennaceous barbules in iridescent region. C, plumulaceous barb and barbules. D,
calamus surface. E, barb cross section. F, rachis cross section.
!
19!
FIG. 7. Structural features of fresh white feather. A, under light microscopy, and B–F, SEM.
B, rachis and pennaceous barbs and barbules. C, plumulaceous barbules. D–E, rachis surface.
F, rachis cross section.
!
20!
FIG. 8. Structural features of fresh horse hair. A, under light microscopy, and B–F, SEM. B–
D, surface. E–F, cross section.
!
21!
FIG. 9. Structural features of fresh turkey reticulate scales. A, under light microscopy, and
B–F, SEM. B–D, outer surface. E–F, cross section. E, exterior surface towards top of image.
F, exterior surface towards right of image.
!
22!
FIG. 10. Structural features of fresh turkey scutate scales. A, under light microscopy, and B–
F, SEM. B–E, outer surface. F, cross section with exterior surface towards top of image.
Decayed samples. The control feathers (darkly pigmented feathers in mineral water)
showed no major signs of degradation even when examined in SEM. The feathers were still
whole and showed full feather morphology (Supporting Information).
!
23!
The decayed black feather showed the highest level of degradation, resulting in a lint-
like mass of tiny feather bits (Fig. 11A). Most of this ‘lint’ was dark with only a few white
remnants of the calamus or rachis present. When examining the sample only rinsed with
ethanol after decay, the dark bits appear to be barbs and barbules that are both interlocking as
well as partially fused together (Fig. 11B). Upon higher magnification, the surface of the
barbs appears to show extreme flaking (Fig. 11C–D), although the exact identity of some of
these structures had to be further examined with the further prepared samples (see below).
They could be bacteria, although this is unlikely given that bacteria would be expected to
burst under the vacuum conditions of SEM. Bits of the rachis or calamus are apparent under
SEM, and the surface shows a degraded, woven texture (Fig. 11E–F).
The sample of decayed black feather that was critically point dried after rinsing with
ethanol appears to also show a flaking pattern to the keratin (Fig. 12A) distinct from well
defined sausage-shaped structures that could either be melanosomes or bacteria (Fig. 12B–C).
The structures can often be seen resting on a degraded keratin surface with a woven texture.
The sample of decayed black feather that was rinsed with ethanol and then treated with Triton
X-100 before critical point drying also shows these sausage-shaped structures (Fig. 12 D–F),
proving that they are melanosomes and not bacteria. The degraded, woven texture of the
keratin is also apparent in these samples.
The decayed iridescent feathers showed the second highest levels of degradation,
resulting in small bits not quite as fine as the ‘lint’ produced from the decayed black feathers
(Fig. 13A). The original feather structure was mostly lost, but more morphological structure
was retained than in the decayed black feathers. As in the decayed black feathers, much of the
remnants consisted of a mass barbs and barbules. Iridescence was even preserved (Fig. 13B).
When examining the sample only rinsed with ethanol after decay, many pennaceous and
plumulaceous barbules are apparent, such as the pennacous barbules consisting of the
!
24!
thickened ridge on one side that supports a flattened vane (Fig. 13C–D). Only a few remnants
of the rachis are present and the surface appears relatively wrinkled (Fig. 13E–F).
The sample of decayed iridescent feather that was critically point dried after rinsing
with ethanol shows fraying of the rachis surface (Fig. 14A), wrinkled keratin surface with
potential bacteria or melanosomes on the surface (Fig. 14B), and exposed medulloid cells
from the pith, some of which appear to be imploded (Fig. 14C). The sample of decayed
iridescent feather that was rinsed with ethanol and then treated with Triton X-100 before
critical point drying shows a degraded, woven texture to the keratin surface (Fig. 14D),
plumulaceous barbules with well-preserved structure and some flaking on their surface (Fig.
14E), and many imploded medulloid cells exposed from the pith.
The decayed white feathers showed the lowest level of degradation among the
experimental feathers. The entire macroscopic feather morphology was preserved (Fig. 15A),
and the decayed white feathers were most similar to the control feathers in the mineral water
in terms of level of degradation. Examining the sample only rinsed with ethanol after decay
revealed that the calamus surface appears to be flaking (Fig. 15B–C). However, much of the
keratin surface appears to be relatively unaltered (Fig. 15D). Interestingly, one region of the
rachis surface shows short, hair-like structures extending off of the surface when viewed
under high magnification (Fig. 15E). The cross section of the rachis is typical with an outer
cortex showing a columnar, radiating structure and and inner pith filled with medulloid cells
(Fig. 15F).
The sample of decayed white feather that was critically point dried after rinsing with
ethanol shows similar features to the sample simply rinsed in ethanol and reveals little keratin
surface degradation on the barbs, barbules, and rachis (Fig. 16A–C). It also shows a normal
rachis cross section with outer cortex and pith represented (Fig. 16D). The sample of decayed
white feather that was rinsed with ethanol and then treated with Triton X-100 before critical
!
25!
point drying provides similar information. The calamus surface shows little degradation (Fig.
16E–F).
FIG. 11. Structural features of decayed black feathers rinsed with ethanol after decay
treatment. A, under light microscopy, and B–F, SEM. B–D, fused mass of barbules. E–F,
rachis or calamus surface.
!
26!
FIG. 12. SEM images of structural features of decayed black feathers rinsed with ethanol
after decay treatment and then prepared further. A–C, samples critically point dried. D–F,
samples treated with Triton X-100 detergent and then critically point dried.
!
27!
FIG. 13. Structural features of decayed iridescent feathers rinsed with ethanol after decay
treatment. A–B, under light microscopy, and C–F, SEM. B, iridescent pennaceous barbules.
C, pennaceous barbules. D, plumulaceous barbules. E–F, rachis surface.
!
28!
FIG. 14. SEM images of structural features of decayed iridescent feathers rinsed with ethanol
after decay treatment and then prepared further. A–C, samples critically point dried. D–F,
samples treated with Triton X-100 detergent and then critically point dried.
!
29!
FIG. 15. Structural features of decayed white feathers rinsed with ethanol after decay
treatment. A, under light microscopy, and B–F, SEM. B–C, calamus surface. D, pennaceous
barb and barbules. E, rachis surface. F, rachis cross section.
!
30!
FIG. 16. SEM images of structural features of decayed white feathers rinsed with ethanol
after decay treatment and then prepared further. A–D, samples critically point dried. E–F,
samples treated with Triton X-100 detergent and then critically point dried. A, pennaceous
barb and barbules. B, rachis and barb surface. C, plumulaceous barbule. D, rachis cross
section. E–F, calamus surface.
!
31!
Matured samples. The following are maturation experiments at 100°C/250 bars for 24
hours.
The matured adult turkey beard showed bristles with a wrinkled texture (Fig. 17A). In
addition to wrinkling, the bristles appear to be flaking (although some of this might be
matured epidermis adhering to the bristles) (Fig. 17B–C). The bristle cross section is strongly
compressed in some areas (Fig. 17D), and this particular example shows how some bristles
are not completely solid, but can have a hollow central region. The epidermis is extremely
degraded and consists of thin flakes (Fig. 17E–F).
The matured juvenile turkey beard shows curling in the bristles and discolored
epidermis (Fig. 18A). The tight curls of the bristles are accompanied by degradation of the
bristle surface as seen by the rough texture (Fig. 18B–C). The cross sections of the bristles
are solid and compressed, but the keratin still appears somewhat organized (Fig. 18D–E). The
epidermis is degraded and appears to be flaking similar to that seen in the matured adult
turkey beard (Fig. 18F).
The matured predominantly black crocodile scale retains some of it’s original color
variation (Fig. 19A). The hard outer layer shows splitting between its sub-layers and cracking
(Fig. 19B–C). The rippled texture of the outer layer’s surface was still retained in some areas
(Fig. 19D). The internal layer of the scale can be distinguished from the hard outer layer and
shows a lot of degradation (Fig. 19E). This internal layer has become soft (Fig. 19F).
The matured predominantly white crocodile scale consists of harder and softer
products representing the outer and inner layers of the scale, respectively, and retains its light
coloration on it’s outer layer, although some discoloration appears to have occurred (Fig.
20A–B). The outer layer shows cracking, flaking, and splitting between its sub-layers (Fig.
20C). These sub-layers still retain the original rippled texture (Fig. 20D). The inner layer is
more degraded and exhibits pock-marking (Fig. 20E–F).
!
32!
The matured black feather shows a breakdown of the overall feather structure,
retention of original coloration variation, and curling of the keratin (Fig. 21A). Many subunits
of the feather are still present, however, such as barbs and barbules, although these are
incomplete and show curling and kinking (Fig. 21B–F). The rachis could not be found in the
sample.
The matured decayed black feather shows extreme degradation resulting in a black
pellet (Fig. 22A). Some of it consists of a mass of fused barbs and barbules (Fig. 22B),
although other areas appear to derive from the rachis and have a flat surface with cracks (Fig.
22C). Yet other regions show a granulated texture (Fig. 22D). Only a small amount of
original structure remains, such as some barbules attached to barbs, but these have highly
degraded surfaces and are flattened (Fig. 22E–F).
The matured iridescent feather consists of clumped, folded barbs and some of the
iridescence remains (Fig. 23A). The morphology of the barbs at low magnification appears
relatively well conserved, despite maturation. The barbs are closely packed together and the
barbs and barbules are often incomplete (Fig. 23B–D). One portion of the sample appears to
represent part of the rachis (Fig. 23E). The keratin shows signs of degradation such as
wrinkling on the barbs and barbules surfaces (Fig. 23D) and flaking of the rachis (Fig. 23F).
The matured decayed iridescent feather may have been exposed to water during
autoclaving as the capsule gained weight, suggesting a failed seal and infiltration of water
(Supporting Information). However, the sample was analyzed anyway. The sample shows
extreme degradation resulting in a black pellet (Fig. 24A) similar to that seen in the matured
decayed black feather. The keratin surface of what appears to be the remnants of the rachis is
cracked and fraying into strips or strands (Fig. 24B–C) These strands are irregular in cross-
sectional shape with folding and creases common and are sometimes associated with thin,
!
33!
filaments (Fig. 24D–E). Some degraded barbules attached to remnants of barbs are visible
and the barbules appear shriveled (Fig. 24F).
The matured white feather shows high levels of degradation and the predominant
remnant is the rachis/calamus (Fig. 25A). However, some plumulaceous barbs are still
apparent whose surfaces are moderately degraded as evidenced by minor wrinkling and
flaking (Fig. 25B–C). The surface of the calamus is highly degraded with wrinkling,
cracking, flaking, and fraying apparent (Fig. 25 D–F). The fraying produces strips of keratin
like that seen in matured decayed iridescent feather.
The matured decayed white feather appears similar to the matured white feather at
low magnification with predominantly the rachis/calamus surviving (Fig. 26A). However, it
might show the highest levels of degradation at the ultrastructural level of all the feather
samples examined in this study other than the highly matured ‘goo’. Plumulaceous barbs are
still apparent at one end of the sample (Fig. 26B). However, the surface of the calamus shows
a woven, and in some places cracked, texture and the proximal end shows internal layering of
strands (Fig. 26C–F) that at first seem similar to the strands seen in the matured white feather
and the matured decayed iridescent feather. However, these strands appear to be well
organized and overlapping with regularly spaced nodes. Overall they appear almost identical
to plumulaceous barbules (Fig. 26E). These strands can be observed originating from more
continuous and less degraded regions of the calamus (Fig. 26D).
