# Coupling gel electrophoresis with photoluminescence reveals biochrome complexes in modern and fossil shells.

Luca Polacchi, Camille Lutet-Toti, Olivier Basuyaux, Pierre Gueriau, Marie Albéric, Hélène Pasco, Jérôme Thomas, Didier Merle, Benoît Habermeyer, Roger Guilard, et al.

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## To cite this version:

Luca Polacchi, Camille Lutet-Toti, Olivier Basuyaux, Pierre Gueriau, Marie Albéric, et al., Coupling gel electrophoresis with photoluminescence reveals biochrome complexes in modern and fossil shells. Methods in Ecology and Evolution, 2025, 16 (12), pp.2807-2818. 10.1111/2041-210X.70176. hal-05384818

HAL Id: hal-05384818

https://ube.hal.science/hal-05384818v1

Submitted on 27 Nov 2025

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Distributed under a Creative Commons Attribution 4.0 International License
# Coupling gel electrophoresis with photoluminescence reveals biochrome complexes in modern and fossil shells

Luca Polacchi<fnref n="1" /> | Camille Lutet-Toti<fnref n="2" /> | Olivier Basuyaux³ | Pierre Gueriau<fnref n="1" />,⁴ |

Marie Albéric⁵ | Hélène Pasco<fnref n="1" /> | Jérôme Thomas<fnref n="2" /> | Didier Merle⁶ |

Benoît Habermeyer⁷ | Roger Guilard⁷,⁸ | Mathieu Thoury<fnref n="1" /> | Frédéric Marin<fnref n="2" />

**Correspondence**

Frédéric Marin

Email: frederic.marin@u-bourgogne.fr

**Funding information**

CNRS Tellus-INTERRVIE (INSU), MAELSTROM (2022); FishMed-PhD program, PhD Fellowship Camille Lutet-Toti; OSU-Theta, SRO project PRELUDE; DIM PAMIR (PAtrimoines Matériels - Innovation, expérimentation et Résilience), PRESERV project

Handling Editor: Natalie Cooper

**Abstract**

1. Polyacrylamide gel electrophoresis is commonly used to visualize mixtures of proteins, such as those embedded in calcareous biominerals. However, it is ineffective for the detection of biochromes, a major class of low molecular weight organic compounds commonly associated with calcified exoskeletons.

2. We describe a novel approach based on the coupling of electrophoresis and photoluminescence (PL) spectral imaging to reveal invisible biochromes, identify them chemically, provide evidence for their putative interaction with exoskeletal macromolecules and purify them in large amounts. Our protocol relies on three key-steps: a mild extraction of all organics from bleached skeletal powder (step 1); an optimized electrophoretic fractionation immediately followed by direct on-gel spectral image acquisition performed before classical gel staining (step 2); a large-scale purification via preparative electrophoresis coupled to PL spectral imaging, to obtain significant amounts of biochromes of interest (step 3).

3. Steps 1–3 were successfully applied to recent gastropod shell extracts, while steps 1 and 2 were applied to their fossil equivalents from the Eocene epoch (≈approximately 45 million years). In both cases, we were able to identify porphyrins associated with the shells via a spectrophotometric measurement of the luminescent bands directly on the analytical gel, even at low concentration in the case of fossil extract. For recent shell extracts, our approach allowed us to identify spectrophotometrically the tubes containing the invisible pigments, after preparative fractionation and fraction collection.

4. Our protocol enables direct on-gel identification of porphyrin molecules, even in trace amounts. It opens up new avenues for studying a wide range of biological

This is an open access article under the terms of the [Creative Commons Attribution License](https://creativecommons.org/licenses/by/4.0/), which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

© 2025 The Author(s). Methods in Ecology and Evolution published by John Wiley & Sons Ltd on behalf of British Ecological Society.

