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Skin, scales, and cells in a Jurassic plesiosaur
Highlights
dThe first in-depth study of plesiosaur soft tissues is reported
dSome plesiosaurs had smooth skin on the body and small
scales on the flippers
dScales likely enhanced swimming and/or grip on the
substrate during feeding
Authors
Miguel Marx, Peter Sjo
¨vall,
Benjamin P. Kear, ..., Klaus Nilkens,
Michiel Op De Beeck, Johan Lindgren
Correspondence
miguel.marx@geol.lu.se
In brief
Marx et al. report the first detailed study
of fossilized plesiosaur soft tissues.
These show that plesiosaurs possessed
both smooth body skin and scaly flippers.
Marx et al., 2025, Current Biology 35, 1–8
March 10, 2025 ª2025 The Authors. Published by Elsevier Inc.
https://doi.org/10.1016/j.cub.2025.01.001 ll
Report
Skin, scales, and cells in a Jurassic plesiosaur
Miguel Marx,
1,7,8,
*Peter Sjo
¨vall,
2
Benjamin P. Kear,
3
Martin Jarenmark,
1
Mats E. Eriksson,
1
Sven Sachs,
4
Klaus Nilkens,
5
Michiel Op De Beeck,
6
and Johan Lindgren
1
1
Department of Geology, Lund University, So
¨lvegatan 12, 223 62 Lund, Sweden
2
RISE Research Institutes of Sweden, Materials and Production, P.O. Box 857, 501 15 Bora
˚s, Sweden
3
The Museum of Evolution, Uppsala University, Norbyv€
agen 16, 752 36 Uppsala, Sweden
4
Naturkunde-Museum Bielefeld, Abteilung Geowissenschaften, Adenauerplatz 2, 33602 Bielefeld, Germany
5
Urwelt-Museum Hauff, Aichelberger Straße 90, 73271 Holzmaden, Germany
6
Centre for Environmental and Climate Science, Lund University, So
¨lvegatan 37, 223 62 Lund, Sweden
7
X (formerly Twitter): @miguelpmarx
8
Lead contact
*Correspondence: miguel.marx@geol.lu.se
https://doi.org/10.1016/j.cub.2025.01.001
SUMMARY
Plesiosaurs are an iconic group of Mesozoic marine reptiles with an evolutionary history spanning over 140
million years (Ma).
1
Their skeletal remains have been discovered worldwide; however, accompanying fossil-
ized soft tissues are exceptionally rare.
2
Here, we report a virtually complete plesiosaur from the Lower
Jurassic (183 Ma)
3
Posidonia Shale of Germany that preserves skin traces from around the tail and front
flipper. The tail integument was apparently scale-less and retains identifiable melanosomes, keratinocytes
with cell nuclei, and the stratum corneum, stratum spinosum, and stratum basale of the epidermis. Molecular
analysis reveals aromatic and aliphatic hydrocarbons that likely denote degraded original organics. The
flipper integument otherwise integrates small, sub-triangular structures reminiscent of modern reptilian
scales. These may have influenced flipper hydrodynamics and/or provided traction on the substrate during
benthic feeding. Similar to other sea-going reptiles,
4–10
scalation covering at least part of the body therefore
probably augmented the paleoecology of plesiosaurs.
RESULTS AND DISCUSSION
The classic life reconstruction of plesiosaurs (Plesiosauria),
incorporating a long neck, compact body, and four propulsive
flippers, has not changed for nearly 200 years.
1,11
However,
the actual external appearance of these famous Mesozoic rep-
tiles is largely unknown. Recent comprehensive microscopic
and spectroscopic analyses of fossilized soft tissues have
shed light on the paleoecology and evolution of other extinct ma-
rine reptiles.
10,12–17
By contrast, with only eight plesiosaur soft
tissue specimens scientifically documented to date,
2,18–24
the
extreme rarity of such non-skeletal remains has hindered equiv-
alent studies.