The matured horse hair shows curling and extreme kinking in the strands (Fig. 27A–
B). Some strands are splitting down their length (Fig. 27C) and the keratin surface is flaking
(Fig. 27D). The kinks sometimes occur in a helical pattern down the strand (Fig. 27E). The
cross section of the strand is solid, yet disorganized (Fig. 27F).
The matured turkey reticulate scales are highly degraded and show softer and harder
remnants under high magnification, representing the inner and outer layers of the scales,
!
34!
respectively (Fig. 28A). However, overall the sample is much more pliable than the fresh
scales. The harder remnants show flaking at the surface and the original layering is revealed
with wrinkled surface texture (Fig. 28B–C). The softer remnants are clumped (Fig. 28D) and
show many pock-marks on their surface (Fig. 28E). In addition to pock-marks, bulges and
folds of the organic material are also apparent along with structures intermediate between the
pock-marks and the bulges (Fig. 28F).
The matured turkey scutate scale is also highly degraded and shows a more pliable
structure than the fresh scales (Fig. 29A) with potentially harder and softer remnants when
viewed under high magnification deriving from the outer and inner layers, respectively. The
surface is wrinkled and flaking and some areas show a rough texture, representing
degradation (Fig. 29B–F).
Regarding the highly matured samples (250°C/250 bars for 24 hours), neither the
‘goo’ derived from the dark nor the white feathers show microscopic features such as
exposed melanosomes, and the only topography derives from uneven dispersal of the ‘goo’
across the capsule surface (Fig. 30A–F). A sample of white feathers matured at 200°C/250
bars for 24 hours also produced a ‘goo’ that unfortunately leaked from the autoclave tube and
could not be analyzed (Supporting Information).
!
35!
FIG. 17. Structural features of matured adult turkey beard. A, under light microscopy, and
B–F, SEM. B–C, bristle surfaces. D, bristle cross section. E–F, epidermal surface.
!
36!
FIG. 18. Structural features of matured juvenile turkey beard. A, under light microscopy, and
B–F, SEM. B–C, bristle surfaces. D–E, bristle cross section. F, epidermal surface.
!
37!
FIG. 19. Structural features of matured predominantly black crocodile scale. A, under light
microscopy, and B–F, SEM. A–D, harder remnants. E, softer and harder remnants. F, softer
remnants.
!
38!
FIG. 20. Structural features of matured predominantly white crocodile scale. A, under light
microscopy, and B–F, SEM. A, C–D, harder remnants. B, E–F, softer remnants.
!
39!
FIG. 21. Structural features of matured black feather. A, under light microscopy, and B–F,
SEM.
!
40!
FIG. 22. Structural features of matured decayed black feather. A, under light microscopy,
and B–F, SEM.
!
41!
FIG. 23. Structural features of matured iridescent feather. A, under light microscopy, and B–
F, SEM.
!
42!
FIG. 24. Structural features of matured decayed iridescent feather. A, under light
microscopy, and B–F, SEM.
!
43!
FIG. 25. Structural features of matured white feather. A, under light microscopy, and B–F,
SEM. A, calamus. B–C, plumulaceous barbs and barbules. D–F, calamus surface.
!
44!
FIG. 26. Structural features of matured decayed white feather. A, under light microscopy,
and B–F, SEM. B, plumulaceous barbules and rachis. C–F, calamus.
!
45!
FIG. 27. Structural features of matured horse hair. A, under light microscopy, and B–F,
SEM. A–E, surface. F, cross section.
!
46!
FIG. 28. Structural features of matured turkey reticulate scales. A, under light microscopy,
and B–F, SEM. B–C, harder remnants. D–F, softer remnants.
!
47!
FIG. 29. Structural features of matured turkey scutate scales. A, under light microscopy, and
B–F, SEM.
!
48!
FIG. 30. SEM images of structural features of highly matured feather ‘goo’. A–C, ‘goo’
produced from dark feathers. D–F, ‘goo’ produced from white feathers. A, D, ‘goo’ can be
seen to have extruded onto the surface of the capsule.
!
49!
3.2 Water solubility test
Neither the ‘goo’ derived from the dark nor the white feathers appears to be water
soluble. Observing spots of the ‘goo’ adhering to the sides of glass vials prior to, immediately
after, and 20 minutes after the addition of water shows no changes in either coloration or the
shape of spots’ edges (Supporting Information). When observed 7 weeks later, there were
still no apparent changes.
3.3 Compression experiment
In both compression runs, the sediment was well consolidated such that the resulting
column could be manipulated without becoming unconsolidated. It felt similar to the
consistency of chalk.
The compressed non-decayed finch (Fig. 31A–C) showed some crushing of the body,
especially in the trunk, and spreading out of the wings and feathers. However, it retained an
unexpected degree of 3-dimensionality, especially in the head. The feathers in areas where
body fluids did not leak out did not stick to the sediment. Body fluids appeared to have
leached into the sediment in some places as a result of crushing. Most importantly, the
feathers did not show signs of clumping and represented the original morphology prior to
burial to a high degree.
!
50!
FIG. 31. Preparation of the sediment columns derived from the compression experiments. A–
C, the non-decayed bird. D–F, the decayed bird. A, ventral view. B, lateral view of head. C,
posterior view. D, overhead view of column early in preparation (high stratigraphically) with
head still present. E, overhead view of column later in preparation (low stratigraphically)
with head removed. F, lower stratigraphic layer in side view.
!
51!
The compressed decayed finch showed a very high degree of degradation resulting
from bacterial decay (Fig. 31D–F). The head and tail feathers had separated from the body
via decay and floated up through the sediment prior to the water evaporating away in the
column, as discussed in Supporting Information. The head retained a high amount of 3-
dimensionality (Fig. 31D). The beak retained its orange color. The rest of the body was kept
at a lower stratigraphic level and was highly decayed with many small, unidentifiable pieces
of feathers and body dispersed throughout the sediment column. Again, the trunk of the body
appeared to be the most affected by crushing, although a good amount of 3-dimensionality
was retained. The amount of decay made it difficult to discern any anatomical features.
Surprisingly, some of the flight feathers showed the least degradation of any feature at this
lower stratigraphic level despite the fact that they likely remained wet for a longer period of
time than the head and tail feathers that had floated to the top of the column – allowing more
time for bacterial decay (Fig. 31F). The feathers at the lower stratigraphic level do appear
slightly discolored, however (brownish instead of grey).
3.4 Chemical analysis
Table 2 shows the compounds identified from the total ion chromatograms (TIC) in
the samples that underwent Py/GC/MS. TIC themselves are presented in Supporting
Information. Compounds could not be confidently identified for every peak of the TIC.
!
52!
Sample
(Abbreviation)
Non-Matured
Matured
250
°
C, 250 bars, 24 hours
White feathers
– matured into
‘goo’ (GW)
NA
!Toluene
!Amides
-!Butanamide
-!3-Methylbutanamide
-!Pentanamide,4-methyl-
!N-[2-Hydroxyethyl]succinimide
!Pyrrole-related compounds
-!5,10-Diethoxy-2,3,7,8-tetrahydro-
1H,6H-dipyrrolo[1,2-a;1’,2’-
d]pyrazine
!Cholestenes
-!17-(1,5-Dimethylhexyl)-10,13-
dimethyl-
4,5,6,7,8,9,10,11,12,13,14,15,16,17-
tetradecahydro-1H-
cyclopenta[a]phenanthrene
-!Cholesta-3,5-diene
-!17-(1,5-Dimethylhexyl)-10,13-
dimethyl-
2,3,4,7,8,9,10,11,12,13,14,15,16,17-
tetradecahydro-1H-
cyclopenta[a]phenanthren-3-ol
!Sterols
-!Cholestanol
Dark feathers
– matured into
‘goo’ (GD)
NA
!Amides
-!Propanamide
-!Butanamide
-!3-Methylbutanamide
-!Pentanamide,4-methyl-
!Pyrrole-related compounds
-!Pyrrolidine,1-acetyl-
-!5,10-Diethoxy-2,3,7,8-tetrahydro-
1H,6H-dipyrrolo[1,2-a;1’,2’-
d]pyrazine
!Sterols
100
°
C, 250 bars, 24 hours
Feathers –
iridescent (FI)
!Toluene
!Phenols
-!Methylphenol
!Pyrrole-related
compounds
-!Pyrollopyrazine
!Cholestenes
!Toluene
!Phenols
-!Methylphenol
!Pyrrole-related compounds
-!Pyrollopyrazine
!Phthalate (Contaminant?)
!Sterols
Decayed
iridescent
feathers (DFI)
!Toluene
!Phenols
-!Methylphenol
!Toluene
!Phenols
-!Phenol
!
53!
!Pyrrole-related
compounds
-!Pyrollopyrazine
-!Methylphenol
!Benzyl nitrile
!Pyrrole-related compounds?
-!Pyrollopyrazine
!Amides
!Cholestenes
-!Cholest-3-ene
!Sterols
Feathers –
white (FW)
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Benzyl nitrile
!Pyrrole-related
compounds
-!Pyrollopyrazine
!Sterols
-!Epicholestanol
!Toluene
!Pyrrole-related compounds
-!Pyrrole
-!Pyrollopyrazines
!Styrene
!Phenols
-!Phenol
-!Methylphenol
-!Trimethyl methoxyphenol
!Benzyl nitrile
!Amides
!Sterols
Decayed
white feathers
(DFW)
!Toluene
!Phenols
-!Methylphenol
!Pyrrole-related
compounds
-!Pyrollopyrazine
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Benzyl nitrile
!Pyrrole-related compounds?
-!Pyrollopyrazine?
!Sterols
Feathers –
black (FB)
!Toluene
!Pyrrole-related
compounds
-!Pyrrolo[1,2-
a]pyrazine-1,4-
dione,hexahydro-
-!Pyrrolo[1,2-
a]pyrazine-1,4-
dione,hexahydro-3-
(phenylmethyl)-
!Sterols
-!Long chain sterol?
!Toluene
!Phenols
-!Methylphenol
-!Trimethyl methoxyphenol
!Benzyl nitrile
!Pyrrole-related compounds
-!Pyrollopyrazine
-!Pyrollopyrazine phenylmethyl
!Sterols
-!Long chain sterols?
Decayed black
feathers
(DFB)
!Toluene
!Phenols
-!Methylphenol
!Indoles
-!Indole
!Pyrrole-related
compounds?
-!Pyrollopyrazine?
!Toluene
!Phenols
-!Methylphenol
!Benzyl nitrile
!Indoles
-!Methylindole
!Pyrrole-related compounds?
-!Pyrollopyrazine?
!
54!
Avian scutate
scales (ST)
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Benzyl nitrile
!Palmitic acid
!Stearic acid
!Amides
-!Octadecanamide
!Pyrrole-related
compounds
-!Pyrollopyrazine
phenylmethyl
!Sterols
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Pyrrole-related compounds
-!Pyrollopyrazine
-!Dipyrollopyrazine
!Amides
-!Octadecanamide
!Sterols
Avian
reticulate
scales (RT)
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indoles
-!Indole
!Pyrrole-related
compounds
-!Pyrollopyrazine
-!Dipyrollopyrazine
!Sterols
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Pyrrole-related compounds
-!Dipyrollopyrazine
!Amides?