<footnote n="1">¹IPANEMA, CNRS, Ministère de la Culture, UVSQ, MNHN, UAR3461, Université Paris-Saclay—Site du Synchrotron SOLEIL, Gif-sur-Yvette, France;</footnote>

<footnote n="2">²Biogeosciences Laboratory, UMR CNRS-EPHE-UBE 6282, University Burgundy Europe, Dijon, France; ³M2e, Mer Expertise Environnement, Blainville-sur-</footnote>

<footnote>Mer, France; ⁴Institute of Earth Sciences, Geopolis—University of Lausanne, Lausanne, Switzerland; ⁵LCMCP Laboratory, Sorbonne University, Paris Cedex 05,</footnote>

<footnote>France; ⁶Department Origines & Evolution, MNHN, Sorbonne University, Paris Cedex 05, France; ⁷PorphyChem SAS, Longvic, France and ⁸ICMUB, UMR CNRS</footnote>

<footnote>6302, University Burgundy Europe, Dijon, France</footnote>
composites, whether mineralized or not, that contain photoluminescent biochromes. It is particularly well-suited to ancient specimens and fossils to trace the origin and evolution of biochrome complexes in the geological record.

### KEYWORDS

biomineral, electrophoresis, fossil, multispectral imaging, pigment, porphyrin

# 1 | INTRODUCTION

Calcium carbonate-based biominerals (e.g. coral exoskeletons, mollusc shells, sea urchin spines) are organo-mineral assemblies that contain small amounts (typically 1% wt/wt or less) of organic macromolecules—collectively called the skeletal matrix—which regulate mineral deposition (Addadi & Weiner, 2014; Marin et al., 2016; Suzuki, 2020). A substantial proportion of biominerals contain biochromes in addition (Amarowicz et al., 2012; Bergamonti et al., 2013; Comfort, 1949, 1951; Soldati et al., 2008; Williams, 2017). Biochromes are small organic molecules, showing absorption in the visible spectral range, produced by living systems. They are composed of a diversified set of chemical structures, from linear chains with conjugated double bonds, such as carotenoids (Beltran & Wurtzel, 2025), to complex structures involving rings and metal ion complexation, like porphyrins (Kadish & Guilard, 2024). They generate colour by absorbing light at specific wavelengths due to their specific molecular structures (Hari et al., 1994). These pigments can be detected in situ, that is directly on the shell, via non-destructive methods (UV light, Raman spectroscopy) that are easy to implement (Caze et al., 2015). However, these approaches analyse shell biochromes in bulk. Thus, at their best, they will give an overall spectral signature that is the sum of the spectral signatures of each biochrome taken individually, according to their concentration in the skeletal tissue. The resulting spectrum may be difficult to interpret and clearly not discriminant enough. Deepened characterization requires the extraction of these colouring molecules from calcified tissues. Common protocols rely primarily on harsh extraction with strong acids to dissolve the mineral fraction (Bonnard et al., 2020; Hou et al., 2019). This approach is practical for quickly obtaining large amounts of biochromes. However, beyond using harmful and environmentally unfriendly chemicals, it irreversibly degrades the skeletal macromolecules and consequently disrupts the biochrome-macromolecule complexes. It is therefore unsuitable for disentangling biochrome-macromolecule interactions and studying the persistence of organics in ancient or fossil biominerals (Caze et al., 2015).

Another difficulty is getting an accurate 'instantaneous' picture of all organic components, that is biochromes and skeletal matrix, together. While gel electrophoresis—referred to as Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis, abbreviated as SDS-PAGE—is the most convenient technique for visualizing macromolecules from biominerals and determining their size distribution (Immel et al., 2016; Osuna-Mascaro et al., 2014), it is ineffective in revealing associated biochromes for three reasons: (a) this technique is not discriminatory enough for small organic molecules around and below 1kDa, typically a molecular weight range of biochromes that are not bound to macromolecules; (b)

in addition, when unbound, biochromes diffuse extremely rapidly out of the gel (and are consequently lost) because of their small size, as soon as the gel is processed for staining; (c) when bound to matrix macromolecules, biochromes may be stabilized in the gel via a fixation step (with trichloroacetic acid, i.e., TCA, for example) but some of them are invisible or, at best, only weakly coloured in natural light; as biochromes do not take on any of the classical gel dyes (Coomassie Blue, silver nitrate), they are irreversibly masked by subsequent gel staining (Pavat et al., 2012). The novel protocol developed here circumvents these three obstacles: in particular, it reveals invisible biochromes or mixtures of biochromes, identifies them chemically via direct on-gel spectral acquisition and evidences their putative complexation with macromolecules. At last, it allows obtaining a considerable amount of purified biochromes when the starting shell material is abundant.