Here, we report a well-preserved plesiosaur (Urwelt-Museum
Hauff, Holzmaden, Germany [MH] 7) from the world-renowned
Lower Jurassic (Lias εII
6C
, lower Toarcian
25,26
) Posidonia Shale
(Posidonienschiefer Formation) of southern Germany. MH 7
was excavated from a quarry near Holzmaden in 1940. Prepa-
ration of the skeleton in 2020 uncovered soft tissue traces
from around the tail and trailing edge of the right forelimb
(Figure 1A). We therefore applied a suite of techniques,
including transmitted light microscopy (TLM), scanning elec-
tron microscopy (SEM), energy-dispersive X-ray spectroscopy
(EDX), electron backscatter diffraction (EBSD), infrared (IR)
microspectroscopy, and time-of-flight secondary ion mass
spectrometry (ToF-SIMS) to examine MH 7 in unprecedented
detail.
Description and analysis
The fossilized soft tissues of MH 7 (Figures 1,2, and 3) are mainly
exposed along the dorsal and ventral sides of the caudal verte-
bral column (Figures 1B and 1C) and behind the bones of the
right forelimb. Small patches also cover the ends of some caudal
ribs, the caudal neural spines, and distal phalanges. We identify
these soft tissue remains as skin based on the presence of a
distinct internal layering that morphologically corresponds to
the stratum corneum, stratum spinosum, and stratum basale of
the integument in living amniotes.
27–29
The stratum spinosum of the tail skin in MH 7 is up to 215 mm
thick, whereas the stratum corneum is relatively thin (15–
25 mm). Fossil keratinocytes are observable within the stratum
spinosum. These are 20 mm in diameter and sub-circular in
outline, with dark centers that we interpret as remnant cell nuclei
(Figures 1D and 1E). The preserved keratinocytes become flat-
tened toward the external surface of the skin (facing into the
sediment matrix) and are thus consistent with corneocytes
(enucleated keratinocytes), which comprise the stratum cor-
neum in modern reptiles (Figure 1F). Clusters of dark-colored
melanosome microbodies derived from decayed melanophores
are also dispersed through the outer skin layers (Figures 1G and
1H). In addition, the stratum basale is demarcated by columnar
keratinocytes (Figures 1I and 1J).
Voids occur throughout the stratum spinosum where keratino-
cytes are not preserved. EDX of petrographic thin-sections de-
tected predominantly calcium and phosphorus in the stratum
Current Biology 35, 1–8, March 10, 2025 ª2025 The Authors. Published by Elsevier Inc. 1
This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).
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Please cite this article in press as: Marx et al., Skin, scales, and cells in a Jurassic plesiosaur, Current Biology (2025), https://doi.org/10.1016/
j.cub.2025.01.001
Figure 1. Plesiosaur specimen (MH 7) with comparisons
(A) MH 7 in ventral view, showing soft tissue sampling sites (arrows) on the dorsal (1) and ventral side (2) of the tail and trailing edge (3) of the right front flipper.
(B) Skin from the ventral side of the tail showing thick folds (white arrows) and an apparently torn surface (black arrow).
(C) Skin from the dorsal side of the tail (white arrows).
(D and E) (D) TLM image of skin from (B) with diagram (E), showing the differentiated stratum corneum (sc) and underlying stratum spinosum (ss), keratinocytes
with cell nuclei (circular structures with black dots), melanoso me aggregates (brown patches), voids (light gray patches), and indeterminate organics or minerals
(dark gray patches).
(F) Comparative thin-section through the eyelid epidermis of an extant Leatherback turtle, Dermochelys coriacea, indicating the similarly differentiated sc, pre-
corneous layer (pl), ss, and keratinocytes (ker).
(G) TLM image of a petrographic thin-section showing skin from the ventral side of the tail in MH 7. Inset: SEM image enlargement of ellipsoidal melanosome
microbodies (red box).
(H) TEM image of carapace skin from D. coriacea, showing differentiated sc and ss, including melanosomes (dark dots).
(I and J) (I) TLM image with diagram (J) showing a petrographic thin-section of skin from the dorsal side of the tail in MH 7 incorporating the sc, ss with a remnant
melanophore, and the stratum basale.
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j.cub.2025.01.001
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spinosum, whereas the stratum corneum is enriched with phos-
phorus but lacks calcium (Figure 2A). EBSD similarly found
calcite in the stratum spinosum but little or no calcite in the stra-
tum corneum (Figures 2B and 2C). This distribution implies pro-
gressive mineralization from the exterior to interior of the skin
and, presumably, involved calcium phosphate replication of
the stratum corneum versus partial replacement of the stratum
spinosum by calcium phosphate and, later, calcite that filled
voids left by more advanced decay.