!Sterols
Turkey beard
– adult (BA)
!Toluene
!Phenols
-!Phenol
-!4-Methylphenol
!Benzyl nitrile
!Benzenepropanenitrile
!Indoles
-!Indole
-!Methylindole
!Pyrrole-related
compounds
-!Pyrollopyrazine
!Sterols?
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indolizine?
!Indoles
-!Indole?
-!Methylindole
!Pyrrole-related compounds?
-!Pyrollopyrazine?
!Cholestenes
-!Cholestadiene
!Sterols
-!Cholesterol
Turkey beard
– juvenile (BJ)
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indoles
-!Indole
!Pyrrole-related
compounds?
-!Pyrollopyrazine?
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indoles
-!Indole
-!Methylindole
!Pyrrole-related compounds?
-!Pyrollopyrazine?
!
55!
!Amides?
!Cholestenes
-!Cholestadiene
!Sterols
-!Cholesterol
!Amides?
!Sterols?
-!Cholesterol?
Crocodilian
scale – black
(CB)
!Pyrrole-related
compounds
-!Pyrrole
-!Methylpyrrole
-!Pyrollopyrazine
-!Dipyrollopyrazine
!Toluene
!Sterols
-!Cholesterol
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indoles
-!Indole
-!Methylindole
!Pyrrole-related compounds
-!Pyrollopyrazine
!Sterols?
-!Cholesterol?
Crocodilian
scale – white
(CW)
!Toluene
!Pyrrole-related
compounds
-!Pyrrole?
-!Methylpyrrole
-!Pyrollopyrazine?
-!5,10-Diethoxy-
2,3,7,8-tetrahydro-
1H,6H-
dipyrrolo[1,2-
a;1’,2’-d]pyrazine?
!Benzyl nitrile
!Amides?
!Sterols?
-!Cholesterol?
!Methyline chloride
!Toluene
!Pyrolle-related compounds
-!Pyrrole?
-!Methylpyrrole
-!Pyrollopyrazine
!Phenols
-!Methylphenol
!Benzyl nitrile
!Indoles
-!Methylindole
!Sterols?
-!Cholesterol?
Mammalian
hair (H)
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Benzyl nitrile
!Indoles
-!Indole
-!Methylindole
!Pyrrole-related
compounds
-!Pyrollopyrazine
-!Dipyrollopyrazine
!Sterols
!Toluene
!Phenols
-!Phenol
-!Methylphenol
!Indoles
-!Indole
-!Methylindole
!Pentenoic acid-amino phenyl?
!Pyrrole-related compounds
-!Pyrollopyrazine
-!Dipyrollopyrazine
!Sterols
Table 2. Compounds identified from the TIC.
!
56!
Fresh samples. Similar pyrolysis breakdown products were observed in the various
fresh tissues, with toluene, phenols (sometimes methylated), indoles (sometimes methylated),
pyrrole-related compounds (especially pyrollopyrazines and sometimes dipyrollopyrazines),
and sterols common. Pyrrole and methylpyrrole were other pyrrole-related compounds
identified. Benzyl nitrile was also present in several fresh samples. Amides were confidently
identified in one sample, the fresh scutate scales, and potentially found in other samples.
Some more unusual compounds identified include palmitic acid, stearic acid, and
pyrollopyrazine phenylmethyl in the fresh scutate scales, cholestenes in the fresh iridescent
feathers, and benzenepropanenitrile in the fresh adult turkey beard. No major differences are
apparent between melanized and non-melanized tissues, such as compounds present in
melanized tissues clearly lacking in non-melanized tissues.
Decayed samples. Similar pyrolysis breakdown products were observed between the
various non-matured decayed feathers, with toluene, phenols (methylated), indoles, and
pyrrole-related compounds (i.e., pyrollopyrazines) common. The compounds in the non-
matured decayed feathers are very similar to those seen in fresh feathers, although larger
compounds like sterols are lacking. However, sterols are seen in the matured decayed
feathers (see below), so differences between fresh and non-matured decayed feathers are not
clear. Furthermore, no differences are apparent between the melanized and non-melanized
non-matured decayed feathers.
Matured samples. The samples matured at 100°C/250 bars for 24 hours showed
similar pyrolysis breakdown products as those seen in the non-matured fresh and decayed
samples. Toluene, phenols (sometimes methylated), indoles (sometimes methylated), pyrrole-
related compounds (especially pyrollopyrazines and sometimes dipyrollopyrazines), and
!
57!
sterols were common. Pyrrole and methylpyrrole were other pyrrole-related compounds
identified. Benzyl nitrile and amides were also found in a number of samples. Cholestenes
were found in matured decayed iridescent feather and matured adult turkey beard. More
unusual compounds identified include phthalate in the matured iridescent feather (likely a
contaminant), styrene in the matured white feather, trimethyl methoxyphenol in the matured
white and black feathers, pyrollopyrazine phenylmethyl in the matured black feather, and a
possible occurrence of pentenoic acid-amino phenyl in the matured horse hair. Differences
between the fresh and matured (100°C/250 bars for 24 hours) samples were not clear. Among
these moderately matured samples, differences between melanized and non-melanized were
also not apparent. Similarly, no obvious differences can be seen between the moderately
matured samples derived from fresh feathers and matured samples derived from decayed
feathers.
The feathers highly matured (250°C/250 bars for 24 hours) into a ‘goo’ seem to have
the most distinct chemical compounds and TIC of any of the samples studied. They have
many of the same categories of compounds as the other samples (i.e., toluene, amides,
pyrrole-related compounds, cholestenes, and sterols), however, many of the specific
compounds identified are unique. The amides are of unique varieties – butanamides and
pentanamides (sometimes methylated) – while the amides that could be identified in the other
samples were octadecanamide. One of the pyrrole-related compounds in the ‘goo’ samples
was diethoxy-tetrahydro-dipyrollopyrazine, which was not confidently identified in any other
samples. The ‘goo’ derived from white feathers contained a diverse group of cholestenes,
some of which were unique from the rest of the samples. Other interesting compounds
identified were n-[2-hydroxyethyl]succinimide in the ‘goo’ derived from white feathers and
pyrrolidine,1-acetyl- in the ‘goo’ derived from dark feathers. Compared to the other samples,
differences between the compounds identified in the melanized and non-melanized ‘goo’
!
58!
were more dramatic. For example, the non-melanized ‘goo’ contains cholestenes while the
melanized ‘goo’ contains only sterols. However, differences between the ‘goo’ derived from
white and dark feathers should probably not be over-interpreted as they might reflect inability
to confidently identify certain peaks and the small sample size. This would make sense given
the inability to distinguish melanized and non-melanized keratins in the less matured
samples.
4. DISCUSSION
4.1 Structural analysis
Fresh samples. The morphology and ultrastructure of the various fresh keratinous
structures match previous descriptions in the literature, as expected. However, there are some
interesting comparisons.
The bristles of the turkey beards have a habit of fraying or splitting at their ends when
cut, unlike feathers and horse hair. This might be due to unique keratin makeup of bristles
(feather-type and avian scale-type φ-keratins) or their internal structure – irregular cross-
sectional shape and often, but not always, solid (unlike feathers) with more organized internal
keratin than horse hair. It was also interesting that the flaking on the outer surface of the
bristle was more reminiscent to the α-keratin horse hair than the φ-keratin in feathers. There
seem to be significant changes in the bristles due to ontogeny, with juvenile bristles being
more irregular in cross-sectional shape than adult bristles. Cross sections of the matured adult
bristles show that some have hollow centers, as has been previously reported. It appears that
cross section morphology is distinct between feathers, turkey beards, scales, and mammalian
hair. Fossil integumentary filaments in dinosaurs have been described as hollow in different
!
59!
instances using conflicting evidence – some citing dark banding on the edges (Chen et al.
1998) and others citing dark banding in the center (Mayr et al. 2002). Given the implications
for how we interpret growth in these integumentary structures, morphology of the cross
sections of primitive integumentary structures could be useful if this information can survive
fossilization.
Feather barbules can show a good amount of variation, and most intriguingly, can
have unique morphology in an iridescent portion of a feather. Plumulaceous barbule nodes in
this study either had ring-like morphology or had a multi-pronged morphology. Some rachis
cross sections lack columnar organization of the outer cortex and cross-sectional shape can
vary in irregularity.
Decayed samples. The most surprising result from the feather decay experiment was
that melanized feathers (black and iridescent) decayed much faster than the non-melanized
feathers (white). In fact, the white feathers were most similar to the control feathers in terms
of their degradation. The overall morphology of the white feathers was preserved and even
the ultrastructural features of the keratin appear little affected by decay contrary to previous
studies (Gunderson et al. 2008), although the hair-like structures on the surface of one portion
of the rachis are peculiar, and may represent early stages of keratin decay. It is possible that,
regardless of the effect of melanin on microbial decay resistance, other factors might be more
important in feather decay such as the structure of the feather or the amount of calcium
phosphate deposition within the keratin. More work is needed to determine the relative
importance of feather features in providing decay resistance.
The contribution of melanin to decay resistance cannot be outright rejected since the
surviving components of the decayed black feather are mostly melanized. The lighter
portions fringing the barbs and the rachis (which was not always fully pigmented) seem to be
!
60!
less represented in the sample. However, a similar pattern of rachis loss and barb/barbule
retention was seen in the decayed iridescent feather, and these were melanized throughout
(including their rachises) implying that the structure of different feather parts has a greater
contribution to decay resistance than melanin.
The connection between the barbules and barbs was preserved more readily than the
connection between the barbs and rachis in the decayed black and iridescent feathers, and in
the decayed black feather, these remnants of barbs and barbules also show semi-fusion into a
mass. Sausage-shaped eumelanosomes from the decayed black and iridescent feathers are
apparent. This is likely due to breakdown of the rachis keratin, which could be seen fraying in
the sample of decayed iridescent feathers. This breakdown of the rachis also reveals the
medulloid cells of the pith, which sometimes appear collapsed (possibly as a result of the
vacuum conditions during SEM). The decayed black and iridescent feathers also both show
keratin surfaces with a woven texture, rather than simply flaking apart into smaller pieces.
Matured samples. Even at relatively low temperatures (100°C), maturation had major
effects on the morphology and ultrastructure of the integumentary appendages.
Both juvenile and adult turkey bristles show surface degradation and cross-sectional
compression. However, the juvenile bristles show extreme curling while the adult bristle
exhibit simple wrinkling or creasing. The flaking of the epidermis is similar in both
ontogenetic stages.
The matured black feather retains the original color range (black with white fringe)
better than did the decayed black feather. Like the decayed black feather, the matured black
feather resulted in a mass of barbs and barbules. While these were semi-fused with highly
degraded surfaces in the decayed black feather, the matured black feather showed no fusion
and the barbs and barbules were kinked rather than degrading along the surface of the keratin.
!
61!
Compared to the decayed black feather, the matured decayed black feather shows
great fusion of the barbs and barbules such that only a few can be identified among the solid
mass. Some of the flaking seen in the keratin surface of the decayed black feather appears to
be intensified into the granulated texture of some regions on the matured decayed black
feather.
Like the decayed iridescent feather, the matured iridescent feather consists of a mass
of barbs and barbules. Both retain some degree of iridescence, although this is reduced in the
matured sample. Similar to the matured vs. decayed black feather, the matured iridescent
feather shows a loss of overall feather morphology, but the keratin surface does not appear as
degraded as that of the decayed iridescent feather.