# 2 | MATERIALS AND METHODS

## 2.1 | Materials

Our study focused on two shelled gastropod species that belong to the same superfamily (Trochoidea), according to the World Register of Marine Species (WORMS) database (Ahyong et al., 2025): the extant *Phorcus lineatus* (da Costa, 1778) (Gastropoda, Vetigastropoda, Trochida, Trochoidea, Trochidae); the fossil *Tectus crenularis* (Lamarck, 1804) (Gastropoda, Vetigastropoda, Trochida, Trochoidea, Tegulidae). Two dozen shells of *P. lineatus* were manually sampled along the Normandy coast near Blainville-sur-Mer by one of us (O.B.). This extremely abundant species is not endangered, not listed in the Washington Convention and consequently was not the subject of any request to an animal ethics committee. Emptied specimens were carefully sorted visually: shells with no defects and no epibionts were selected. Five fossil shells of *T. crenularis* of the middle Lutetian (≈45 million years) of the Paris Basin were used for the analysis: they came from the invertebrate fossil collection of the Natural History National Museum of Paris.

## 2.2 | Matrix extraction

The shells were scrupulously brushed, rinsed and carefully decontaminated for several days in a tenfold aqueous solution of sodium hypochlorite (0.6%-1.4% active chlorine, Merck ref. 1.05614.2500). After thorough rinsing with ultrapure water and drying, they were coarsely crushed and bleached again before being mechanically comminuted
(Pulverisette 2, Fritsch) to a powder with a particle size of less than 200µm, obtained by sieving. The powder was suspended in ultrapure water and slowly demineralized overnight with cold dilute acetic acid, according to a published protocol (Immel et al., 2016; Osuna-Mascaro et al., 2014). The resulting solution was centrifuged (30min, 3900 G) to separate the insoluble and soluble organic fractions. The insoluble pellet (defined as AIM, 'Acetic acid-Insoluble Matrix') was rinsed at least five times by cycles of suspension in ultrapure water, centrifugation and removal of the supernatant. For *P. lineatus*, the soluble fraction (ASM, 'Acetic acid-Soluble Matrix') was concentrated in a 400mL ultrafiltration stirred cell (Millipore, model 8400, ref. UFSC40001) equipped with a cellulose membrane with a 10kDa cut-off (Millipore 76mm diameter disc, ref. PLGC07610). The viscous retentate (about 10–15mL) was kept while the collected filtrate was ultrafiltered again on a membrane of 1kDa cut-off (Millipore 76mm diameter disk, ref. PLAC07610). The two retentates (ASM >10kDa and 1<ASM<10kDa) were extensively dialyzed against ultrapure water in dialysis tubing of 1kDa cut-off (SpectraP/Por 6 pre-wetted RC tubing) with at least 6 water changes. The insoluble and the two soluble organic matrices were freeze-dried (Telstar Cryodos). For the fossil *T. crenularis*, the insoluble AIM pellet contained clay minerals and was not used further. Since the ASM supernatant resulting from the decalcification was not concentrated in organics, we performed a single ultrafiltration in the Millipore model 8400 ultrafiltration cell equipped with a 1kDa cut-off membrane, all the other subsequent steps being identical to those of *P. lineatus*.

**2.3 | Analytical gel electrophoresis and acquisition of spectral images**

For visualizing the result of the extraction, the standard electrophoretic mini-gel system (Bio-Rad, Mini Protean III) was used. To improve the resolution of the low to very low (<1kDa) molecular weight components, we combined the use of discontinuous Tris-tricine gels run with anode and cathode buffers (Schägger & von Jagow, 1987) and a high percentage (15%) of acrylamide. Samples were prepared and denatured in Laemmli sample buffer as previously described (Immel et al., 2016). Note that the AIM of *P. lineatus* was only partially solubilized in this buffer. The soluble portion of this fraction is referred to as the Laemmli-soluble-AIM (LS-AIM). All fractions were run at 120V. The use of a pocket UV torch (NS UV 395 model, NightSearcher, UK) allowed the migration of the invisible pigments to be checked preceding the migration front and the run to be stopped when the pigment reached the bottom of the gel. The gel was then de-cast, placed on a flat clean support, and multispectral images were acquired by using the home-built imaging system, as described in Figure S1. The detection part consisted of a 4 megapixels CMOS camera (ORCA-Flash 4.0 LT Plus, Hamamatsu) with a sensitivity ranging from 350 to 1100nm. The camera was equipped with a UV-VIS-IR 60mm 1:4 Apo Macro lens (CoastalOptics), in front of which was positioned a filter wheel fitted with 8 interference bandpass filters (Semrock) to perform multispectral acquisitions. The illumination part was composed of 16 LED lights ranging from 365 up to 700nm (CoolLED pE-4000), coupled to a liquid