Ellipsoidal melanosome microbodies are evident within the
tail skin of MH 7 after demineralization (Figures 2D–2G). The in-
ternal skin surface (facing away from the sediment matrix)
is covered by irregular pits and what appear to be thickened
folds (Figure 2H). Several overlying skin fragments were
Figure 2. Plesiosaur tail skin
(A) EDX elemental map of a petrographic thin-section through skin from the ventral side of the tail in MH 7, indicating the phosphorus (P)-rich sc and P/calcium
(Ca)-rich ss. Colors: cyan, silicon (Si); purple, P; orange, sulfur (S); blue, Ca; yellow, fluorine/aluminum (F, Al).
(B) EBSD map of (A). Color: blue, calcite (Cal).
(C) EBSD map of skin from the dorsal side of the tail, showing the ss permeated with Cal. Colors: green, apatite (Ap); purple, Cal; red, quartz (Qtz).
(D and E) (D) Light microscopy (LM) image of skin from the ventral side of the tail in MH 7 showing clusters of melanosome microbodies (dark dots) with TLM image
of the same skin after demineralization (E).
(F and G) (F) SEM image of demineralized skin from the ventral side of the tail in MH 7 with enlargement (G) of melanosome microbodies.
(H and I) (H) Skin from the ventral side of the tail showing pitted ss with apparent folding (white arrow) and tearing (black arrows). Smooth areas of the ss (red
arrow); (I) enlargement of possible torn skin from (H).
See also Figures S3 and S4.
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j.cub.2025.01.001
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seemingly torn to expose an interdigitating ‘‘zig-zagged’’
texture (Figure 2I).
By contrast, skin from the right forelimb incorporates small
and irregularly sub-triangular structures that we recognize as
scales (Figure 3A). These were coated in sulfur precipitates prior
to demineralization (Figure 3B) but retained structural integrity
after demineralization to reveal smooth surfaces that lack mela-
nosomes (Figures 3C and 3D). They clearly differ from the soft,
scale-less skin occurring around the tail of MH 7 and instead
compare closely with carapace scutes of fossil and living turtles
(Figures S1A and S1B), as well as mosasauroid marine lizard
scales (Figures S1C and S1D), which can similarly exhibit
Figure 3. Plesiosaur flipper scales
(A) Scales from the trailing edge of the right flipper in MH 7 (see Figure 1A), showing their irregularly sub-triangular shape and light-colored midline sediment infill.
(B) SEM image with inset EDX elemental map (orange dashed line: scale boundary). Colors: red, S; blue, P; yellow, Si.
(C) SEM image of demineralized scale fragment from MH 7 showing a smooth surface.
(D) Enlarged SEM image showing absence of melanosome microbodies on the scale surface of MH 7.
(E) SEM image of the scale margin from MH 7 showing internal layering (white arrows).
(F and G) (F) TLM image with diagram (G) of a petrographic thin-section through the scaly flipper skin from MH 7 showing ker (black arrows), cell nuclei (black
dots), and the outermost dense corneocyte layer (brown fill) covered by mineral deposits (white fill).
See also Figures S1 and S2.
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j.cub.2025.01.001
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sediment infilling along their midline, exposed by erosion of the
longitudinal keel
7
(contrary to smooth ichthyosaur skin; Figure
S1E). Diagnostically, the stratified edges of the scales in MH 7
(Figure 3E) have a thick (36 mm) outermost corneocyte layer
(Figures 3F and 3G) that is identical to the tough stratum cor-
neum characterizing the scaly skin of modern reptiles.
30–32
Our ToF-SIMS analyses (Figures S2 and S3) detected polycy-
clic aromatic hydrocarbons (PAHs) and aliphatic hydrocarbons
indicative of asphaltene-like molecular structures in the tail and
flipper skin of MH 7. Eumelanin was not identified, and may
have been diagenetically transformed into PAH-like compounds
during fossilization. IR spectra also produced peaks attributable
to hydrocarbons: 1,378 cm
1
(d
s
(CH
3
)) and 1,458 cm
1
(d
as
(CH
3
)d
s
(CH
2
)) (C–H distortion)
15
; 1,714 cm
1
(v(C=O))
33
; 2,852
and 2,923 cm
1
(C–H stretches).
