
==== Front
Sci Rep
Sci Rep
Scientific Reports
2045-2322
Nature Publishing Group UK London

39256439
71313
10.1038/s41598-024-71313-6
Article
Biomolecules in Pleistocene fossils from tropical cave indicate fossil biofilm
de Sousa Daniel Vieira daniel.vsousa@univasf.edu.br

1
Maia Paulo Victor Sciammarella 2
Eltink Estevan 3
de Moura Guimarães Luciano 4
1 https://ror.org/00devjr72 grid.412386.a 0000 0004 0643 9364 Colegiado de Geografia, Universidade Federal do Vale do São Francisco, Senhor do Bonfim, Petrolina, 48970-000 Brazil
2 Instituto Federal de Educação de Minas Gerais, Arcos, Minas Gerais Brazil
3 https://ror.org/00devjr72 grid.412386.a 0000 0004 0643 9364 Colegiado de Ecologia, Universidade Federal do Vale do São Francisco, Senhor do Bonfim, Petrolina, 48970-000 Brazil
4 https://ror.org/0409dgb37 grid.12799.34 0000 0000 8338 6359 Departamento de Física, Universidade Federal de Viçosa, Viçosa, 36570-000 Brazil
9 9 2024
9 9 2024
2024
14 210719 2 2024
27 8 2024
© The Author(s) 2024
2024
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Finding biomolecules in fossils is a challenging task due to their degradation over time from physical, chemical, and biological factors. The primary hypothesis for explaining the presence of biomolecules in fossilized bones tissues suggests their survival in the fossilization process. In contrast, some of these biomolecules could either derive from bacteria biofilm, thus without a direct relationship with the fossil record or could be an artifact from measurement procedures. Raman spectroscopy studies across various fossil ages and environments have detected multiple bands ranging from 1200 to 1800 cm−1 associative of organic compounds. However, the significance of these bands remains elusive. Our research aims to address this issue through a deep Raman spectroscopy investigation on Pleistocene teeth from Tayassu and Smilodon populator. These fossils were obtained from a well-preserved stratigraphic succession in Toca de Cima do Pilão cave, near the National Park of Serra da Capivara in semiarid Brazil. We propose two hypotheses to explain the presence of organic compounds related to 1200 to 1800 cm−1 Raman spectral range in fossil tissues: (i) these bands are biological signatures of preserved fossil biomolecules, or (ii) they are exogenous biological signatures associated with the bacterial biofilm formation during post-depositional processes. Our results align with the latter hypothesis, followed by biofilm degradation. However, the specific mechanisms involved in the natural biofilm degradation in fossil records remain unexplored in this study. In our case, the formation of biofilm on fossil bones is attributed to the oligotrophic conditions of the cave sediment matrix. We present a comprehensive model to elucidate the existence of biofilm on fossilized tissues, emphasizing the pivotal role of post-depositional processes, especially water action, in the cave environment. As the fossils were discovered in a cave setting, post-depositional processes significantly contribute to the formation of the biofilm matrix. Although our study provides insights into biofilm formation, further research is needed to delve into the specific mechanisms driving natural biofilm degradation in fossils.

Subject terms

Palaeontology
Mineralogy
Chemical biology
http://dx.doi.org/10.13039/501100019831 Conselho Nacional das Fundações Estaduais de Amparo à Pesquisa 432075/2018-6 de Sousa Daniel Vieira http://dx.doi.org/10.13039/501100006162 Fundação de Amparo à Ciência e Tecnologia do Estado de Pernambuco 0936-21-52799 de Sousa Daniel Vieira http://dx.doi.org/10.13039/501100003593 Conselho Nacional de Desenvolvimento Científico e Tecnológico 102032/2024-6 de Sousa Daniel Vieira issue-copyright-statement© Springer Nature Limited 2024
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pmcIntroduction

Fossil biomolecules are organic compounds, mostly composed of ancient DNA, proteins, amino acids, lipids, and others, which are the remains of ancient organisms1. Although finding biomolecules in fossil records is not an easy task, as they tend to degrade over time due to physical, chemical, and biological factors, knowledge of biomolecules in fossils is a valuable source that can provide useful insight concerning the evolution of life on Earth2. Additionally, fossil biomolecules knowledge could be a key to understanding the preservation of ancient life in astrobiology studies3.

The first hypothesis to explain the presence of biomolecules in fossil bone tissues deals with them being preserved during the fossilization process. Somehow the fossil diagenesis occurs in a slow way, most likely with poor mineral matrix dissolution and replacement, which may be linked to different diagenetic routes4. The diagenetic process in fossil records is site-specific5–7 and even varies in distinct locations in the same cave4. For example, an environment rich in carbonate water can help preserve collagen while a location without carbonate-rich water helps decompose the fossil organic fraction4. Additionally, some minerals, like calcite, can form the so-called mineral-binding proteins, which help preserve ancient biomolecules in the fossil record2, thereby preserving fossil organic matrix7,8. On the other hand, some of these biomolecules could derive from bacteria biofilm without a direct relationship with the fossil records9,10. Additionally, they could stem from laboratory contaminants, or result from instrumental artifacts as well as data processing11,12.