light-guide fibre fitted with a fibre-optic ring light-guide to allow homogeneous illumination. Emission and excitation spectra were collected using a modified spectrofluorometer (Fluorolog 3-22, HORIBA Jobin Yvon) which allows spectra to be collected directly on the gels using an optical bundle connected to a focusing lens. The entrance and exit slit widths of the monochromators were set at 10nm to collect spectra with good signal-to-noise and a spectral resolution compatible with the detection of Q-bands. Emission spectra were collected using an excitation at 400nm. Excitation spectra were collected using a 732±34nm bandpass filter (from Semrock) at the entrance of the emission monochromator to eliminate any stray light. The spectra on tube were collected under a UV excitation generated by a Jaxman U1C torch and using a JETI 1211 portable spectrometer sensitive between 300 and 1000nm, fitted with a high-pass transmission filter with a cut-off wavelength at 409 nm to eliminate the reflectance of the UV excitation.

**2.4 | Preparative gel electrophoresis**

As the shell matrix of *P. lineatus* was abundant, we performed preparative gel electrophoresis using the Bio-Rad preparative cell (model 491 Prep Cell) equipped with its cooling unit, its peristaltic pump (EP-1 Econo Pump), its detection cell (EM-1 Econo UV Monitor) and its fraction collector (Model 2110) as previously described (Marin et al., 2001). As for analytical gel, we used 15% acrylamide Tris-tricine gel together with cathode/anode buffers to obtain the best resolution for very low molecular weights. The gel was cast and run according to the manufacturer's instructions (available at https://www.bio-rad.com/webroot/web/pdf/lsr/literature/M1702925.pdf) and the progression of the biochrome's migration was followed from time to time by rapidly illuminating the gel tube with the pocket UV torch. Fractions of 5mL (flow rate 0.5mL/min, 10min per tube) started to be collected continuously after 5h of migration, when the pigment signal reached the bottom of the gel tube. Spectra were directly acquired on the different tubes. The free pigment was eluted from tube 4 to tube 7 (with the highest amounts of materials in tubes 4 and 5) while the migration front (bromophenol blue) was eluted from tube 8 to tube 12.

All multispectral data obtained on analytical and on preparative gels were deposited in the Dryad Digital Repository and are accessible at https://datadryad.org/dataset/doi:10.5061/dryad.rr4xgxdjc (Thoury et al., 2025).

**3 | RESULTS**

**3.1 | A non-standard and optimized protocol in three steps**

As shown in Figure 1, our protocol is based on the association of three key steps, namely a mild extraction, consisting of dissolving slowly mineralized tissues with cold dilute acetic acid, in order to recover all organic molecules embedded in the calcified skeleton and to minimize the possible degradation of biochrome-macromolecule
<figure><img src="image_3.png" /><figcaption>**FIGURE 1** Scheme of the procedure for the analysis of invisible shell biochromes describing the three key-steps (explained in the text): In brief, they involve a mild extraction (upper panel) and the coupling of photoluminescence spectral imaging with mono-dimensional electrophoresis, either analytical on mini-gels (left panel) or preparative (right panel).</figcaption></figure>

complexes (step 1); the non-standard coupling of electrophoretic fractionation (on analytical mini-gels) with a custom-made photoluminescence (PL) spectral imaging, according to a proper sequence (step 2); whenever possible, the adaptation of step 2 to large quantities of shell extracts, by preparative electrophoresis, in order to obtain workable quantities of biochrome (step 3).

Steps 1 and 2 are suited only to minute quantities of skeletal organic matrices, typically extracted from precious samples, including recent but rare materials, archaeological or fossil shell specimens; the full procedure (steps 1-3) can be applied to organic mixtures extracted from abundant raw material (fresh shells).