15
Peaks in the 1,000 to
1,100 cm
1
region correspond to the phosphate (PO
4
3
)
group
15,34
(Figure S4).
Interpretations of plesiosaur skin
The body of MH 7 was seemingly covered in a mosaic of smooth
skin and (possibly keeled) scales like some turtles.
10
Indeed, the
epidermal microstructure of MH 7 corresponds well to that of
living amniotes, with an outermost stratum corneum comprising
compacted corneocytes and an underlying stratum spinosum
composed of younger, sub-spherical skin cells.
27–29
Frey et al.
2
suggested that the rarity of recovered plesiosaur skin
might be due to their epidermis being thin, as in extant snakes,
35
and thus susceptible to rapid breakdown after death. Conversely,
the combined depthof the stratum spinosum andstratum corneum
in MH 7 (which was at least 250 mm) implies a thick epidermis,
more compatible with that of living sea turtles (Figures S1Fand
S1G). The low preservation potential of plesiosaur skin in the Pos-
idonia Shale could, therefore, bea consequence of their infrequent
burial in dysoxic seafloor deposits conducive to soft tissue fossil-
ization
24,36
or because their skin remnants were unintentionally
removed during mechanical/chemical preparation.
Melanophore traces are situated close to the outer surface of
the tail skin in MH 7, which contrasts with many modern reptiles,
where pigment cells are typically concentrated in the deep
epidermis and superficial dermis.
37,38
However, epidermal
pigment cells can also be located just below (but not within)
the stratum corneum,
30
thereby potentially explaining the
pigment cell arrangement that we observe in MH 7.
The flipper skin of MH 7 is otherwise very similar to that of
modern scaly reptiles.
32,39
In particular, the outer corneocyte
layer is devoid of melanophore traces and substantially thicker
than that of the scale-less skin from around the tail. This differ-
ence is expected because the highly keratinized outermost
epidermis of reptilian scales usually comprises a deep layer of
compacted corneocytes
40
(Figure S1H). These cells are charac-
terized by corneous beta-proteins that make living reptile scales
hard and immobile.
41
Yet, our ToF-SIMS analyses failed to
detect any residual proteinaceous matter, probably because
the molecular content of MH 7 was completely transformed by
diagenetic processes during fossilization.
Functional roles of plesiosaur skin
In conjunction with MH 7, we examined other Posidonia Shale
plesiosaurs that preserve soft tissue remains. The most
informative were specimens of the microcleidids (Micro-
cleididae) Seeleyosaurus guilelmiimperatoris (Museum fu
¨r
Naturkunde Berlin, Germany [MB].R.1992) and Microcleidus
brachypterygius (Pal€
aontologische Sammlung, Universit€
at Tu
¨-
bingen, Germany [GPIT]-PV-30094). These complete skeletons
exhibit associated integument remnants (incorporating dark,
presumably melanic residues and embedded ‘‘fibrous struc-
tures’’
15,17,42,43
) behind the bones of the forelimbs (Figures
4A–4C). Such tissue traces derive from the flexible trailing
edge of the flippers that served to generate thrust as hydrofoils
in underwater flight.
44,45
Our novel detection of scales on the
flipper of MH 7 might, therefore, imply a hydrodynamic function
because these integumentary appendages may have stiffened
the trailing edge during swimming.
9,10
Not surprisingly, soft tis-
sue trailing edges have also been reported on the hindlimb hy-
drofoils
46
of Posidonia Shale microcleidids (Staatliches
Museum fu
¨r Naturkunde Stuttgart, Germany [SMNS] 51945)
24
and in the Late Cretaceous short-necked polycotylid (Polycoty-
lidae) Mauriciosaurus fernandezi, which purportedly possessed
scale rows along the body
2
(although these could be artifacts of
taphonomic deformation and/or cracking). Furthermore, MB.R.
1992 has a large, fleshy tail fin (Figures 4D and 4E) that was
either vertically
19,47
or horizontally
48
oriented and potentially
acted as a rudder for maneuverability
47
or to facilitate caudally
driven propulsion.