Biofilms can be found in many environments on Earth, including fossils, representing a communal way of living for microorganisms that are held together by extracellular polymeric substances (EPS)13. These EPS matrices can be formed from polysaccharides, lipids, proteins, genetic materials, and humic-like substances13,14; however, the EPS composition also changes depending on the nature of the microorganism, in addition to depending on environmental conditions such as pH, temperature, oxygen, nitrogen levels, and the surface properties13,15. Biofilms can protect microorganisms against extreme conditions such as ultraviolet (UV) radiation, extreme temperature, extreme pH, high salinity, high pressure, poor nutrients, antibiotics, among others13.

Fossilized bacteria biofilms have been reported in various places on Earth for example, from banded iron-formations16,17 to the stromatolite rocks18. Recent studies demonstrated that the biofilms have an important role in fossil preservation. They can mediate preservation of leaf 19, influence soft tissue preservation19–23, also mediate embryo preservation in Neoproterozoic and Cambrian fossils24. The mediation of the preservation of soft tissues occurs due to the biofilm can cover the soft tissues with fine detail24,25, encapsulating it and protecting it from diagenesis.

Raman Spectroscopy is a powerful tool8,23,26–28 when studying biofilms and soft tissue preservation on fossil records. Raman spectroscopy is a non-destructive technique that relies on the inelastic scattering of light. This process involves a small amount of monochromatic light changing color when scattered by a material. The resulting color change can be quantified and correlated with the vibrational modes of molecules or atoms in the sample, effectively serving as a fingerprint. Studies on fossilization processes covering different fossil ages (from Quaternary to Cretaceus) and environmental conditions using Raman spectroscopy report the presence of several bands between 1200 to 1800 cm−1 spectral range5,14,27,29. Piga et al.27 were the first to suggest their organic origin. Barros et al.29 try to link some of these bands to D and G bands characteristic to graphitic related material, whereas Jurašeková et al.14 related these bands to proteins and other organic molecules. However, such a pattern in some fossilized bone tissues is not completely understood14.

This study aims to contribute to this debate by deciphering this Raman signature through a deep Raman spectroscopy investigation on Pleistocene tooth from Tayassu and Smilodon populator collected from a well-preserved stratigraphic sequence in Toca de Cima do Pilão cave, near National Park of Serra da Capivara, in semiarid Brazil (Fig. 1). We have two hypotheses that can explain the existence of these organic signatures on fossil bones: i) these biological signatures are related to preserved fossil biomolecules, or ii) this organic signature is related to bacteria biofilm formation during post-depositional processes.Fig. 1 Location of the study area. (A,B) Global and Satellite image of the study area from Google Earth Pro Software (version 7.3.6.9796); The Yellow line represents the marble massif while the red line represents the cuestas landform.; (C) Panoramic view along of the marble massif were Toca de Cima dos Pilão is located. The white doted box represents the Fig. 1D; (D) Detail of Marble massif; (E) Cave entrance; (F) Cave map with the sampling location shown by the red rectangle. The cave map was courtesy by the Fundação Museu do Homem Americano. These figures are created using Google Earth Pro (https://www.google.com/earth/about/versions/#download-pro) and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/). Figure Created by Daniel V. de Sousa.

Methods

We analyzed fossils teeth samples from Tayassu (13 samples) and Smilodon populator (6 samples) collected across the trench profile of the Toca de Cima do Pilão cave (Fig. 2) (UTM coordinates: 23 L Zone, 768698.59 m E, 9019327.47 m S), near Parque Nacional Serra da Capivara, Piauí state, Brazil. The fossiliferous material was deposited in the scientific collection of the Paleontology Laboratory of Fundação Museu do Homem Americano (FUNDHAM), São Raimundo Nonato, Piauí state, Brazil. Some fossil samples exhibit fresh fractures that occurred during sample collection, revealing the inside of the teeth. Additionally, two samples were sectioned to access the innermost part of the teeth (Smilodon samples 4 and 6). These fractures are identifiable by their stark white hue and lack of siliciclastic sediment (refer to Supplementary Figure S1). For this study, these features are classified as the internal surfaces of the fossils.Fig. 2 Trench profile and fossil sampling. The illustrations of a pig and a paw denote the depths at which the Tayassu and Smilodon samples were collected. The geochronological data were taken from de Sousa et al.4. Figure was created using Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

The teeth samples were chosen for carbon isotopic analysis for paleoecology and fossil diagenesis study. However, during the Raman spectroscopy measurements, we found uncommon peaks and significant fluorescence which almost obliterated the hydroxyapatite and B-type carbonate peaks, which would preclude a fossil diagenetic study using Raman spectroscopy4. These uncommon peaks most likely point to biomolecules; therefore, we decided to investigate their origin more deeply which resulted in this study. We analyzed thirteen fossil samples from Tayassu and six fossil samples from Smilodon, within these samples just the first sample of Tayassu is a metapodial bone fragment, all the others are teeth samples.