The matured decayed iridescent feather has turned into a pellet similar to that of the
matured decayed black feather. Most of the features still present in the decayed iridescent
feather are lost in the matured decayed iridescent feather, although barbules attached to
degraded barbs can still be made out. Unlike the woven texture seen in the decayed iridescent
feather, the keratin of what was presumably once the rachis is peeling into large strips or
strands in the matured decayed iridescent feather.
The matured white feather shows degradation of the overall feather morphology and
also shows the peeling off of large strips or strands on its rachis/calamus. It is hard to
compare maturation to decay in this instance since the white feather decayed at such a slow
rate in the experiment, resulting in little signs of degradation.
The matured decayed white feather shows similar loss of overall feather morphology
to the matured white feather, but effect on the rachis/calamus keratin is more extreme. Instead
of simply peeling off into strips or strands, an organized mesh of filaments is exposed. A
previous study used microbial degradation of the rachis keratin to better study the
filamentous hierarchy within. The reported filaments, similar to those observed here, are the
!
62!
thickest of any keratin filament (6 µm), extend along the axis of the rachis with a small outer
circumferential layer, and have thickened nodes staggering between adjacent filaments along
two and three dimensions (Lingham-Soliar et al. 2010). In that study and here, filaments
appear almost identical to plumulaceous barbules – a fact that these researchers cite as
evidence for an evo-devo model of feather evolution in which barbs fuse together during
development to form the rachis, as proposed by Prum & Brush (2002). While the results here
support the existence of these filamentous subunits of the rachis, it is interesting that the other
researchers observed them through decay alone. Here, they were only apparent in the matured
decayed white feather. The peeling off of strands or strips in the matured decayed iridescent
and matured white feathers as well as the woven texture seen in the decayed black and
decayed iridescent feathers might represent early stages of degradation in which these
filamentous subunits are not fully exposed.
Although the decay experiments found the more melanized feathers to degrade faster,
maturation seemed to show the opposite trend. The greatest morphological and ultrastructural
changes as well as the greatest amount of rachis/calamus keratin degradation as a result of
maturation occurs in the white feathers. While other factors might be more important in
conferring decay resistance to feathers than melanin, melanin might be important in reducing
the rate of degradation due to maturation.
In all three feather colors, maturation did not produce identical effects as decay,
particularly at the ultrastructural level of the keratin surface. This is reflective of the activity
of bacteria degrading the keratin surface during decay, while the intense temperature and
pressure of maturation can affect the keratin structure as a whole, rather than just at its
surface. However, maturation and decay can both break down the overall structure of the
feather. Of all samples, the greatest amount of degradation occurred in the matured decayed
feathers, as might be expected.
!
63!
Differences in degradation patterns between matured filaments might relate to their
keratin type: helical curling and kinking in the matured horse hair (α-keratin), curling of the
matured juvenile turkey bristles (both feather-type and avian scale-type φ-keratins), wrinkling
and creasing of the matured adult turkey bristles (both feather-type and avian scale-type φ-
keratins), and the kinking of the matured black feather (feather-type φ-keratins) were
observed. These differences could be useful only if they left some sort of signature in fossils
(perhaps by affecting melanosome positioning).
Both predominantly black and predominantly white crocodile scales show cracking of
their outer layer, splitting between the sub-layers, and retention of the rippled ultrastructural
surface. While the predominantly white scale was slightly discolored, the black retained its
original range of coloration. The internal layer of the scales matured into a soft substance.
Like the matured crocodile scales, the matured reticulate turkey scales show harder
and softer remnants. The softer remnants, likely derived from the inner layer of the scales,
show interesting features that come in a range of morphologies. The extreme morphology
looks like a concave disk bulging out of the surface of the organic material. Superficially,
they look like human erythrocytes. However, some are irregular in shape and many
intermediates are present between the extreme morphology and the folding or pock-marking
of the surface. There are also some bulges that are simple spheres. Thus, they likely represent
a taphonomic artifact of maturation in which the organic material folds outward or gasses
escape from the interior. This is important as similarly shaped features have been interpreted
as fossilized erythrocytes in Mesozoic dinosaur bones (Bertazzo et al. 2015). According to
the authors, bird erythrocytes are 350-650% larger than the average size of their proposed
erythrocytes. Not only are the proposed dinosaur erythrocytes potentially taphonomic
artifacts from organic material, they are also more similar in shape to human rather than oval
bird or crocodilian erythrocytes (Claver & Quaglia 2009). Their putative fossil erythrocytes
!
64!
are, therefore, rejected here. This is in agreement with anthropological study of human blood
in bones showing that well-defined erythrocytes are morphologically unidentifiable through
histological analysis after a week of decomposition. Their breakdown products can only be
detected using immunohistochemical analysis for a few years after death.
Immunohistochemical analysis cannot detect degraded erythrocytes in bones of archeological
age (Cappella et al. 2015), suggesting protein breakdown and corresponding loss of the
antigen binding site.
The matured scutate turkey scale does not show as many features as did the matured
reticulate scales and the harder and softer portions are more difficult to distinguish in the
sample. All of the various scale types in this study matured into pliable masses, although the
original outer and inner layers of the scales were more recognizable after maturation in the
scales made of non-featherlike β-keratins than those made of φ-keratins (the scutate scales).
The rippled ultrastructural surface of the outer layer of fresh and matured crocodile
and fresh turkey reticulate scales is more complex than the relatively smooth keratin surface
of feathers. These scales are made of non-featherlike β-keratin. The scutate scales show a
relatively smooth ultrastructural surface, more like feathers (both consisting of φ-keratin).
Some might take this ultrastructural similarity as further evidence that scutate scales evolved
via modification of leg feathers (Zheng et al. 2013). Even if the rippled texture is an accurate
indicator of keratin type, and it survives moderate heat and pressures, the question remains as
to whether it can survive the extreme conditions of fossilization.
The ‘goo’ produced from highly matured feathers shows complete degradation of the
keratin ultrastructure. Melanosomes cannot be recognized in the ‘goo’ derived from dark
feathers, which is surprising considering their widespread presence in fossils. It is possible
that melanosomes did survive maturation but became obscured by the ‘goo’.
!
65!
More importantly, the presence of this ‘goo’ contradicts previous work in which
feathers exposed to comparably high temperatures and pressures (200°C/250 bars and
250°C/250 bars) for 24 hours retained the majority of their original morphology. Repeating
the experiments detailed by McNamara et al. (2013) failed to reproduce their results.
Furthermore, retention of feather morphology was not observed here even after a lower
temperature maturation of white feathers (200°C, likely lower than temperatures during
fossilization), which also produced a ‘goo’ that leaked out of the tube (Supporting
Information). Repeating the exact temperatures, pressure, and duration they reported (and in
the maturation runs where the samples were lost, using an Ar gas, rather than a wet,
autoclave) failed to produce results even vaguely similar to those they report. In their
supplemental material, however, McNamara et al. state, “Experiments were undertaken at:
200°C, 1 bar; 200°C, 117 bar; 200°C, 250 bar; 200°C, 500 bar; 270°C, 500 bar; 25°C, 500
bar, each for 24 hours. 1 hour experiments were run at 200°C, 250 bar and 250°C, 250 bar”.
Possibly, the results they present were actually from maturation experiments run for 1 hour
rather than 24 hours, and the main text of the paper reports an incorrect procedure.
4.2 Water solubility test
The ‘goo’ produced from the highly matured feathers is not water soluble, so
dissolution is not a method by which keratin can be lost from the fossil. However, this does
not mean that the ‘goo’ is not volatile and cannot be lost in other ways.
!
66!
4.3 Compression experiment
The results from the compressed un-decayed finch show that compression after sub-
aqueous burial does not greatly alter the appearance of feather morphology. The results of
Foth (2012) derive from unrealistic simulation of taphonomic conditions. The more realistic
taphonomic simulation reported here shows that feather morphology is likely preserved
through burial and compression and that complex feathers do not clump to look like proto-
feathers.
The compressed decayed finch provided little information as the decay process had
proceeded further than expected and little of the bird remained. However, the survival of
some flight feathers on the wing suggests that these feathers might be able to resist decay and
degradation longer than other feathers.
Much more 3-dimensionality was preserved in both finches than was expected. This is
because the side walls of the container in which the sediment was compressed exert forces
back onto the sediment during compression. However, the current design (Supporting
Information) does produce a consolidated column that can be easily manipulated. For future
experiments, a second compression step will likely be needed in which this moist,
consolidated sediment column is compressed but allowed to expand outward in order to
replicate the extreme flattening in some fossils. Future trials will likely avoid a decay step
prior to compression as it is difficult to monitor the rate of decay when the finch is buried.
Finally, if the compressed bird and sediment column can be matured under high temperature
and pressure, then it might be possible to produce specimens that are essentially ‘artificial
fossils’. However, an autoclave with a very large sample chamber would be required.
!
67!
4.4 Chemical analysis
The pyrolysis breakdown products generally match those reported from previous
Py/GC/MS studies of keratin (Brebu & Spiridon 2011). These include aromatics like toluene
and styrene, phenols, nitriles, pyrroles, pyrazines, amides, and indoles. Phthalate found in one
of the samples is likely a contaminant as it is a plasticizer (Xiao et al. 2010). Lipids such as
sterols, cholestenes, and fatty acids (e.g., palmitic and stearic acid) observed in this study are
known to derive from keratinizing cells and have been found on feather plumage and wool
fibers (Carruthers and Suntzeff 1953, Bolliger & Varga 1961, Negri et al. 1991, Tomlinson et
al. 2004). The possible presence of pentenoic acid (also known as valeric acid) in the matured
horse hair is not completely unexpected as pentenoic acid has been identified in the grease of
dog hair (Brouwer and Nijkamp 1953).
The Py/GC/MS results here can identify the presence of keratin but cannot easily
distinguish different keratin types (e.g., α-keratin and β-keratin) or keratin from different
taxa.
Py/GC/MS has previously been run on fresh and fossil melanin. Major pyrolysis
products of melanin are pyridine, pyrrole, toluene, C1 pyridine, C1 pyrrole, phenol, 4- and 5-
methylphenol, 2-penylacetonnitrile, C2 phenol, 3-phenylpropanenitrile, indole, C1 indole,
palmitic acid, and stearic acid. The fossil sample showed additional products as a result of
diagenesis: thiophene, C1 thiophene, C2 thiophene, C2 benzene, C3 thiophene, and C3 benzene
(Glass et al. 2012). Many of these major pyrolysis products of melanin (e.g., pyrroles,
toluene, phenols, nitriles, indoles, and fatty acids) are also major pyrolysis products of
keratin. This similarity would explain why Py/GC/MS was unable to distinguish between
melanized and non-melanized keratins in this study.
!
68!