48
Another purpose of the flipper scales in MH 7 may have been
to provide a protective covering for traction on the seafloor dur-
ing benthic ‘‘grazing.’’ This is consistent with plesiosaur ‘‘bot-
tom-walking’’
49,50
and feeding traces,
51
as well as preserved
gastric contents,
24,52
which comprise coarse sediment masses
in SMNS 51945
24
and MB.R.1992—the latter individual had
also swallowed a mixture of small gastropods and cephalopods
(Figures 4F and 4G). Extant sea turtles
53
and dolphins
54
likewise
ingest large volumes of sand and mud when sifting through sea-
floor sediments for prey.
In summary, our comprehensive morphological, micro-
scopic, and spectroscopic investigation of the soft tissue res-
idues in MH 7 suggests that plesiosaurs (and more basal sau-
ropterygians
22
) retained reptilian scaly skin throughout their
land-to-sea transition and later specialization for life in the
open ocean. This contrasts with other Mesozoic marine rep-
tiles, including ichthyosaurs
15,55
and metriorhynchid crocody-
lomorphs,
56
which lost or reduced their scalation to reduce
drag. Although the scale-less tail skin of MH 7 hints at an anal-
ogous external body appearance, the presence of at least par-
tial squamation along the trailing edges of the flippers un-
doubtedly fulfilled some functional role and presumably
conferred a selective advantage for plesiosaurs during their
protracted evolution as one of the most successful pelagic
tetrapod clades.
RESOURCE AVAILABILITY
Lead contact
Further information and requests for resources and reagents should be
directed to and will be fulfilled by the lead contact, Miguel Marx (miguel.
marx@geol.lu.se).
Materials availability
This study did not generate new unique reagents.
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Data and code availability
MH 7 is accessioned and permanently housed in the publicly accessible
collection at MH. All experimental samples derived from MH 7 are stored at
the Department of Geology, Lund University (LU), Sweden. Any information
required to reanalyze the data reported in this paper is available from the
lead contact upon request.
ACKNOWLEDGMENTS
We thank R. Hauff, F. Hauff, B. Hauff, and A. Fichtner (MH); O. Hampe and S.
Thiel (MB); I. Werneburg and A. Krahl (GPIT); E.E. Maxwell (SMNS); R.M.
Carney (University of South Florida); D.K. Johansson (University of Copenha-
gen Zoological Museum); and D.A. Winkler and M.J. Polcyn (Southern Meth-
odist University) for access to specimens and information, and for their
hospitality during our research visits. F. Hauff contributed to the editing. C.
Sandt assisted with IR microspectroscopy. F. Borondics prepared samples
for SMIS analysis at SOLEIL. C. Alwmark, G. Zach
en, J. Martell, and A. Plan
contributed to SEM, EDX, and EBSD data processing. H.J. Go
¨tz (MB) took
UV photographs of MB.R.1992. O. Gustafsson and C. Rasmussen provided in-
formation on histological samples. Financial support included Swedish
Research Council grants to J.L. (2020-03542), B.P.K. (2020-03423), P.S.
(2019-03731), and M.E.E. (2019-03516). M.M. also acknowledges travel fund-
ing from the Royal Physiographic Society of Lund (42011).
AUTHOR CONTRIBUTIONS
Conceptualization, M.M., J.L., B.P.K., and S.S.; methodology, M.M., J.L., P.S.,
M.J.,and M.O.D.B.;investigation, M.M.,J.L., P.S.,and M.O.D.B.;writing– original
Figure 4. Plesiosaur soft tissues and gut contents
(A) UV image (cutoff at 365 nm) of soft tissues and glued edges (yellow arrows) from the trailing edge of the right flipper in Seeleyosaurus guilelmiimperatoris
(MB.R.1992). White box indicates enlargement (B).
(B) Enlargement from (A) showing skin traces with embedded 3D ‘‘fibrous structures’’
15,17,42,43
(white arrows) and surrounding glue (red arrows).
(C) Left forelimb trailing edge (white arrows) of Microcleidus brachypterygius (GPIT-PV-30094).
(D) UV image (cutoff at 365 nm) of the incomplete and partially restored
19,47,48
(yellow arrows) tail fin from S. guilelmiimperatoris (MB.R.1992).
(E) Enlargement from (D) showing fibrous structures.
15,17,42,43
(F) Preserved gut contents from S. guilelmiimperatoris (MB.R.1992), comprising a coarse sediment mass with a gastropod shell (white arrow).
(G) Possible belemnite phragmocone/guard (white arrow) and onychite (arm hook: cyan arrow) within the preserved gut contents from S. guilelmiimperatoris
(MB.R.1992).