## 3.2 | Revealing biochromes in gels (Figure 2e,f)

We applied our protocol to shell extracts of two mollusc models, the living *P. lineatus* (Figure 2a,b), an extant gastropod extremely common

along the Atlantic and Normandy coasts (Cabral, 2020) and an equivalent 45-million-year-old fossil specimen, *T. crenularis* (Figure 2c,d), belonging to the same superfamily (Trochoidea) and found in the Lutetian of Paris Basin (Gain et al., 2019). The visualization of biochromes in analytical gels requires two sine qua non conditions: (1) the use of hand-cast unconventional high acrylamide (15, 16%) Tris-tricine gels for increasing the resolution of low molecular weight components, around 1kDa and less. These components include, for example, free biochromes, oligopeptides and oligosaccharides. (2) The acquisition of images in natural light and PL modes, just after gel running/demoulding but before any type of gel fixation and staining. Figure 2e,f illustrate results obtained in these two modes. The left gel (Figure 2e) is an image obtained from the freshly run and unstained gel in natural light. Large diffuse smears of dark brown pigments are visible in lanes 2, 3 and 4, corresponding to different *P. lineatus* extracts, while a very faint orange smear can be observed in lane 5 (*T. crenularis*). Interestingly, when the same gel is illuminated with UV light (Figure 2f), additional biochromes,
<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /><figcaption>FIGURE 2 Analytical SDS-PAGE of shell extracts of the extant *Phorcus lineatus* (Figure 2a,b) and of the 45-million-year-old fossil *Tectus crenularis* (Figure 2c,d). Figure 2e represents the Tris-tricine gel (15% acrylamide) just after migration (no staining), in visible light. Dark biochromes migrate mostly as polydisperse smear in lanes 2 and 3. The migration front is visualized by blue bands in all lanes. Note that some pigments migrate faster than this migration front in lanes 2–4. Figure 2f: False colour photoluminescence image of the same gel, using 385 nm excitation and generated by associating three black and white images (red channel: 708±37 nm; green channel: 571±36 nm; blue channel: 472±15 nm). The image reveals additional biochromes in lanes 3, 4 and 5, mostly at the bottom of the gel. In Figure 2e,f, lanes 2–4 are extracts of *P. lineatus*, while lane 5, that of *Tectus*; lane 1: Pre-stained molecular weight markers (Fermentas, from 11 to 170 kDa); lane 2: Laemmli-soluble/acetic acid-insoluble matrix; lane 3: acetic acid-soluble matrix, fraction >10 kDa; lane 4: acetic acid-soluble matrix, 1 kDa < fraction <10 kDa; lane 5: acetic acid-soluble fraction >1 kDa. Figure 2g: Excitation and emission spectra obtained directly in situ from the electrophoresis bands (of Figure 2f) from *P. lineatus* ASM (pink-red, lane 4) and from *T. crenularis* ASM (purple, lane 5). The emission and excitation spectra were collected using 700 nm excitation and 405 nm detection, respectively. The spectra obtained correspond to uroporphyrins. The developed chemical formula indicated on the right side of Figure 2g are that of uroporphyrin I and III, the most commonly found in natural systems.</figcaption></figure>

almost or totally invisible in natural light, are observed in a zone preceding the migration front (i.e. below the line formed by bromophenol blue) using an optimized multispectral set-up: in the bottom of lanes

3 and 4, they consist of a discrete predominant band, strongly luminescent in the red spectral domain, surmounted by weaker luminescent smearing signals (yellowish to red). The electrophoretic pattern of
lane 5 (fossil) is different and composed of successive discrete bands of weak PL, and comprising three cyan and one light red (underlined by a white rectangle), separated according to their migration properties. Lane 5 is crowned by an upper smearing fat cyan signal.

## 3.3 | In-gel identification of luminescent biochromes (Figure 2f,g)

Our protocol not only reveals the presence of all biochromes (visible and non-visible in natural light) but allows the chemical identification of porphyrinic ones, based on their spectral signature. In the example of Figure 2f, both emission and excitation spectra were acquired in situ from the bottom luminescent band of *P. lineatus* extracts (lanes 3 and 4), using a custom-modified spectrofluorometer. In Figure 2g, the Soret band observed in the excitation spectrum at 408 nm and five Q-bands at 508, 542, 573, 623 and 649 nm (red continuous curve) in addition to three bands at 623, 653 and 685 nm in the emission spectrum (red dotted line) are characteristic of a non-metallated porphyrin (Ji et al., 1990; Williams et al., 2016). A subsequent database search (wwwphotochemcad.com/databases/common-compounds/porphyrins) indicates that the excitation spectrum corresponds either to protoporphyrin IX or to uroporphyrins (Taniguchi & Lindsey, 2018; Toussaint et al., 2023). However, the emission spectrum, with its two peaks at 623 and 653 nm, suggests the second possibility (i.e. uroporphyrin, which peaks are located around 620 and 655 nm) rather than the first one: indeed, protoporphyrin IX emission peaks are slightly shifted (635 and 705 nm) compared to those of uroporphyrin. The definitive identification of the nature of porphyrin in the shell of *P. lineatus* will, however, require extra analyses, such as HPLC or mass spectrometry, far beyond the scope of this paper.