Analytical techniques

Raman spectroscopy measurements were conducted in different regions of the samples (Figs. 3, 4). All raw data can be accessed in the Mendeley data repository (https://data.mendeley.com/datasets/nscghhnt63/2). These analyses were performed using a Renishaw micro-Raman inVia Spectrometer equipped with a diode laser at 785 nm. This specific choice of laser line is aimed at minor fluorescence effects. Fluorescence is a phenomenon typically present in natural materials, which hinders achieving the Raman spectrum. Higher wavelengths such as 785 or 1024 nm are used to reduce the fluorescence effect. We used a laser at 785 nm, and to ensuring that the laser intensity did not induce photodegradation of the sample, the laser power used is about 0.20 mW and the exposure time was 10 s with 1 accumulation. We obtained the spectra using a 50 × objective and NA = 0.75; in some cases, we used 50 × long distance objective. The spectra‘s baseline was extracted using a computational cubic mathematical model for all samples. After removing of the baseline, the Lorentzian curve fitting was used for adjusting of all Raman bands. Table1 shows the frequency of bands used in the deconvolution process to identify bone materials and biomolecules. All spectra were analyzed in MagicPlot 2.5.1 software.Fig. 3 Tayassu samples used in this study. The numbers in the pictures A to Ga represent the site of Raman measurements. The white bar in the bottom of the images is a scale of 1 cm. (A–C) sample 1, small bone fragment. (D,E) sample 2; (F–H) sample 3 cracked during sampling collection; (I,J) sample 4; (L) sample 5, the red box in the picture is the image M and N; (M) sample 5 were we note small patches of red sediments attached on the teeth (1a, 1b and 1c point analysis); (N) sample 5 red and black spots attached on the teeth (3d and 3b point analysis respectively); (O,Q): sample 6, the red box is represented in the picture P; (P) sample 6, a cracked filling by red sediment (2a, 2b 2c and 2d points); (Q) sample 6 the red box is the image R; (R) sample 6, the Raman measurements are taken at the edge of sample showing black and red pigments; (S) sample 7; (T) sample 8, the measurements was take on dentin, the red box is seen in image U; (U) sample 8, transition of dentin (4 and 5 points) and enamel (points 6, 7, 8 and 9), showing red sediments filling the cracks; (V) sample 9, the red box represents the site of point analysis found in detail in image X; (X) The edge of enamel (2a, 2b, 3a and 3b points) and dentin (1a, 1b and 1c) showing red sediments filling the cracks); (Z) sample 10 the red box represents the image Aa. Aa – sample 10, the measurements were taken on the enamel (6, 7, and 8 points) and dentin (1, 2, 3, 4, and 5 points), s showing the red sediments filling the cracks); Ba: sample 11 the red box represents the image Ca; Ca: sample 11, the point analysis was taken on the dentin; Da: sample 12 the red box represent the image Ea; Ea: sample 12, the Raman analysis was taken from dentin (point 6) and enamel (1 to 4 points), showing red sediments; Fa: sample 13 red box represents the image Ga; Ga: sample 13 the measurements were taken from the enamel, with the enamel fully coating by a black and red thin film. The sample in figure G shows a recently cracked face. The figure was created using microsoft power point (https://www.microsoft.com/pt-br/microsoft-365/).

Fig. 4 Smilodon populator samples used in this study. (A) sample 1, the red and black boxes point to the different analyzed regions. The detail of the red box is shown in Fig. 4B. In this sample, the interior faces were exposed during sample preparation; (B) sample 1 –analysis of the dentin and enamel. (C) Sample 1 – the red box is shown in detail in image D; (D) Dentin point analysis, showing a cracked filling by red sediments. (E–I) sample 2; (F) represent the red box seen in image E, showing black lines; (G) sample 2 – the measurements were taken in the yellow box of the image E, showing a few black features; (H) sample 2 – showing a region almost free of red and black features; the red box can be seen in detail in image I. In this sample, the interior faces were exposed during sample preparation; (I) sample 2 – the measurements were taken in the region fully cover by red and black features; (J) sample 3 – the site analysis can be seen in detail in image K; (K) sample 3 – a variety of the color features on the teeth, represented by white, gray, and black; (L–N) sample 4 – the red box in image L appears in image M; (M) sample 4 – dentin with black features; (N) sample 4 – the point analysis was taken in a recent cracked, completely free of red and black features. This sample was cutting for Raman measurements; (O–R) sample 5 O –the site of measurements on the enamel, and P analysis on dentin; the red box in image O appears in image Q, and the red box in image P appears in image R; (S–X) sample 6 – the red box shows the local analysis on the enamel, and appears in detail in image T; (T) detail of the red box in image S, showing a yellow sediment filling a crack; U sample 6 – showing a red and black boxes; the black box can be seen in detail in image V, and the red box appears in image X. The samples in figures U This sample was cutting for Raman measurements. The figure was created using Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

Table 1 Band assignment of bone and biomolecules materials used in this study.

Material type	Band assignment	Initial band position (cm−1)	Source	
Bone material	Hydroxyapatite (v1)	960	Thomas et al30; Piga et al27; de Sousa et al4	
Hydroxyapatite (v2)	432	
Hydroxyapatite (v3)	1049	
Hydroxyapatite (v4)	590	
Biomolecules	C = O strech, C–C	1721	Chao & Zhang31; Wiemann et al.26	
Pepitide, C = O strech	1682	
amide I, Tysosine, phenylalanine; polycyclic ring system	1614	Jenkins et al.32; Chao & Zhang31; Wiemann et al.26; Ivleva et al.33; Puppels et al.34; Jehlička & Edwards35	
N-H2 C-H band, C = C strech	1538	Chao & Zhang31	
C-H2 deformation associated with proteins, polysaccharides, and lipid	1445	Ivleva et al.36; Chao & Zhang31; Schuster et al.37; Gieroba et al.38	
C–H bend protein	1320	Chao & Zhang31; Löchte39; Schrader40	
Protein, Amide III	1287	Gieroba et al.38	
 = C δ(CH), lipids, proteins	1262	Gieroba et al.38	
Amide III	1233	Ivleva et al.33; Gieroba et al.38	
Carbohydrates, mainly C–C, C-O	1190	Schrader40	
 = C–C = unsaturated FA; (COC), glycosidic bonds	1117	Jehlička & Edwards35; Schuster et al.37; Gieroba et al.38	
V(C–C)	1192	Edwards et al.41	