Py/GC/MS was also unable to distinguish between fresh, decayed, moderately
matured (100°C/250 bars for 24 hours), and moderately matured decayed keratins. However,
the highly matured ‘goo’ (250°C/250 bars for 24 hours) produced from feathers presented
intriguing results. The TIC and identified pyrolysis breakdown products from these ‘goo’
samples were the most unique in the study, suggesting that Py/GC/MS is a possible tool for
distinguishing between fresh and highly matured keratin. The phenanthrene found in the
‘goo’ produced from white feathers is interesting as this compound is composed of three
fused benzene rings – benzene being known as a pyrolysis breakdown product of keratin
(Brebu & Spiridon 2011). Also of interest is the succinimide found in the ‘goo’ produced
from white feathers. This compound is known to be linked to protein degradation reactions
(Gieger & Clarke 1987). Specifically, Stephenson & Clarke (1989) described how the
formation of succinimide provides a model for spontaneous protein degradation. They state,
“Nonenzymatic intramolecular reactions can result in the deamidation, isomerization, and
racemization of protein and peptide asparaginyl and aspartyl residues via succinimide
intermediates”. In combination with the loss of all morphological structure in the ‘goo’, these
Py/GC/MS results suggest a breakdown of the keratin protein structure and peptide bonds
(and possibly, further breakdown of the free amino acids) in the highly matured feathers. This
casts serious doubt as to whether keratin protein can be preserved in fossils, especially of
Mesozoic age. Melanosomes and calcium phosphates salts are likely the only remnants of
keratinous structures in fossils.
Manning et al. (2009) found the following products of Py/GC/MS of “mummy”
hadrosaur skin: “n-alkanes/n-alken-1-ene homologues ranging in carbon number from C9 to
C36 with a trimodal distribution of n-alkanes (maxima C11, C15 and C27)”, aldehyde,
“contaminant”, silicon-containing compounds, benzonitrile, benzothiophene, dibenzofuran,
benzene derivatives with 2 or 3 carbons in the alkyl group, and naphthalene derivatives with
!
69!
1 or 2 carbons in the alkyl group. Edwards et al. (2011) used Py/GC/MS of fossil reptile skin
to identify straight chain n-alkane/n-alkenes (carbon chain lengths: 10 and 15) and branched
n-alkane/n-alkenes (carbon chain lengths: 11, 13, 16, 18, and 19). These results show some
pyrolysis breakdown products that might be expected from keratin such as nitriles,
thiophenes, and benzene, but these are also known from Py/GC/MS of fossil melanin. The
group of compounds they identified match with those expected from keratin to a far lesser
extent as those from this study on fresh, decayed, and matured keratin. Additionally, their
results seem to match less well with the highly matured feather ‘goo’ than they do with the
fresher keratin samples. One would expect to have similar pyrolysis breakdown products in
these fossils to those in highly matured keratin. The abundance of aliphatic compounds in
their samples are also reminiscent of reports of aliphatic compounds derived from lipids as a
result of in situ polymerization in some fossils, rather than proteins. These researchers also
used FTIR to identify the presence of amide groups in fossil reptile and “mummy” hadrosaur.
However, the presence of diverse amides in the highly matured feather ‘goo’ shows that
amides present in Py/GC/MS do not necessarily derive from intact keratin proteins. It seems
that these previous studies may have found potential evidence for organic material that
derives from keratin breakdown, but failed to provide convincing evidence for the presence
of intact or original keratin as they suggest.
5. CONCLUSION
Differences in decay and maturation patterns between different keratin types and
structures were observed, but none of the described keratin textures have been confidently
identified in fossils. Contrary to previous claims, keratin does not likely survive in fossils or
leave any obvious signatures, since the high temperature maturation experiments (250°C/250
!
70!
bars for 24 hours) are probably more realistic in terms of fossilization conditions than the
other experimental treatments and resulted in feathers degrading into ‘goo’. Of all proteins, it
seems that collagen, rather than keratin, is the most likely protein to survive over geologic
time, but claims of original collagen protein in Mesozoic fossils (Schweitzer et al. 2007) may
still require reexamination given the thermodynamic instability of peptide bonds and
difficulties of immunohistochemistry. Future improvements to the compression experiments
in this study might be able to produce ‘artificial fossils’ through which diagenetic interactions
between soft tissues and sediment could be observed.
REFERENCES
Alibardi, L. and Toni, M. 2006. Cytochemical, biochemical and molecular aspects of the
process of keratinization in the epidermis of reptilian scales. Progress in
Histochemistry and Cytochemistry, 40, 73–134.
Alibardi, L., Knapp, L. W., and Sawyer, R. H. 2005. Beta-keratin localization in developing
alligator scales and feathers in relation to the development and evolution of feathers.
Journal of Submicroscopic Cytology and Pathology, 38, 175–192.
Allison, P. A. and Briggs, D. E. G. 1993. Exceptional fossil record: Distribution of soft-tissue
preservation through the Phanerozoic. Geology, 21, 527–530.
Barden, H. E., Bergmann, U., Edwards, N. P., Egerton, V. M., Manning, P. L., Perry, S., van
Veelen, A., Wogelius, R. A., and van Dongen, B. E. 2015. Bacteria or melanosomes?
A geochemical analysis of micro-bodies on a tadpole from the Oligocene Enspel
Formation of Germany. Palaeobiodiversity and Palaeoenvironments, 95, 33–45.
Benton, M. J., Zhou, Z., Orr, P. J., Zhang, F., and Kearns, S. L. 2008. The remarkable fossils
from the Early Cretaceous Jehol Biota of China and how they have changed our
!
71!
knowledge of Mesozoic life. Proceedings of the Geologists' Association, 119, 209–
228.
Bergmann, U., Manning, P. L., and Wogelius, R. A. 2012. Chemical mapping of
paleontological and archeological artifacts with synchrotron X-rays. Annual Review of
Analytical Chemistry, 5, 361–389.
Bertazzo, S., Maidment, S. C., Kallepitis, C., Fearn, S., Stevens, M. M., and Xie, H. N. 2015.
Fibres and cellular structures preserved in 75-million-year-old dinosaur specimens.
Nature Communications, 6.
Bolliger, A., and Varga, D. 1961. Feather lipids. Nature, 190, 1125.
Brebu, M., and Spiridon, I. 2011. Thermal degradation of keratin waste. Journal of Analytical
and Applied Pyrolysis, 91, 288–295.
Briggs, D. E. G. 1999. Molecular taphonomy of animal and plant cuticles: selective
preservation and diagenesis. Philosophical Transactions of the Royal Society B:
Biological Sciences, 354, 7–17.
Briggs, D. E. G., and Summons, R. E. 2014. Ancient biomolecules: their origins,
fossilization, and role in revealing the history of life. BioEssays, 36, 482–490.
Brooks, J. D. and Smith, J. W. 1967. The diagenesis of plant lipids during the formation of
coal, petroleum and natural gas—I. Changes in the n-paraffin hydrocarbons.
Geochimica et Cosmochimica Acta, 31, 2389–2397.
— 1969. The diagenesis of plant lipids during the formation of coal, petroleum and natural
gas—II. Coalification and the formation of oil and gas in the Gippsland Basin.
Geochimica et Cosmochimica Acta, 33, 1183–1194.
Brouwer, E., and Nijkamp, H. J. 1953. Occurrence of two valeric acids (β-methylbutyric acid
and α-methylbutyric acid) in the hair grease of the dog. Biochemical Journal, 55, 444.
Brush, A. H. 1978. Feather keratins. Chemical Zoology, 10, 117-139.
!
72!
Cappella, A., Bertoglio, B., Castoldi, E., Maderna, E., Giancamillo, A. D., Domeneghini, C.,
Andreola, S., and Cattaneo, C. 2015. The taphonomy of blood components in
decomposing bone and its relevance to physical anthropology. American Journal of
Physical Anthropology, doi: 10.1002/ajpa.22830.
Carruthers, C., and Suntzeff, V. 1953. Biochemistry and physiology of epidermis.
Physiological Reviews, 33, 229–243.
Chen, P. J., Dong, Z. M., and Zhen, S. N. 1998. An exceptionally well-preserved theropod
dinosaur from the Yixian Formation of China. Nature, 391, 147–152.
Claver, J. A., and Quaglia, A. I. 2009. Comparative morphology, development, and function
of blood cells in nonmammalian vertebrates. Journal of Exotic Pet Medicine, 18, 87–
97.
Edwards, N. P., Barden, H. E., Van Dongen, B. E., Manning, P. L., Larson, P. L., Bergmann,
U., Sellers, W. I., and Wogelius, R. A. 2011. Infrared mapping resolves soft tissue
preservation in 50 million year-old reptile skin. Proceedings of the Royal Society of
London B: Biological Sciences, 278, 3209–3218.
Foth, C. 2012. On the identification of feather structures in stem-line representatives of birds:
evidence from fossils and actuopalaeontology. Paläontologische Zeitschrift, 86, 91–
102.
Fraser, R. D. and Parry, A. D. 2008. Molecular packing in the feather keratin filament.
Journal of Structural Biology, 162, 1–13.
Fraser, R. D. B., MacRae, T. P., and Rogers, G. E. 1972. Keratins- their Composition,
Structure and Biosynthesis, Charles C. Thomas, Springfield, Illinois, US, 304.
Geiger, T., and Clarke, S. 1987. Deamidation, isomerization, and racemization at asparaginyl
and aspartyl residues in peptides. Succinimide-linked reactions that contribute to
protein degradation. Journal of Biological Chemistry, 262, 785–794.
!
73!
Glass, K., Ito, S., Wilby, P. R., Sota, T., Nakamura, A., Bowers, C. R., Vinther, J., Dutta, S.,
Summons, R., Briggs, D. E. G., Wakamatsu, K., and Simon, J. D. 2012. Direct
chemical evidence for eumelanin pigment from the Jurassic period. Proceedings of
the National Academy of Sciences, 109, 10218–10223.
Glass, K., Ito, S., Wilby, P. R., Sota, T., Nakamura, A., Bowers, C. R., Miller, K. E., Dutta,
S., Summons, R. E., Briggs, D. E. G., Wakamatsu, K., and Simon, J. D. 2013. Impact
of diagenesis and maturation on the survival of eumelanin in the fossil record.
Organic Geochemistry, 64, 29–37.
Greenwalt, D. E., Goreva, Y. S., Siljeström, S. M., Rose, T., and Harbach, R. E. 2013.
Hemoglobin-derived porphyrins preserved in a Middle Eocene blood-engorged
mosquito. Proceedings of the National Academy of Sciences, 110, 18496–18500.
Greenwold, M. J. and Sawyer, R. H. 2010. Genomic organization and molecular phylogenies
of the beta (β) keratin multigene family in the chicken (Gallus gallus) and zebra finch
(Taeniopygia guttata): implications for feather evolution. BMC Evolutionary Biology,
10, 148.
Gunderson, A. R., Frame, A. M., Swaddle, J. P., and Forsyth, M. H. 2008. Resistance of
melanized feathers to bacterial degradation: is it really so black and white?. Journal of
Avian Biology, 39, 539–545.
Gupta, N. S., Briggs, D. E. G., and Pancost, R. D. 2006a. Molecular taphonomy of
graptolites. Journal of the Geological Society, 163, 897–900.
Gupta, N. S., Michels, R., Briggs, D. E. G., Evershed, R. P., and Pancost, R. D. 2006b. The
organic preservation of fossil arthropods: an experimental study. Proceedings of the
Royal Society B: Biological Sciences, 273, 2777–2783.
Gupta, N. S., Briggs, D. E. G., Collinson, M. E., Evershed, R. P., Michels, R., Jack, K. S.,
and Pancost, R. D. 2007a. Evidence for the in situ polymerisation of labile aliphatic
!
74!
organic compounds during the preservation of fossil leaves: implications for organic
matter preservation. Organic Geochemistry, 38, 499–522.