A similar in situ spectrum obtained from the faint red band of the fossil *T. crenularis* (Figure 2f; lane 5, white rectangle) corresponds to a porphyrin that also shows all the excitation and emission features of an uroporphyrin, with excitation bands at 400, 504, 534, 567, 609 and 649 nm (pink continuous line) and emission bands at 617, 648 and 683 nm (pink dotted line). However, it is interesting to note that the two uroporphyrins identified in the extant and fossil samples are not identical: (1) first, they exhibit a different electrophoretic migration pattern, with *P. lineatus* uroporphyrin migrating faster than the fossil one; (2) secondly, one notes a shift of few nanometres (4–8 nm) in the peak values of the emission and excitation spectra of both porphyrins (fossil vs. extant): 400 versus 408, 504 versus 508, 534 versus 542, 567 versus 573 nm for the excitation spectra and 617 versus 623 and 648 versus 653 nm for the emission spectra. The origin of the spectrophotometric shift of the peaks, as well as the difference in migration patterns, cannot be attributed to a difference in the extraction procedure or in the medium containing the two porphyrins, since both extractions were similar and the electrophoretic medium consisting of a stationary phase (polyacrylamide) and a mobile phase (aqueous solution containing Tris, tricine and SDS) was identical for both extracts. Further mass spectrometric and NMR characterizations are needed to understand the differences in

spectrophotometric shifts between the two uroporphyrins, which may be attributed to the low biochrome content in *T. crenularis* compared to that in *P. lineatus*.

## 3.4 | Analysis of biochrome-macromolecule complexes

An additional advantage of coupling electrophoresis and PL spectral imaging for the visualization of extracts containing biochromes is the ability to infer the molecular interactions of biochromes with other organic macromolecules of the skeletal matrix. This information is deduced from the migration behaviour of biochromes. In the present example, extracts of the living *P. lineatus* and of the fossil *T. crenularis* are characterized by brown smears visible under natural light (Figure 2a, lanes 2, 3 and 5). The molecular distribution of these pigmented smears extends to relatively high molecular weights (far above 10 kDa), clearly suggesting that the biochrome is associated with polydisperse macromolecules (proteins or polysaccharides) of the shell matrix. It is indeed known for a long time that skeletal matrices of calcified tissues exhibit such an electrophoretic behaviour (Marin & Luquet, 2007). In addition, as the electrophoresis was performed in denaturing conditions (SDS+β-mercaptoethanol), this may indicate that the dark brown biochrome and the polydisperse macromolecules are strongly bound together, suggesting the involvement of covalent bonds. On the contrary, the thick discrete porphyrin signal observed in red in the false colour PL image in lanes 3 and 4 of Figure 2f (*P. lineatus*) is located at the bottom of the gel in a zone corresponding to components of very low molecular weight. Its position suggests two possibilities: (a) it is not bound to any matrix macromolecules, the most likely case; (b) it was bound, but the bond was labile and easily disrupted by the denaturing conditions of the electrophoretic migration.