Results

Raman spectroscopy analysis–tentative peak interpretation

Tayassu samples

Raman spectroscopy analysis of the thirteen Tayassu samples revealed a signature pattern that comprised a high fluorescence and several bands related to organic compounds between 1000 and 1800 cm−1 spectroscopy range (Fig. 5). While examining the fossil teeth, the phosphate signature presented a relatively low intensity concerning hydroxyapatite v1 PO4−2. In some cases, the phosphate peak is almost undetectable resulting from greater fluorescence and the high intensity of the organic compounds. The hydroxyapatite vibration modes (v1, v2, v3, and v4) appear only in a few parts of the teeth.Fig. 5 An example of curve fitting and bands assignments. (A) Extended Raman spectrum without treatment, showing the high fluorescence signal and (B) Curve fitting and deconvolution showing the bands assignments. The figure was created using Microsoft Excel and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

The spectral deconvolution revealed that the fingerprint of the Tayassu samples is comprised of thirteen bands, most of them assigned as biomolecules (Fig. 5). The first peak appears at 962 cm−1 and represent the hydroxyapatite v1. However, the most characteristic peak of this spectroscopy signature appears close to ~ 1287 and ~ 1543 cm−1, which are assigned as protein amide III and C-H bend protein, or N–H2, C = C stretch. The first biomolecule band appears at ~ 1117 cm−1. This peak was assigned as C–C unsaturated fatty acids, (COC), glycosidic bonds, appearing clearly in samples 1, 2, 3, 4, 6, and 12 (Fig. 7, and Supplementary material); in samples 5, 7, 8, 9, 10, and 11, this peak looks like a shoulder (Fig. 8, and Supplementary material). The peaks ~ 1190, 1223, 1262 cm−1 appear as a shoulder in all samples except for samples 7, 8, 10, and 11 (Fig. 8, and Supplementary material), which is easily recognized. These bands are assigned as carbohydrates, mainly C–C, C–O, = C–H in-plane and = C δ(CH), lipids, and proteins respectively (Fig. 5). In all samples, the 1319 cm−1 peak appears as a shoulder with medium or strong intensity, although in samples 1, 2, and 7, it looks more like a peak. We attributed this peak (1319 cm−1) to C–H bend protein24. Next, we have a shoulder comprising two peaks, a strong one at ~ 1403 cm-1 and a small peak at ~ 1444 cm−1 (Figs. 5, 7, 8). We could not find the origin of the 1403 cm−1 peak, but the 1444 cm−1 is attributed to C-H2 deformation lipids38. Following, we have a small shoulder at ~ 1616 cm−1 present in all samples (Fig. 5 and Supplementary material), which is attributed to the amide I band, tyrosine, or phenylalanine28,29,38 . Finally, the last shoulder is composed of two peaks (Fig. 5). The first has a weak intensity at ~ 1681 cm−1, and the second has moderated intensity at ~ 1717 cm−1. These two peaks were attributed to peptide or C = O stretch and C–C, respectively8,38. All spectra show the same pattern (Figs. 7, 8, and Supplementary material). However, the frequency of these thirteen bands has a blueshift or redshift depending on the site point analysis, also varying with the samples.

Despite the fossil samples being found in a carbonate cave, we did not observe peaks related to calcium carbonate minerals of diagenetic origin (v1–CO32− 1085 cm−1). Although it is know that the high fluorescence and organic signatures can obliterate the carbonate peak, as occurs with the phosphate matrix.

In the Raman experiments, we found micro features on the samples, as shown in (Fig. 6). Some samples have features with black pigments, like manganese oxide precipitation (Fig. 6E,L,M). Others have reddish sediments filling holes and fissures (Fig. 6D,G,I,K), besides some white regions with no pigments (Fig. 6D,E,G,I–K,M), at least visible under the light microscope. Figures 7 and 8 show the normalized spectra of these distinct features. In sample 3 (Fig. 6A, B), the vibration modes of hydroxyapatite are easily recognized just in point analysis 3 (Fig. 7A). At points 1 and 2 the hydroxyapatite v1 peak appears weak relative intensity. There is a high intensity of organic compounds.Fig. 6 Typical micro features variability found on Tayassu samples and point analysis. (A,B) Sample 3, there is the reddish sediment attached to the sample (A) and a recent crake (B); (C,D) (E) Sample 5, C and D show can observe the reddish sediment filling the fissures and holes; Fig. 5E shows the black pigment similar to manganese oxide. (F,G) Sample 6. (H,I) Sample 8. (J, K) Sample 10. (L,M) Sample 13, there is a black pigment principal on enamel. The red dots are the point analysis, and the number represents the ID of the point. The Raman spectra appears in (Figs. 7, 8). The white bar in the bottom of the image (imagens D, E, G, I, K, and M) represents 200 µm. The figure was created using Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