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draft, M.M.; writing – review and editing, B.P.K., M.M., J.L., M.E.E., M.J., S.S.,
K.N., and M.O.D.B.; visualization, M.M., P.S., and K.N.; funding acquisition,
J.L., B.P.K., M.E.E., P.S., and M.M.
DECLARATION OF INTERESTS
The authors declare no competing interests.
STAR+METHODS
Detailed methods are provided in the online version of this paper and include
the following:
dKEY RESOURCES TABLE
dMETHOD DETAILS
BSample preparation
BTLM and thin-sectioning
BSEM, EDX, and EBSD
BIR microspectroscopy and ToF-SIMS
BComparative samples
SUPPLEMENTAL INFORMATION
Supplemental information can be found online at https://doi.org/10.1016/j.
cub.2025.01.001.
Received: March 15, 2024
Revised: October 28, 2024
Accepted: January 3, 2025
Published: February 6, 2025
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STAR+METHODS
KEY RESOURCES TABLE
METHOD DETAILS
Sample preparation
Samples of fossilized soft tissues from MH 7 were removed using hand tools and loosely wrapped in aluminum foil before being
placed inside individual plastic containers that were rinsed with ultrapure water (Milli-Q) and sterilized with 96% ethanol (Avantor
Inc., Radnor, USA). Following transport to LU, the samples were rinsed in Milli-Q water to remove adhering debris; they were then
sterilized again in 96% ethanol. All sample handling was done with forceps rinsed with Milli-Q water and sterilized using either
70% or 96% ethanol.
Prior to destructive processing, all samples were photographed using an Olympus SC30 digital camera mounted on an Olympus
SZX16 stereomicroscope, as well as an Olympus SZX10 stereomicroscope equipped with an Olympus SC50 digital camera. Sam-
ples selected for demineralization were then cut using a sterilized Dremelsaw and placed into sterile 5 mL jars before being rinsed
with 96% ethanol and set to air dry on aluminum foil. The dried samples were then stored at 4C in separate sterile 5 mL jars capped
with aluminum foil.
TLM and thin-sectioning
For TLM, demineralized tail and forelimb skin samples were mounted onto Menzel SuperfrostPlus (Thermo Scientific Inc., Wal-
tham, USA) glass slides with Menzelmicroscope cover slips (Thermo Scientific Inc., Waltham, USA) for observation using an
Olympus BX51 fluorescence microscope with an Olympus DP74 digital camera. For petrographic thin-sectioning, non-demineral-
ized/untreated tail and forelimb skin samples were embedded in a 5:1 mixture of Araldite DBF (ABIC Kemi AB, Norrko
¨ping, Sweden)
epoxy casting resin and RenHY 956 hardener (Huntsman Corporation, The Woodlands, USA), which was set to cure in an oven at
45–50C for 24 hrs. The samples were then ground until transparent using 600 and 1200 grit resin-bonded diamond discs (Struers
Inc., Ballerup, Denmark), and polished using polishing pads and diamond paste. Cyanoacrylate glue prevented breakage during
grinding. Imaging of petrographic thin-sections was undertaken on an Olympus BX50 polarized light microscope equipped with
an Olympus SC50 digital camera.
SEM, EDX, and EBSD
Prior to demineralization, forelimb skin samples were imaged at LU on a TESCAN Mira3 High Resolution Schottky Field Emission Gun
Scanning Electron Microscope (FEG-SEM) under low vacuum and without coating. Working distance was 6–9 mm with an electron
energy of 5 keV. EDX and EBSD of thin-sectioned skin samples was also carried out at LU with an X-MaxN 80 detector (124 eV,
80 mm
2
) and a NordlysNano detector (Oxford Instruments, Abingdon, UK) attached to the FEG-SEM. EDX and EBSD used working
distances of 16/19 mm and an electron energy of 20 keV for skin samples from dorsal/ventral sides of the tail, respectively. EDX and
EBSD data was processed using AZtecLive 6.1 (Oxford Instruments, Abingdon, UK), with post-processing for removal of wild spikes
using AZtecCrystal (Oxford Instruments, Abingdon, UK).