Gupta, N. S., Briggs, D. E. G., Collinson, M. E., Evershed, R. P., Michels, R., and Pancost,
R. D. 2007b. Molecular preservation of plant and insect cuticles from the Oligocene
Enspel Formation, Germany: evidence against derivation of aliphatic polymer from
sediment. Organic Geochemistry, 38, 404–418.
Gupta, N. S., Michels, R., Briggs, D. E. G., Collinson, M. E., Evershed, R. P., and Pancost,
R. D. 2007c. Experimental evidence for the formation of geomacromolecules from
plant leaf lipids. Organic Geochemistry, 38, 28–36.
Gupta, N. S., Cambra-Moo, O., Briggs, D. E. G., Love, G. D., Fregenal-Martinez, M. A., and
Summons, R. E. 2008. Molecular taphonomy of macrofossils from the Cretaceous Las
Hoyas Formation, Spain. Cretaceous Research, 29, 1–8.
Gupta, N. S., Cody, G. D., Tetlie, O. E., Briggs, D. E. G., and Summons, R. E. 2009. Rapid
incorporation of lipids into macromolecules during experimental decay of
invertebrates: Initiation of geopolymer formation. Organic Geochemistry, 40, 589–
594.
Gupta, R. and Ramnani, P. 2006. Microbial keratinases and their prospective applications: an
overview. Applied Microbiology and Biotechnology, 70, 21–33.
Haddar, H. O., Zaghloul, T. I., and Saeed, H. M. 2009. Biodegradation of native feather
keratin by Bacillus subtilis recombinant strains. Biodegradation, 20, 687–694.
Hill, P., Brantley, H., and Van Dyke, M. 2010. Some properties of keratin biomaterials:
kerateines. Biomaterials, 31, 585–593.
Hofreiter, M., Collins, M., and Stewart, J. R. 2012. Ancient biomolecules in Quaternary
palaeoecology. Quaternary Science Reviews, 33, 1–13.
!
75!
Korniłłowicz-Kowalska, T. and Bohacz, J. 2011. Biodegradation of keratin waste: theory and
practical aspects. Waste Management, 31, 1689–1701.
Lingham-Soliar, T., Bonser, R. H., and Wesley-Smith, J. 2010. Selective biodegradation of
keratin matrix in feather rachis reveals classic bioengineering. Proceedings of the
Royal Society of London B: Biological Sciences, 277, 1161–1168.
Manning, P. L., Morris, P. M., McMahon, A., Jones, E., Gize, A., Macquaker, J. H. S., Wolff,
G., Thompson, A., Marshall, J., Taylor, K. G., Lyson, T., Gaskell, S., Reamtong, O.,
Sellers, W. I., van Dongen, B, E., Buckley, M., and Wogelius, R. A. 2009.
Mineralized soft-tissue structure and chemistry in a mummified hadrosaur from the
Hell Creek Formation, North Dakota (USA). Proceedings of the Royal Society of
London B: Biological Sciences, 276, 3429–3437.
Mayr, G., Peters, S. D., Plodowski, G., and Vogel, O. 2002. Bristle-like integumentary
structures at the tail of the horned dinosaur Psittacosaurus. Naturwissenschaften, 89,
361–365.
McNamara, M.E., Briggs, D. E. G., Orr, P. J., Field, D. J., and Wang, Z. 2013. Experimental
maturation of feathers: implications for reconstructions of fossil feather colour.
Biology Letters, 9, 20130184.
Murphy, M. E., and King, J. R. 1982. Amino acid composition of the plumage of the white-
crowned sparrow. Condor, 435-438.
Nakai, S., and Modler, H. W. 1996. Food proteins: properties and characterization. John
Wiley & Sons, Hoboken, New Jersey, US, 544.
Negri, A. P., Cornell, H. J., and Rivett, D. E. 1991. The nature of covalently bound fatty acids
in wool fibres. Crop and Pasture Science, 42, 1285–1292.
O'Connor, S., Solazzo, C., and Collins, M. 2014. Advances in identifying archaeological
traces of horn and other keratinous hard tissues. Studies in Conservation.
!
76!
Orlando, L., Ginolhac, A., Zhang, G., Froese, D., Albrechtsen, A., Stiller, M., Schubert, M.,
Cappellini, E., Peterson, B., Moltke, I., Johnson, P. L. F., Fumagalli, M., Vilstrup, J.
T., Raghavan, M., Korneliussen, T., Malaspinas, A-S., Vogt, J., Szklarczyk, D.,
Kelstrup, C. D., Vinther, J., Dolocan, A., Stenderup, J., Velazquez, A. M. V., Cahill,
J., Rasmussen, M., Wang, X., Min, J., Zazula, G. D., Seguin-Orlando, A., Mortensen,
C., Magnussen, K., Thompson, J. F., Weinstock, J., Gregersen, K., Røed, K. H.,
Eisenmann, V., Rubin, C. J., Miller, D. C., Antczak, D. F., Bertelsen, M. F., Brunak,
S., Al-Rasheid, K. A. S., Ryder, O., Andersson, L., Mundy, J., Krogh, A., Gilbert, M.
T. P., Kjær, K., Sicheritz-Ponten, T., Jensen, L. J., Olsen, J. V., Hofreiter, M.,
Nielsen, R., Shapiro, B., Wang, J., and Willerslev, E. 2013. Recalibrating Equus
evolution using the genome sequence of an early Middle Pleistocene horse. Nature
499, 74–78.
Pauling, L., Corey, R. B., and Branson, H. R. 1951. The structure of proteins: two hydrogen-
bonded helical configurations of the polypeptide chain. Proceedings of the National
Academy of Sciences, 37, 205–211.
Prum, R. O. and Brush, A. H. 2002. The evolutionary origin and diversification of feathers.
The Quarterly Review of Biology, 77, 261–295.
Sawyer, R.H., Glenn, T., French, J. O., Mays, B., Shames, R. B., Barnes, G. L., Rhodes, W.,
and Ishikawa, Y. 2000. The expression of beta (β) keratins in the epidermal
appendages of reptiles and birds. American Zoologist, 40, 530–539.
Sawyer, R. H., Washington, L. D., Salvatore, B. A., Glenn, T. C., and Knapp, L. W. 2003.
Origin of Archosaurian integumentary appendages: The bristles of the wild turkey
beard express feather-type β keratins. Journal of Experimental Zoology Part B
Molecular and Developmental Evolution, 297, 27–34.
Schorger, A. W. 1957. The beard of the wild turkey. The Auk, 74, 441–446.
!
77!
Schweitzer, M. H., Watt, J. A., Avci, R., Knapp, L., Chiappe, L., Norell, M., and Marshall,
M. 1999. Beta-keratin specific immunological reactivity in feather-like structures of
the cretaceous alvarezsaurid, Shuvuuia deserti. Journal of Experimental Zoology, 285,
146–57.
Schweitzer, M. H., Suo, Z., Avci, R., Asara, J. M., Allen, M. A., Arce, F. T., and Horner, J.
R. 2007. Analyses of soft tissue from Tyrannosaurus rex suggest the presence of
protein. Science, 316, 277–280.
Shames, R.B., Knapp, L. W., Carver, W. E., Washington, L. D., and Sawyer, R. H. 1989.
Keratinization of the outer surface of the avian scutate scale: interrelationship of alpha
and beta keratin filaments in a cornifying tissue. Cell and Tissue Research, 257, 85–
92.
Solazzo, C., Wadsley, M., Dyer, J. M., Clerens, S., Collins, M. J., and Plowman, J. 2013a.
Characterisation of novel α!keratin peptide markers for species identification in
keratinous tissues using mass spectrometry. Rapid Communications in Mass
Spectrometry, 27, 2685–2698.
Solazzo, C., Wilson, J., Dyer, J. M., Clerens, S., Plowman, J. E., von Holstein, I., Rogers, P.
W., Peacock, E. E., and Collins, M. J. 2013b. Modeling deamidation in sheep α-
keratin peptides and application to archeological wool textiles. Analytical Chemistry,
86, 567–575.
Solazzo, C., Rogers, P. W., Weber, L., Beaubien, H. F., Wilson, J., and Collins, M. 2014.
Species identification by peptide mass fingerprinting (PMF) in fibre products
preserved by association with copper-alloy artefacts. Journal of Archaeological
Science, 49, 524–535.
Steinert, P. M. and Freedberg, I. M. 1991. Molecular and cell biology of keratins. In
Physiology, Biochemistry and Molecular Biology of the Skin (Vol 1), (Goldsmith, L.
!
78!
A., ed.), Oxford University Press, Oxford, UK, 113–147.
Stephenson, R. C., and Clarke, S. 1989. Succinimide formation from aspartyl and asparaginyl
peptides as a model for the spontaneous degradation of proteins. Journal of Biological
Chemistry, 264, 6164–6170.
Tomlinson, D. J., Mülling, C. H., and Fakler, T. M. 2004. Invited review: formation of
keratins in the bovine claw: roles of hormones, minerals, and vitamins in functional
claw integrity. Journal of Dairy Science, 87, 797–809.
True, L. D. 2008. Quality control in molecular immunohistochemistry. Histochemistry and
Cell Biology, 130, 473–480.
Vinther, J. 2015. A guide to the field of palaeo colour. BioEssays, 37, 643–656.
Vinther, J., Briggs, D. E. G., Prum, R. O., and Saranathan, V. 2008. The colour of fossil
feathers. Biology Letters, 4, 522–525.
Wang, S. Y., Cappellini, E., and Zhang, H. Y. 2012. Why collagens best survived in fossils?
Clues from amino acid thermal stability. Biochemical and Biophysical Research
Communications, 422, 5–7.
Williams, C. M., Richter, C. S., Mackenzie, J. M., and Shih, J. C. 1990. Isolation,
identification, and characterization of a feather-degrading bacterium. Applied and
Environmental Microbiology, 56, 1509–1515.
Wyld, J. A., and Brush, A. H. 1979. The molecular heterogeneity and diversity of reptilian
keratins. Journal of Molecular Evolution, 12, 331–347.
Wyld, J. A., and Brush, A. H. 1983. Keratin diversity in the reptilian epidermis. Journal of
Experimental Zoology, 225, 387–396.
Xiao, N. Y., Lu, X. C., Guo, Q. B., Tan, G. L., and Yang, H. Q. 2010. Research Progress of
Phthalate Plasticizer Migration in Plastic Food Packaging. Packaging Engineering,
11, 036.
!
79!
Xu, X. and Guo, Y. 2009. The origin and early evolution of feathers: insights from recent
paleontological and neontological data. Vertebrata PalAsiatica, 47, 311–329.
Zheng, X., Zhou, Z., Wang, X., Zhang, F., Zhang, X., Wang, Y., Wei, G., Wang, S., and Xu,
X. 2013. Hind wings in basal birds and the evolution of leg feathers. Science, 339,
1309–1312.
!
A!
SUPPORTING INFORMATION
FURTHER DESCRIPTION OF MATERIALS AND METHODS
Fresh samples
Samples were gathered from a range of extant Archosaurian integumentary structures,
including feathers, scales, and bristles. Mammalian hair was also included as an outgroup.