Fig. 7 Typical raman signature found on Tayassu samples. (A) Spectra from sample 3, showing the high intensity of biomolecules on points 1 and 2; at point 3 there is only hydroxyapatite signals. (B) Spectra from Sample 5 showing goethite in reddish sediments and lower biomolecules signal on white region. (C) Spectra from Sample 5 taken from black and white regions. (D) In Sample 6, the measurements were taken from white regions. (E) In Sample 6, the Raman data are from the reddish sediments filling the fissures, the hydroxyapatite signal is undetectable. (F) In Sample 8 the spectra were taken from a white region on dentin. Hap – Hydroxyapatite, Gh – Goethite. The figure was created using Microsoft Excel and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

Fig. 8 Typical Raman signature found on Tayassu samples. (A) Sample 8 – The spectra were taken from the border of dentin and enamel; there is a strong signal of biomolecules on reddish sediments. (B,D) Sample 10. Image B shows the spectra taken only from reddish sediments, whereas (C) shows the data from the white region, showing only hydroxyapatite signals. (D) Sample 13 shows a high fluorescence at points 6 and 7, taken from the black and yellow region on the sample. Hap–hydroxyapatite. The figure was created using Microsoft Excel and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

Sample 5 (Fig. 6C) shows microsites with reddish and black pigments (Fig. 6D, E). The Raman analysis reveals the same pattern of biomolecules signature; however. the hydroxyapatite v1 (960 cm−1, P–O symmetric stretching) is absent in point analysis 1a, 1b, and 1c (Fig. 6D and 7B). The absence of v1 is most likely due to the thickness of reddish sediment which does not allow the ion beam to reach the phosphate matrix. Besides the biomolecule signature, the spectrum of the 1a point reveals the goethite mineral (Fig. 7B). In the point analyses 2a and 2b (Fig. 6D and 7B), the hydroxyapatite v1 is strong and easily identified, as opposed to points 3a, 3b, 3c, and 3d (Fig. 6E), where the hydroxyapatite v1 is almost undetectable (Fig. 7C). The point analysis 3b shows a black pigment like manganese oxide (Fig. 6E); however, the manganese was not found in Raman analysis (Fig. 7C) possibly due to the high fluorescence signals. In sample 6 (Fig. 6F, G), the hydroxyapatite v1 shows weak intensity on the white regions (Fig. 7D). By analyzing the reddish clays that fill the fissure, the hydroxyapatite was undetectable, appearing only a small peak in the point analysis 2b (Fig. 6G and 7E). Otherwise, the biomolecule signature and fluorescence show a high intensity.

The same pattern observed for samples 3, 5, and 6 appears in other samples. The dentin of sample 8 shows a high intensity of hydroxyapatite v1 (point analysis 1, 2, 3; Fig. 6H,I and Fig 7F and more 8A). Except for points 4 and 9, which were taken from a white region, all others (point 5, 6, 7, 8; Fig. 6I) were collected from reddish sediments that fill a fissure on the teeth surface. The reddish features show a low intensity of hydroxyapatite v1 and a high signal of biomolecules (Fig. 8). In sample 10 (Fig. 6J, K) the biomolecules signal is stronger on dentin, even if derived from a white region, the enamel points show a high signal of hydroxyapatite, with all vibrational modes; in addition, the biomolecules signature is undetectable (Fig. 8B, C). In Sample 13, the vibrational mode of hydroxyapatite appears on a white region of dentin and enamel (Fig 6L,M and Fig 8 D) (Fig. 6 L,M and Fig 8D) at points 1, 2, 3, and 5. There is a high intensity of biomolecule signatures and fluorescence in the black and yellow region of Sample 13 (points 6 and 7 in Fig. 6M), where the phosphate signature is almost unnoticeable (Fig. 8D).

Smilodon populator samples

The teeth samples from Smilodon have similar features to those found on Tayassu (Fig. 9). The fossil teeth have fissures filled with reddish sediments (Fig. 9 A, B), black pigments like manganese oxide impregnation (Fig. 9 C,D,H), and white regions with no pigment were found (Fig. 9 B,D,F,H,I). Figure 9 illustrates the typical features and Raman signatures of the six Smilodon samples and 65 Raman point analyses. The Raman spectrum displayed the same pattern as the Tayassu samples.Fig. 9 Typical Raman signature and features found on Smilodon samples. (A) In Sample 1, the red box appears in image B. (B) Red box in detailed is shown in image A, with a reddish sediment filling a fissure. (C,E) Sample 2 – the red boxes correspond to images D and F. (D) Detail of the region of interest in image C. There is yellow and black pigments on the sample. (F) Red box in detail is shown in image E, showing that the region of analysis is free of red and black pigments. (G) Sample 3 –Red box in detail is shown in image H. (H) The region of analysis of sample 3, showing the black and yellow pigments. (I) Sample 4 – the measurements on the recently cracked region. (J) Raman spectrum from a white region in sample 1, image B. (K) Raman spectrum from reddish sediment in Sample 1, image B. (L) Raman spectrum from black pigments of sample 2 at image D. (M) Raman spectrum from a white region in Sample 2, image F. (N) Raman spectrum from black pigments on Sample 3, image H. (O) Raman spectrum from a white region in Sample 4, image I. – Hap – Hydroxyapatite. The red dots are the point analysis, and the number is the ID of the point. The white bar at the bottom of the images (images B, D, F, and H) represents 200 µm. The figure was created using Microsoft Excel and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