REAGENT or RESOURCE SOURCE IDENTIFIER
Software and algorithms
AZtecLive 6.1 Oxford Instruments https://nano.oxinst.com/products/azteclive
AZtecCrystal Oxford Instruments https://nano.oxinst.com/azteccrystal
OCTAVVS 0.1.22 Troein et al.
57
https://pypi.org/project/octavvs/
OMNICSpectra Software Thermo Fischer Scientific https://www.thermofisher.com/order/catalog/product/833-036200
Quasar Toplak et al.
58,59
https://quasar.codes/
SurfaceLab, version 7.1 IONTOF GmbH https://www.iontof.com/
Biological samples
MH 7 fossilized tissue This paper MH 7
Dermochelys coriacea This paper ZMUC-R2106; LO unnumbered
Ctenochelys acris This paper SMU 76353
Trachemys scripta elegans This paper SMU R133
Caretta caretta This paper ZMUC-KPC16030906
Varanus exanthematicus This paper LO 10298
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j.cub.2025.01.001
Report
FEG-SEM imaging was also undertaken at RISE Research Institutes of Sweden in Bora
˚s, Sweden. Tail and forelimb skin samples
were imaged both before and after demineralization, and under high vacuum without coating using a Zeiss Supra 40VP FEG-SEM
equipped with an Everhardt-Thornley type electron detector (SE2). The working distance was 3–5 mm, and the electron energy
was 1–2 keV. EDX used X-Max 50 mm
2
detector (Oxford Instruments, Abingdon, UK) set at a working distance of 8.5 mm and an
electron energy of 15 keV.
IR microspectroscopy and ToF-SIMS
Our skin samples were separated into two subsets for demineralization: (1) those taken from the ventral and dorsal sides of the tail
(N=2); (2) those taken from the ventral side of the tail and flipper (N=3). Demineralization of the subset (1) samples from the ventral side
of the tail proceeded in a molecular biology grade ethylenediaminetetraacetic acid (EDTA) solution of 0.5 M and pH 8 (AppliChem,
Darmstadt, Germany) for one week, during which the supernatant was replaced with fresh EDTA every other day. The samples were
then washed by exchanging the supernatant with a Bioultra grade solution of 0.25 M ammonium formate (Sigma-Aldrich, St. Louis,
USA) 14 times, and then rinsed six times with a solution of 0.15 M ammonium formate. After rinsing, pieces of the demineralized skin
were pipetted onto two silicon wafers (Sci-Mat Silicon Materials, Kaufering, Germany) for ToF-SIMS analysis, and onto two CaF
2
win-
dows (Crystran Ltd, Dorset, UK) for IR microspectroscopy; these were left to air dry. The CaF
2
window samples were placed in a
vacuum chamber (10
-7
mbar) for 1.5 hrs to evaporate the ammonium formate salt (NH
4
HCO
2
). Additional rinsing in 15% aqueous
sodium chloride (Sigma-Aldrich, BioXtra grade) was also performed to remove residual NH
4
HCO
2
prior to IR microspectroscopy.
Demineralization of the subset (1) samples from the dorsal side of the tail took place in EDTA for 20 days with the supernatant being
replaced every other day. This was followed by 20 rinses in Milli-Q water before drying on a CaF
2
window and sterile silicon wafer.
The subset (2) samples from the ventral side of the tail and the flipper were first inspected under a stereomicroscope to detect any
residual glue. They were then placed in an EDTA solution (refreshed three times for the subset [2] flipper tissue samples) for 22 and
26 days, respectively. One of the subset (2) tail samples was then rinsed with 0.25 M NaCl and 0.15 M NaCl sequentially 10 times for
each before being examined using IR microspectroscopy. The other subset (2) tail sample was rinsed in 0.25 M ammonium formate
and 0.15 M ammonium formate 10 times for each prior to ToF-SIMS analysis. The subset (2) flipper samples were rinsed 10 times with
0.25 M ammonium formate, and six times in 0.15 M ammonium formate before ToF-SIMS examination.
IR microspectroscopic measurements were undertaken at the SOLEIL synchrotron facility in France, and independently at LU.
SOLEIL used a Nicolet Continuum FT-IR microscope (Nicolet CZ, Prague, Czech Republic) with a synchrotron light source in trans-
mission mode. IR spectra were visualized and studied using both OMNICSpectra Software and Quasar.