Feathers were collected from a deceased male bronze turkey (Meleagris gallopavo)
and male light Sussex chicken (Gallus gallus) ordered from UK farms. Feathers were
plucked from the body and sorted according to body region (back, belly, neck, wing coverts,
primaries, secondaries, tail, legs) and stored in a laboratory freezer to prevent any decay until
experimentation. The light Sussex chicken was selected for its black and white plumage,
acting as a control when examining taphonomic differences between pigmented and non-
pigmented feathers. It is mostly white, but has black feathers on the wings and tail. Many
feathers on the neck are black with a white fringe. The male turkey was selected for its beard,
as discussed below. The feathers from the turkey are heavily pigmented and some are
iridescent. This study focused on white, black, and iridescent feathers.
Legs from the turkey were dissected off and placed in the freezer. Scutate and
reticulate scales were later dissected off of the legs.
The male bronze turkey provided was younger than expected – it only had very short
bristles. The beard, along with the epidermal outgrowth that supports the bristles, was
dissected and placed in the freezer. To supplement this, dried beards of wild turkeys (same
species as the domestic turkey) from older individuals with longer bristles were shipped from
the US. Although their dried nature might change degradation processes in these samples,
!
B!
they will allow for characterization of beard taphonomy according to ontogeny (important
due to the unusual developmental patterns of beards). Dried beards were not placed in the
freezer, as this was deemed unnecessary.
Pigmented and non-pigmented scales were obtained from the flank of a large, male
Nile crocodile (Crocodylus niloticus). No single scale is a solid color, so the designation
refers to the predominant level of pigmentation across the scale. The skin sample was
provided by La Ferme Aux Crocodiles (Pierrelatte, France). Scales were stored in the freezer.
The mane of a domestic horse (Equus ferus) was obtained from the Equine Centre of
Langford Veterinary Services (Bristol, UK). This sample was also not placed in the freezer.
Decayed samples
Naturally occurring turkey feather bacteria were cultured. One liter of salt broth was
prepared in a sterile (autoclaved) flask based on previous feather decay studies (Williams et
al. 1990). The recipe is as follows: (in grams/liter) 0.5 NH4CL; 0.5 NaCl; 0.3 K2HPO4; 0.4
KH2PO4; 0.1 MgCl-6H2O; 0.1 yeast extract. Non-experimental samples of darkly pigmented
turkey belly feathers were placed in the broth. An automated shaker/incubator (~37.5°C, 200-
150 rpm) allowed the naturally occurring bacteria to multiply in the broth for 56 days. It was
noted that after a month of culturing the naturally occurring bacteria on the turkey belly
feathers, clears signs of decay were present. The broth became very murky. Murkiness was
apparent even after one day of culturing. Many small bits of feathers separated and floated in
the broth, indicating protein degradation. However, most of the feathers still retained much of
their original form.
After this bacterial broth was cultured, it was applied to various experimental feather
samples. Black, white, and iridescent feathers (Table 1) were placed in Pyrex jars with a
!
C!
plastic spacer on top to keep the feathers submerged. The clean salt broth was then poured
into the jars with a small amount of the cultured bacterial broth added to introduce the
microorganisms needed for rapid decay. During pouring, the cultured bacterial broth was
filtered through pantyhose to prevent the previous feathers from contaminating the
experimental samples. As a control, another jar contained four darkly pigmented turkey back
feathers in mineral water. The four jars were then placed into an incubator at ~37°C for 50
days (although a power outage caused temperatures to drop for several days during this time).
At the end of the decay process, decayed feathers were removed from the Pyrex jars with
tweezers or filtered out with pantyhose depending on their level of degradation and then
rinsed with ethanol on top of filter paper to clean them and prevent further decay.
Maturation
Some fresh and decayed samples were subjected to maturation through autoclaving
(Table 1). Samples were decontaminated by rinsing in acetone prior to loading into Au90 Pd10
capsules that were then sealed and placed into a water-pressurized autoclave. Capsules were
loaded with as many/as much of the samples while still allowing for the capsule to be cleanly
welded shut. Samples were cut into pieces in order to load into capsules. The weights of the
capsules were taken prior to and after autoclaving to determine if water infiltrated the capsule
due to a failed weld (see below).
Some fresh feathers (white and dark) were matured at 250°C/250 bars in order to
produce a ‘goo’ that was accidentally discovered during failed maturation attempts. During
these failed attempts, white feathers were first placed in an Ar gas autoclave at 250°C/250
bars for 24 hours, but the sample was lost as the resulting ‘goo’ flowed out of the tube. A
second attempt using white feathers matured at 200°C/250 bars for 24 hours in an Ar gas
!
D!
autoclave produced similar results and the sample was also lost. These failed attempts
prompted the use of a wet autoclave (and sealed capsules) at 250°C/250 bars for 24 hours to
produce the ‘goo’ that was ultimately analyzed here.
The rest of the samples matured in this study were heated to 100°C/250 bars to allow
for maturation while still maintaining enough morphological structure for microscopic
analysis. All runs lasted for 24 hours.
Structural analysis
Structure was analyzed using light microscopy and SEM on portions of the matured
and non-matured samples listed in Table 1.
Fresh samples underwent critical point drying prior to microscopy (decayed and
matured samples did not). Samples were left on an automated roller while washed in a series
of chemicals. Samples were fixed in 10% neutral buffer formalin for 24 hours then placed
through a series of 20-minute washes – two washes of distilled water followed by an ethanol
series of 30, 50, 70, and 90% ethanol concentration and, finally, two washes at 100%
concentration. Samples were then critically point dried using hexamethyldisilazane.
The decayed feathers underwent three different treatments in preparation for
microscopy. For each feather type, some of the sample either had 1) no treatment prior to
gold-coating, 2) critical point drying, or 3) 2% Triton X-100 detergent wash (30 minutes on
an automated roller) followed by critical point drying. Critical point drying alone was done in
the hopes of preserving the microbial community on the feathers while the Triton X-100
wash was intended to remove the microbial community to allow for clear study of the feather.
Samples were then affixed to SEM stubs using double-sided carbon tape. Some larger
samples had to be cut in order to fit well onto the stub (e.g., feathers) and others were cut in
!
E!
order to view them in cross section (e.g., feather rachises, turkey bristles, horse hair, and
scales) Prior to SEM, they were studied under a light microscope (Leica M205 C
stereomicroscope with a Leica DFC425 C camera).
SEM was conducted using a Hitachi S3500N variable-pressure scanning electron
microscope under vacuum (scanning electron mode) and samples were gold coated to reduce
charging. Silver paint was sometimes applied to the base of the sample to further reduce
charging.
The ‘goo’ samples were also observed in SEM by cutting open the capsule used
during autoclaving and allowing the ‘goo’ to extrude out onto the surface of the capsule. The
capsule was then affixed to SEM stubs using double-sided carbon tape. These samples were
not critically point dried, nor were they gold coated as the Au90 Pd10 capsule onto which the
‘goo’ extruded provided enough charge dissipation.
Water solubility test
The ‘goo’ produced at 250°C/250 bars underwent water solubility tests. The capsule
used for autoclaving was cut open and the half of the capsule not sent off for chemical
analysis was dropped into a glass vial to allow the ‘goo’ to extrude out and stick to the walls
of the vial. The original capsule was later removed for SEM (see above) and the vial was
filled with water. Photos of the ‘goo’ on the walls of the vial were taken with a light
microscope (Nikon Eclipse LV100D–U stereomicroscope) prior to, immediately after, and 1,
2, 5, 10, 15, and 20 minutes after the addition of water. The vials were then inspected ‘by
eye’ on a weekly basis to verify the initial observations.
Compression experiment
!
F!
Two small zebra finches (Taeniopygia guttata) were obtained from a local pet store
for compression experiments. Work with animals was approved by the UK Home Office and
all animals were euthanized according to Schedule One of the Animals (Scientific
Procedures) Act of 1986. The first bird was compressed fresh and the second was compressed
after a decay treatment. A subaqueous compression rig was designed (see below) consisting
of a container that can be placed into a bench press. The container was a stainless steel
cylinder with an 8 cm internal diameter. It sat on a specially designed base, made of recycled
plastic with grooves allowing water to escape from the bottom/sides during compression. To
load the container, sewing thread was passed into the container through two grooves on the
base. A few centimeters of sand/chalk/dirt mix was then added to the bottom.
For the first run, a finch was then passed under the thread, and the slack in the thread
was removed by pulling on the ends outside of the chamber in order to fasten the bird to the
sediment and prevent floating. The bottom was taped to seal the container, maintain tightness
on the thread, and prevent the water from flowing out. Water was then added and the bird
was buried in a chalk/dirt mix. A sieve was used to drop the calk/dirt mix into the water in
order to replicate low-energy, gradual burial. Fine grain sand was obtained from a local
garden shop. Chalk was purchased online. Dirt was obtained from a local park (with the
purpose of introducing bacteria for the next run where a decayed finch was compressed).
A thick plastic disk fit snuggly into the top of the cylinder of the container, allowing
for compression. A stainless steel spacing ring of the same diameter sat directly above, and in
contact with, the plastic disk and could be threaded onto the mainframe of the bench press,
adding stability. After the container was loaded, the tape was removed, and water was
allowed to flow out into a plastic bag. The container, still in the plastic bag, was then placed
into the bench press and the spacing ring was threaded on by spinning the container. The base
!
G!
of the bench press, on which the container sat, was raised using a manual crank. This
increased pressure on the sediment column and bird within the container as they are pressed
upwards. Pressure was monitored using a built-in pressure gauge. Pressure was increased to
approximately 25 tonnes and this pressure was applied for a minute. Water escaping from
pore spaces during compression flowed out of the specially designed base, while almost all
sediment was retained.
The container was taken out of the bench press and the sediment column was pushed
out of the cylinder. The sediment column was left in the fume hood for 6 days and then
placed into an oven for 24 hours at 60°C to dry the sediment. The column was then prepped
using a microblaster (without abrasive) and a dental pick. Photos were taken throughout the
prepping process with a Nikon D90 DSLR camera.
The second finch was crushed and prepared in the same manner as the first, with
several modifications. First, above the basal sand/chalk/dirt mix, chalk/dirt mix was added
before fastening the bird to the sediment so as to surround the bird in pure mud. Second, the
loaded container (with the water added, bird buried, and bottom taped) was partially sealed
with strips of parafilm at the top opening and placed into a plastic bag. It was left in an
incubator at ~37°C for 46 days (although a power outage caused temperatures to drop for
several days during this time). When it was taken out, it was noted that much of the sediment
had dried and that the head and part of the tail feathers had floated up to the top of the
column, so the next day chalk/dirt/sand mix was added to rebury the bird followed by pure
sand as a cap. The whole container was then submerged in water and the sediment was
probed in order to facilitate rehydration of the pore spaces. After a day of water submergence,
the bird was crushed. Third, the compression rig was slightly modified as the thread on the
bench press was partially stripped during the first run. The stainless steel ring was now
permanently attached to the thread that fits into the mainframe of the press and the plastic
!
H!
disk was replaced with a thinner plastic disk. Fourth, after compression, the sediment column
was left in a fume hood for three days before placing in the oven for drying.
Chemical analysis
Chemical analysis was done using Py/GC/MS. Thirteen samples were sent to
Newcastle University for decontamination (using methanol and dichloromethane) and
analysis. These were fresh and matured samples of FB, FW, ST, RT, and H as well as GD,
GW, and fresh BA (see Table 1 for abbreviations). The rest of the samples were also sent to
Newcastle University for Py/GC/MS but were decontaminated at the University of Bristol by
rinsing them in methanol and then dichloromethane for a several minutes. Data was analyzed
using HP Agilent ChemStation at Newcastle and Xcalibur at Bristol.