The signatures of biomolecules are stronger on the reddish sediments than fill fissures and holes (Fig. 9A, B), which also occurs for the black pigments on the sample surface (Fig. 9L, N), while the signature of hydroxyapatite is almost undetectable (Fig. 9J,N). However, bands relating to biomolecules are also as strong as the same bands in the white regions (e.g., Fig. 9K). The white regions show a moderate signal of hydroxyapatite, along with a strong signature of biomolecules (Fig. 9C,G,H,M). However, point analysis on the recent cracks or broken faces (Fig 9E,F and more Fig 9I) reveals a very weak, or absent, of biomolecules signatures (Fig. 9M, O).

Discussion

Our findings align with earlier reports of organic signatures from fossil samples4,14,27,29. Piga et al.27 observed several bands between 1200 to 1800 cm−1 spectroscopy range in dinosaur bone from Spain, thus suggesting an organic origin. Barros et al.29 found comparable bands in a shrimp fossil from the Araripe Basin (Cretaceous) in Brazil, indicating a graphitic-related material, de Sousa et al.4 reported a several peaks within the same range from a Pleistocene fossil from a limestone cave in Brazil’s Diamantina range. Jurašeková et al.14 recorded the same findings when studying different fossil tissues from vertebrates, spanning the Jurassic to the Cretaceous period. Conducted in a varied environmental context, these studies suggest that the Raman signature found may represent a recurring pattern in certain fossilized tissues, whose implications are not yet fully understood.

We have two hypotheses that could explain the existence of these organic signatures on fossil bones: (i) these biological signatures are linked to preserved fossil biomolecules, such as found in dinosaurs26 or (ii) the biological signature is related to biofilm formation during the post-depositional process. Our experiment design provides an outstanding potential to understand the spatial distribution of these biological signatures and evaluate both hypotheses. If these signatures are related to teeth fossilized biomolecules, as reported by Wiemann et al.26, we expected to find them preserved mainly at the innermost part of the teeth, since they would be more protected from diagenetic processes. If these spectral signatures are not related to the individual when alive, they would be related to the post-depositional processes. Therefore, we could find this spectral signature related to the different post-depositional features (Figs. 6, 7, 8, 9).

Based on the results of the 134 Raman spectra, we might suggest that the inner part of the teeth (the new broken faces) is virtually absent of biological signature (Figs. 4, 5 and 6). It is worth highlighting that out of the 134 Raman spectra, only 13 were made in areas considered to be inside the teeth. However, prior studies reported the existence of this spectral signature inside bone tissues4,14. De Sousa et al.4 studied vertebrae of Pleistocene ground sloth found in limestone cave and found this signature with a strong relative intensity concerning the hydroxyapatite peak, on the outer surface, decreasing until disappearing at the innermost part of the bones. Jurašeková et al.14 report the same findings pointing out that enigmatic peaks occur preferentially in the outermost part of the bone tissues, despite being found in both outer and inner parts.

Our Raman results reveal an organic composition of = C–C unsaturated fatty acids or glycosidic bonds (~ 1117 cm−1), carbohydrates, mainly C–C and C-O (~ 1190 cm−1), = C–H in-plane (~ 1223 cm-1), = C δ(CH), lipids, proteins (~ 1262 cm−1), Protein, mainly amide III (~ 1287 cm−1) C–H bend protein (~ 1319, and ~ 1543 cm−1), CH2, saturated lipids (~ 1444 cm-1), C–H bend protein, N–H2 C–H band, C = C stretch (~ 1543 cm−1) amide I, tyrosine, phenylalanine (~ 1616 cm−1), peptide, C = O stretch (~ 1681 cm−1) C = O stretch, C–C (~ 1717 cm−1)27,31–35,37,39–42. This composition is reported for biofilms and exopolymeric substances (EPS), which comprise polysaccharides, lipids, fatty acids, proteins, nucleic acids, and carbohydrates27,31,33–36. It is worth noting that teeth and bone organic tissues tend to exhibit stonger amide I signals from proteins43,44, while amide III signals appear at lower intensities, which is in contrast with our results that reveal amide III at strong intensities and amide I only as a small shoulder (Figs. 5,7,8 and 9). This suggest that the amide I and III signals  observed in our samples are not associated with organic matrix in the fossil tissue. Therefore, this help to support our interpretation that comprise the organic signature found on the outer face in all fossil samples can be explained as biofilm deposition, as the second hypothesis predicts.

Biofilms are the main form of microbial life association on Earth33 and represent a collective way of living for microorganisms that can reach a few mm in thickness, with spatially organized microbial cells within a self-produced matrix EPS. The EPS is composed of proteins, lipids, polysaccharides, and extracellular DNA45. Biofilms are resistant to extreme environments, and work as protective clothing13,14 for the microorganisms against ultraviolet (UV) radiation, extreme temperature, extreme pH, high salinity, and other extreme conditions. The existence of fossilized biofilm and biofilm on fossils is well known20,23,46–49, and the importance of biofilm to preserving soft tissue in the fossil record has been recent demonstrated25,47–50.