58,59
LU used a Hyperion
3000 IR microscope coupled to a Tensor 27 spectrometer (Bruker Corp., Billerca, USA). IR spectra were recorded in transmission
mode using a Focal Plane Array detector with 64 364 elements. They were constructed from 1024 averaged scans and corrected
for atmospheric contributions from CO
2
and H
2
O using rubber band baseline correction in OCTAVVS v.0.1.22.
57
ToF-SIMS was carried out at RISE on a TOFSIMSIV instrument (IONTOF GmbH, Mu
¨nster, Germany) using 25 keV Bi
3+
primary ions
and low-energy electron flooding for charge compensation. Positive and negative ion data were acquired in the static SIMS regime
(accumulated primary ion dose density kept below 3 310
12
cm
-2
), with optimization for either high mass resolution (bunched mode,
m/Dmz5,000, lateral resolution 3–5 mm) or high image resolution (m/ Dmz300, lateral resolution 0.5–1 mm). Spectra and images
were generated using SurfaceLab v.7.1 (IONTOF GmbH, Mu
¨nster, Germany). Mass spectra of the glue and surrounding rock matrix
were measured as controls to compare with spectra from the fossilized soft tissues.
Comparative samples
Our comparative samples included carapace scutes from the fossil sea turtle, Ctenochelys acris (Southern Methodist University, USA
[SMU] 76353), and the extant Red-eared slider turtle, Trachemys scripta elegans (SMU R133), flipper skin from a juvenile Loggerhead
turtle, Caretta caretta (University of Copenhagen Zoological Museum, Denmark [ZMUC]-KPC16030906), dorsal body skin from an
adult Savannah monitor, Varanus exanthematicus (Department of Geology, Lund University, Sweden [LO] 10298), flipper skin
from a juvenile Leatherback turtle, Dermochelys coriacea (ZMUC-R2106), and carapace and eyelid skin from an adult D. coriacea
(LO unnumbered sample provided by Marine Turtle Permit 073 from the US Fish and Wildlife Service and Florida Fish and Wildlife
Conservation Commission).
Scute tissue was removed from SMU 76353 using hand tools and demineralized before being stored in EDTA for an extended
period of time. The samples were then mounted on SEM stubs for imaging at RISE using a Zeiss Supra 40VP FEG-SEM equipped
with an Everhardt-Thornley type electron detector (SE2).
Untreated scute tissue from SMU R133 was coated with a 15 nm thick AuPd film prior to imaging at RISE on a Zeiss Supra 40VP
FEG-SEM equipped with an Everhardt-Thornley type electron detector (SE2).
Flipper skin from ZMUC-KPC16030906 and ZMUC-R2106 were fixed in 2.5% glutaraldehyde and 70% ethanol, respectively. They
were then sectioned using a sterile scalpel and imaged using an Olympus SZX10 stereomicroscope equipped with an Olympus SC50
digital camera.
Dorsal body skin from LO 10298 was fixed in 4% So
¨rensen phosphate buffered formalin (pH 7.2). The tissue was then embedded in
EPON (Agar Scientific, Stansted, UK) and stained with Richardson’s solution (methylene blue and Azure II)
60
before being cut into
3mm sections with a Leica UC7 ultramicrotome. Imaging used an Olympus BX35 polarized light microscope equipped with an
Olympus UC30 digital camera.
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The LO unnumbered D. coriacea carapace and eyelid skin were fixed overnight using 2.5% glutaraldehyde and 2% paraformal-
dehyde in a 0.1 M sodium cacodylic buffer solution. They were then post-fixed using 2% osmium tetroxide in distilled water at 7
C for 1 hr. Dehydration used a graded ethanol series of 70% for 2 310 min, 96% for 2 310 min, and 100% for 2 315 min before
being embedded in Agar 100 resin (Agar Scientific, Stansted, UK) with acetone. Ultrathin sections were cut using a Leica UC7 ultra-
microtome to 50 nm for the carapace skin, and 1 mm for the eyelid skin which was stained with Richardson’s solution
60
and imaged
using an Olympus BX35 polarized light microscope equipped with an Olympus UC30 digital camera. The carapace skin sections were
mounted on copper grids and stained with 2% uranyl acetate for 30 min, and Reynolds lead citrate for 3 min before imaging using a
JEOL 1400 Plus Transmission Electron Microscope at 100 kV equipped with a CMOS camera.
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