!
I!
SUPPORTING FIGURES AND TABLES
Dark and white feather types that were highly matured (250
°
C) into ‘goo’ for analysis.
!
J!
Sample (See Table 1
for abbreviations)
Number of
samples loaded
Did the capsule gain
weight after
autoclaving?
Was the capsule
crushed after
autoclaving?
CB
Small portion of
1 scale
No
Yes
CW
Small portion of
1 scale
No
Yes
BJ
Multiple bristles
with attached
epidermis
No
Yes
BA
Multiple cut
bristles with
attached
epidermis
No
No
FI
Tips of 2
feathers
No
Yes
DFI
Capsule half full
Yes
Yes
DFW
1 feather; Cut in
half
No
Yes
DFB
Capsule half full
No
Yes
H
Multiple cut
hairs
No
Yes
FB
22/3 feathers; Cut
No
No
ST
1 scale
No
Yes
RT
Multiple scales
with attached
epidermis
No
Yes
FW
2 feathers; Cut
No
Yes
GD
1 feather; Cut
No
Yes
GW
2 feathers; Cut
No
Yes
Autoclave capsule information.
!
K!
Compression rig. A–D, first version. E–F, second version. A, container, thick plastic disk,
and stainless steel spacing ring (from left to right). B, container placed in bench press (but not
with disk and ring threaded onto the frame). C, recycled plastic base with grooves. D,
container sitting on base. E, thin plastic disk and permanently attached stainless steel ring
threaded onto a subcomponent of mainframe. F, subcomponent with permanently attached
ring bolted to the bench press.
!
L!
Decayed feathers
Decayed white feathers. A–B, in Pyrex jars and salt/bacterial broth. C, after removing from
broth.
!
M!
Decayed black feathers. A–B, in Pyrex jars and salt/bacterial broth. C, after removing from
broth.
!
N!
Decayed iridescent feathers. A–B, in Pyrex jars and salt/bacterial broth. C, after removing
from broth.
!
O!
Decayed control feathers. A–B, in Pyrex jars and salt/bacterial broth. C, after removing
from broth.
!
P!
Structural features of decayed control feathers rinsed with ethanol after decay
treatment. A, under light microscopy, and B–F, SEM. B, plumulaceous barbs. C–E,
pennaceous barbs and barbules. F, cross section of rachis.
!
Q!
Water solubility test on ‘goo’
Light microscopy images of the ‘goo’ in vials during water solubility testing. A–C, ‘goo’
produced from dark feathers. D–F, ‘goo’ produced from white feathers. A, D, the vials prior
to the addition of water. B, E, the vials immediately after the addition of water. C, F, the vials
20 minutes after the addition of water.
!
R!
Total ion chromatograms
TIC of the ‘goo’. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
S!
TIC of iridescent feathers. Prominent mass spectrum base peaks (m/z) are labelled above
the chromatogram peaks.
!
T!
TIC of white feathers. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
U!
TIC of black feathers. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
V!
TIC of avian scales. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
W!
TIC of turkey beards. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
X!
TIC of crocodilian scales. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
!
Y!
TIC of mammalian hair. Prominent mass spectrum base peaks (m/z) are labelled above the
chromatogram peaks.
ResearchGate has not been able to resolve any citations for this publication.
Article
Full-text available
Exceptionally preserved organic remains are known throughout the vertebrate fossil record, and recently, evidence has emerged that such soft tissue might contain original components. We examined samples from eight Cretaceous dinosaur bones using nano-analytical techniques; the bones are not exceptionally preserved and show no external indication of soft tissue. In one sample, we observe structures consistent with endogenous collagen fibre remains displaying ~67 nm banding, indicating the possible preservation of the original quaternary structure. Using ToF-SIMS, we identify amino-acid fragments typical of collagen fibrils. Furthermore, we observe structures consistent with putative erythrocyte remains that exhibit mass spectra similar to emu whole blood. Using advanced material characterization approaches, we find that these putative biological structures can be well preserved over geological timescales, and their preservation is more common than previously thought. The preservation of protein over geological timescales offers the opportunity to investigate relationships, physiology and behaviour of long extinct animals.
Article
Full-text available
Many exceptionally preserved fossils have long been thought the product of preservation by bacterial autolithification, based largely upon the presence of, micron-sized, spherical or elongate bodies on their surface. This has recently been challenged by studies of similar fossils which cite morphological and geochemical evidence that these structures could be fossilized melanosomes, melanin-containing organelles. We geochemically analysed a tadpole from the Oligocene Enspel Formation, Germany, which displays such spherical bodies on its surface. Pyrolysis gas chromatography mass spectroscopy (Py-GCMS) and Fourier transform infrared spectrometry (FTIR) indicate that the organic remains of the tadpole are original and are not the result of external contamination, shown by the different chemical compositions of the fossil and its enclosing matrix. Py-GCMS also demonstrates the presence of bacterial and plant biomarkers in the matrix but not the tadpole, suggesting that the spherical bodies are unlikely to be bacterial, and also that such fossils do not develop their dark colour from incorporating plant material, as has been suggested. X-ray absorption spectroscopy (XAS) shows high levels of organically bound Zn(II) in the fossilized soft tissue, a metal known to chelate both eu- and pheomelanin. The zinc in the tadpole shows greater similarity to that bound in pheomelanized extant samples than to that in eumelanized ones. Though further geochemical analysis of both pure pheomelanin and bacterial samples is required to completely exclude a bacterial origin, these results are in line with a pheomelanic origin for the spherical bodies on the tadpole.
Article
Full-text available
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
Article
The variation and persistence of blood components, in particular red blood cells (RBCs), within bone tissue during the decomposition process, especially at the early stages and in different taphonomic conditions, has never been thoroughly investigated, regardless of the fact that knowing how blood survives or degrades within bone could be of help in solving many anthropological issues, such as trauma analysis and interpretation. This research investigated the influence of time and taphonomy on the persistence and detectability of blood components in parietal bone fragments (of different post mortem periods and taphonomic conditions) through histological (Hematoxilin and Eosin, HE) and immunohistochemical (Glycophorin A, GYPA) analyses. The immunohistochemical investigation for GYPA showed the presence of RBCs under the form of erythrocyte debris or residues otherwise morphologically unidentifiable using only HE staining. Hence, while well-defined RBCs can be observed only in the first week of decomposition, afterward these structures can be detectable with certainty only by immunohistochemical analysis, which reveals discrete quantities of RBC residues also in dry bone (post mortem interval, or PMI, of 15 years), but not in archaeological samples, in which the greater PMI and the different taphonomic conditions together could be the answer behind such difference. This study highlights the usefulness and potential of immunohistochemical detection of GYPA in RBC investigation and gives a realistic idea of the persistence and detectability of erythrocytes in different osteological taphonomic conditions, in contrast to results reported by some authors in literature. Another important result concerns the detection of RBC residues in dry bone, which opens the way to the possible use of RBCs in trauma interpretation. Am J Phys Anthropol, 2015. © 2015 Wiley Periodicals, Inc. © 2015 Wiley Periodicals, Inc.
Article
The integuments of extant vertebrates display a variety of epidermal appendages whose patterns, morphology and terminal differentiation (epidermal keratins) depend upon interactions between ectodermal (epidermis) and mesodermal (dermis) tissues. In reptiles and birds, appendage morphogenesis precedes terminal differentiation. Studies have demonstrated that appendage morphogenesis influences the expression of the appendage specific keratin genes. However, little is known about the nature of the structural genes expressed by the epidermal appendages of reptiles. How pattern formation and/or appendage morphogenesis influence terminal differentiation of reptilian appendages is not known. The epidermal appendages of reptiles and birds are characterized by the presence of both alpha (α) and beta (β) type keratin proteins. Studies have focused on the genes of avian β keratins because they are the major structural proteins of feathers. The occurrence of β keratin proteins in the scales and claws of both birds and reptiles and their immunological cross-reactivity suggest that the genes for reptilian β keratins may be homologous with those of birds. In bird appendages, the β keratins are the products of a large family of homologous genes. Specific members of this gene family are expressed during the development of each appendage. Recent sequence analyses of feather β keratins, from different orders of birds, demonstrate that there is more diversity at the DNA level than was implied by earlier protein sequencing studies. Immunological techniques show that the same antibodies that react with the epidermal β keratins of the chicken (Gallus domesticus) react with the epidermal β keratins of American alligators (Alligator mississippiensis). Furthermore, a peptide sequence (20 amino acids) from an alligator claw β keratin is similar to a highly conserved region of avian claw, scale, feather, and feather-like β keratins. These observations suggest that the β keratin genes of avian epidermal appendages have homologues in the American alligator. Understanding the origin and evolution of the β keratin gene families in reptiles and birds will undoubtedly add to our understanding of the evolution of skin appendages such as scales and feathers.
Article
Melanin, and other pigments have recently been shown to preserve over geologic time scales, and are found in several different organisms. This opens up the possibility of inferring colours and colour patterns ranging from invertebrates to feathered dinosaurs and mammals. An emerging discipline is palaeo colour: colour plays an important role in display and camouflage as well as in integumental strengthening and protection, which makes possible the hitherto difficult task of doing inferences about past ecologies, behaviours, and organismal appearance. Several studies and techniques have been presented in the last couple of years that have described ways to characterize pigment patterns. Here, I will review the available methods and the likely applications to understand past ecologies. A golden age of colourized dinosaurs and other animals is now dawning upon us, which may elucidate the nature of ancient predator prey interactions and display structures. Also watch the Video Abstract. © 2015 WILEY Periodicals, Inc.
Article
To provide baseline data for estimating the dietary amino acid requirements of molting birds, we measured the amino acid composition, nitrogen and sulfur content, and heat of combustion of the homogenized plumage of six White-crowned Sparrows (Zonotrichia leucophrys gambelii). On the average, the plumage contained 15.22% nitrogen, 3.14% sulfur, and 0.86% ash. Of the 17 amino acids measured, serine, proline, cystine/2, and glycine (in that order) were most abundant, comprising 46 mol% of the hydrolysate. Nonessential amino acids predominated (68 mol%). Cystine/2 contributed 11 mol%. Variation of amino acid composition among the plumages of the six birds was small (coefficient of variation = 2-7%). The heat of combustion of the homogenized plumages was 21.69 kJ/g of dry mass. Although exact quantitative comparisons are not possible, the amino acid composition of White-crowned Sparrow plumage in general parallels that of other species. Its cystine/2 and sulfur content, however, are noticeably greater than that reported for any other species thus far examined (primarily pale and domesticated).
Article
Fibre products, such as textiles and animal pelts, are often recovered in the corrosion crust of archaeological metal artefacts. Because clothed burials are an important resource for the study of past societies, accurate fibre identification is important. However, extreme mineralisation of animal fibres can render microscopic visualisation difficult for species identification. Peptide mass fingerprinting (PMF) has been successfully used to identify the species origin in both collagen and keratin-made archaeological artefacts. The approach requires little material but the state of degradation (protein hydrolysis) is a limiting factor as it might impact on the identification of key markers. In this study we analysed pelt and textile fragments found in association with copper-alloy objects with different degrees of mineralisation; samples were obtained from a Viking-Age (10th c.) grave in Britain and from a burial in Mongolia (3rd c. BC to 2nd c. AD). Species identification was possible in all but one sample, revealing PMF can be applied to corrosion products, thereby further expanding the value of these objects for textile research.