In the case of our study, the biofilm formation on fossil bones tissues can be explained by the oligotrophic condition of the cave sediment matrix (Fig. 10). The fossil mineral and organic matrix decomposition will be an important source of nutrients. Studies51–53 about chemical composition of teeth (enamel and dentin) show that P, N, Ca, Mg, Na and S are the most abundant elements. While enamel is the most mineralized bone tissue composed of 95%wt of hydroxyapatite and 1%wt of organic matrix (proteins and lipids)53–56, the dentin shows 70%wt of mineral phase and 20%wt of organic matrix55. Nitrogen and organic carbon were higher in dentine than in enamel with 1.94% to 2.93% for nitrogen and 6.34% to 11.5% for organic carbon53. The microorganisms existing in sediment likely will be fixed on the fossil (Fig. 10A) and attracted to the eutrophic condition prevailing on the fossil surface due to diagenesis51,53,56.Fig. 10 Model of microbial biofilm formation: Biofilm formation consists of five distinct steps: (A) Attachment, the planktonic state: microbial organisms adhere to the bone surface with weak bound, most likely van der Waals forces; (B) Colonization: microbial organisms attach to the surface almost irreversibly via stronger hydrophilic/hydrophobic interactions and produce extracellular polymeric substances (EPS). (C,D) evelopment of multilayers: as nutrient consumption increases the auto-induced production and the biofilm passes to development of multilayers. (D) Mature and dispersion: stable formation of a three-dimensional community. Microorganisms are disseminated from the aggregate biofilm and return to a planktonic state. (E) Degradation of biofilm: natural anti-biofilm agents most likely require specific enzymes include in degrading and reconfiguring biofilm, resulting not simply in partial matrix degradation. This general model was draw by Daniel V. Sousa made based on Boles and Horswill58 theoretical model. The figure was draw by hand using the software Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

The biofilm formation starts with the interaction of the microorganisms and fossil surface, an interaction that is reversibly adsorbed to a surface due to weak bound, most likely van der Waals forces49. As bone colonization progresses, the EPS matrix also increases between microorganisms and fossil surface, thus nutrient consumption will increase (Fig. 10B). In this stage, the interaction of the microorganism and fossil surface become almost irreversibly via stronger hydrophilic/hydrophobic interactions by flagella, pili, lipopolysaccharides, exopolysaccharides, and collagen-binding adhesive proteins25,50. As nutrient consumption increases the auto-induced production and the biofilm passes to a development of multilayers (Fig. 10C), latter the maturation and dispersion of the microorganisms occur (Fig. 10D)57. This general model, based on Boles and Horswill studies58, explain the existence of biofilm on fossil bones; however, as the fossil was found in a cave environment the post-depositional process plays a key role in the formation of the biofilm matrix.

Post-depositional processes in the cave environment depend on various site conditions, such as sediment mineralogical composition, pH, temperature, and water characteristics1,23. In the case of our study the sediment geochemistry, mineralogical assemblage, pH, and temperature do not change significantly4. Water condition is the only feature that changes through the stratigraphy. De Sousa et al.4 studied the sediment of the Toca de Cima do Pilão cave and report many redoximorphic features in the lowermost layers, pointing to water table fluctuation. The layers dated after 103.5 kyr show more intense redoximorphic features than the layer dated before 82.4 kyr. The redox is responsible for generating post-depositional features on the fossil bones; for instance, impregnation by iron oxides and even clayey material filling the voids, fractures, and breaks, which explain the presence of goethite in our samples (Fig. 7b). However, the most prominent post-depositional features are the manganese oxide impregnation. Our samples show many black pigments, like manganese oxide precipitation (Figs. 6E, 6L, 9D, 9H); however, our Raman spectrums do not attest the presence of Mn oxide. These results may suggest that black pigment was formed due biofilm action, with few or no relationship with the redox process, pointing out that black pigments commonly found on fossilized materials are not always related to oxidoreduction of manganese oxide. Or else, the fluorescence of the biofilms obliterated the manganese oxide signal, which is more likely to have occurred.

Considering that the peaks between 1200–1800 cm−1 spectral range are linked to biofilm and develop possibly due to greater fluorescence, a ratio between the v1 hydroxyapatite peak vs. the peaks of the highest intensity of organic compounds (Amide III and N-H2), would estimate the greater or lesser biofilm formation in each point analysis. In addition, an analysis of the ratios of v1 hydroxyapatite vs. amide III and hydroxyapatite vs. N-H2 in a stratigraphic sequence, would allow us to observe the effect of depth and/or post-depositional processes on biofilm formation. As a result, there is a clear relationship depth and biofilm production (Fig. 11). Figure 11A, the Precari teeth samples show the amide III and N–H2 raise in deep, also Smilodon samples (Fig. 11B). However, we suggest that such a result does not have any relationship with the long-term fossilization or the age of the fossil, but most likely there is a link with the post-depositional process, mainly associated with water action. Figure 11C shows the stratigraphic sequence and predominance of redoximorphic features, revealing that the ratio of hydroxyapatite/amide III drastically decreased after 82,4kyr, exactly where the features become more marked in the stratigraphic profile. It is worth highlighting those redoximorphic features were linked to water action4. The water favors the microorganism in the locomotion and the fossil organic phase decomposition. Moreover, microorganisms produce organic acids as endmembers of the decomposition process, this acidification contributes to promoting the dissolution of hydroxyapatite and the organic matrix.Fig. 11 Post-depositional process on biofilm formation. (A) Ratio of Hydroxyapatite vs. Amide III through the deep. (B) The ratio of Hydroxyapatite vs. N-H2 through the deep. (C) Stratigraphic sequence, geochronology, and oxidoreduction features. Chronology data collected from de Sousa et al.51. The sizes of the reddish circles on graph C indicate the intensity of the oxidoreduction process, and the white circle indicates the absence of redoximorphic features. The black paw on graphs A and B indicates the data of Smilodon (orange dots), and the little pig indicates the data of Tayassu (blue dots). The stratigraphic and geochronological data were collected from de Sousa et al.4 . The figure was created using Microsoft Excel and Microsoft Power Point (https://www.microsoft.com/pt-br/microsoft-365/).

Following the biofilm hypothesis, a question arises: Is this fossil biofilm or is it a recent biofilm? Comparing our data with the literature on biofilm, we might suggest that they are fossil biofilms. Despite the similar composition, our Raman spectra differ from those reported in the literature for recent biofilm32,34,38,41. Our spectrum does not show nucleic acids. Furthermore, the peaks found in this study show a blueshift or redshift. The amide III band shows this low position at 1269 cm−1 and a high position at 1295 cm−1, while the literature reports it with a position at 1262 cm−1; N–H2 shows a low position at 1493 cm−1 and the high position at 1544 cm−1, while the literature reports it with a position at 1538 cm−1. These shifts could be explained by biofilm degradation or the generated of mineral bind biomolecules. The process of biofilm degradation requires specific enzymes included in degrading and reconfiguring biofilm, resulting not simply in partial matrix degradation but also in the active dispersal of biofilm and subsequent surface recolonization34. Our study does not cover the specific mechanism that involved the natural anti-biofilm agents in fossil records.

Conclusions

This study demonstrate that the Raman signature (bands between 1200 to 1800 cm−1 spectral range) commonly found in many fossil records is linked to biomolecules from degraded biofilm. We show that through a deeply Raman spectroscopy investigation on Quaternary fossils found in Toca de Cima do Pilão cave, Brazil.

Our experimental design is the first to examine biomolecules from biofilms from fossils along a well-dated and well-known stratigraphic profile. We demonstrate the relationship between biofilm and the post-depositional process by analyzing the ratio of hydroxyapatite v1/amide III across the stratigraphic sequence. This approach allowed us to suggest that the increased biofilm production at depth is unrelated to the age of the fossil but rather to post-depositional processes. These biofilms show a spatial distribution associated with post-depositional features on the teeth surface (reddish sediments and black pigments). Moreover, these organic compounds were almost exclusively found on the teeth’s outer surface.

We propose a general model based on Boles and Horswill studies58 to explain the existence of biofilm on fossil bone tissues buried in a siliciclastic sediment matrix. In the case of our study, the biofilm formation on fossil teeth is explained by the oligotrophic condition of the cave sediment matrix. Afterward the maturation and dispersion of the microorganisms the biofilm becomes to degrade. However, our study does not address the specific mechanism that involves the natural anti-biofilm agents and biofilm degradation in fossil records. As the fossil were found in a cave environment the post-depositional process drives the biofilm formation and the water action plays a key role on biofilm development. This study opens new avenues in paleontology, biofilm, and even astrobiology studies, by proposing for the first time, that type of Raman signatures commonly found in a broad of fossil records originated from biofilm production.

Supplementary Information

Supplementary Figure S1.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-024-71313-6.

Acknowledgements

The authors would like to thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Fundação de Amparo a Pesquisa do Estado de Pernambuco (FACEPE) for funding the research of Daniel Vieira de Sousa (Research grant CNPq 432075/2018-6; 102032/2024-6; FACEPE APQ-0936-21-52799) and the Universidade Federal do Vale do São Francisco, Brazil, for supporting the fieldwork of this research [PRPPGI PC 860/2017; PRPPGI PC 1610/2018]. We would also like to thank the Instituto do Patrimônio Histórico e Artístico Nacional–IPHAN (nº 8/2019/ETSRN-PI/IPHAN-PI), Centro Nacional de Arqueologia–CNA (196.2019 ACON/CNA/DEPAM) and the Agência Nacional de Mineração–ANM (ANM nº 48051.003438/2019-99.), for providing the required authorizations for the adequate execution of this research. Additionally, we are deeply indebted to the Fundação Museu do Homem American (FUMDHAM), which provided us with access to the material deposited in its paleontological museum and Dra. Janaína Carla Santos from UNIVASF who help us during the field work. We would also like to thank the anonymous reviewers for refining the paper.

Author contributions

D.V.S., P.V.S.M, L.M.G, E.E. conceived the experiments, and designed the study. P.V.S.M, D.V.S. L.M.G performed the experiments and data analysis of Raman Spectroscopy. D.V.S, P.V.S.M. figure generation with contribute from all authors. D.V.S. drafted the manuscript with input from all authors.

Data availability

Sequence data that support the findings of this study have been deposited in the Mendeley data repository (https://data.mendeley.com/datasets/nscghhnt63/2).

Competing interests

The authors declare no competing interests.

Publisher's note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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