## 3.5 | Upscaled purification of porphyrins

In the present example, the peculiar migration pattern of the porphyrin biochrome of *P. lineatus* (i.e. discrete band migrating far ahead of the migration front of the gel) can be exploited to upscale its purification by preparative electrophoresis, as shown by Figure 3. This aspect is particularly useful if the raw shell material is abundant. This technique, primarily used for purifying proteins from bulk shell extracts (Marin et al., 2001; Marxen et al., 2003), has been specifically adapted here to biochromes, which can be eluted from the gel and recovered with a fraction collector (Figure 3a). Similarly to Step 2, the fractionation is performed on a non-standard 15 or 16% acrylamide Tris-tricine preparative gel. The technique allows the purification of large amounts of biochromes, eluted early in the different tubes (tubes 4–8). The PL spectral imaging system allows porphyrin visualization directly during the gel run and in the fractions collected in the tubes (Figure 3b). Note that the elution of the porphyrin in the tubes follows an asymmetric normal law. As for gel
<figure><img src="image_6.png" /><figcaption>**FIGURE 3** Results of the preparative electrophoresis (Tris-tricine gel, 15% acrylamide) of the ASM >10kDa of *Phorcus lineatus*, visualizing the eluted fractions in tubes. Electrophoresis fractions were collected 'blind' as in Marin et al. (2001). In situ luminescent spectral imaging was applied to detect the tubes containing the biochrome. (a) Twelve first collected tubes shown in visible light. Note that tubes 10–12 are darker due to the elution of the migration front containing bromophenol blue. (b) Same image as (a), but obtained with a false colour photoluminescence (PL) of the tubes using a 385 nm excitation and generated by associating three black and white images (red channel: 708 ± 37 nm; green channel: 571 ± 36 nm; blue channel: 472 ± 15 nm); interestingly, most of the porphyrin biochrome is eluted in tubes 4 and 5, before the migration front. (c) PL spectra directly acquired on tubes 4–9, between 580 and 800 nm. We used a 385 nm excitation wavelength and a portable spectrophotometer equipped with a high-pass inferential filter to cut-off the contribution of the UV excitation. Note the attenuation of the signal from tubes 4 to 9, consistent with that observed in (b).</figcaption></figure>

bands of Step 2, PL spectra are obtained directly from each tube, and the corresponding signals are integrated for relative quantification. The porphyrin elution before the migration front of the electrophoresis guarantees a high purity of the biochrome, since it is not contaminated by macromolecules of the matrix, which elute after the migration front. Consequently, the different fractions obtained can be pooled together and extensively dialyzed in dialysis bags of very low cut-off (500 Da) to remove molecules of the eluent (Tris, glycine). The pure biochrome is then ready to be freeze-dried and analysed further.

# 4 | DISCUSSION

## 4.1 | Porphyrins in shells: comparison with previous studies

In the present paper, we have set up a novel and original approach to identify and characterize porphyrins in gastropod shells by coupling electrophoresis to PL spectroscopy and imaging. Our spectrophotometric findings are congruent with the literature

describing the presence of porphyrins in gastropod shells: this includes the precursor works of Comfort and coworkers (Comfort, 1949, 1951; Nicholas & Comfort, 1949) and more recent studies (Williams, 2017; Williams et al., 2016). In more detail, our spectra—acquired on two vetigastropod species (of the Trochida order)—are in complete agreement with that obtained by Williams and coworkers on the marine gastropods *Clanculus pharaonius* and *Clanculus margaritarius*, two vetigastropod representatives of the Trochidae family, like *P. lineatus*. Using a different set of techniques, these authors identified the presence of uroporphyrins I and III (Williams et al., 2016), the two most commonly found uroporphyrins in Nature. It is puzzling to observe that in the shell of three other gastropods belonging to another subclass (Caenogastropoda, order Neogastropoda), protoporphyrin IX and not uroporphyrin, was identified (Verdes et al., 2015). In the future, it will be interesting to check whether this difference, observed so far in few specimens of two clades (Vetigastropoda vs. Caenogastropoda), applies to all members of these two major and well-defined gastropod subclasses. If so, as uroporphyrins and protoporphyrins are end products of different biochemical synthesis pathways (Kadish & Guilard, 2024), this suggests a
fundamental evolutionary change between these two clades in the gene network involved in porphyrin synthesis and secretion. As illustrated here, the coupling of electrophoresis and PL imaging and spectroscopy—besides advancing the chemical characterization of porphyrins directly on gel, even with an attenuated signal due to fossilization—will contribute to comprehend the macroevolution of shell pigments in molluscs (Williams et al., 2017).

## 4.2 | Unconventional features of our protocol

In calcified tissues, biochromes are poorly studied in relation to biomineralization processes and to their association with the skeletal matrix. When pigmented skeletal matrices are extracted for subsequent analysis, biochromes are often lost during key steps (ultrafiltration, dialysis) of the extraction procedure (Immel et al., 2016; Pavat et al., 2012). When bound to macromolecular components, some biochromes go unnoticed simply because they are neither detected nor the central focus of the study. By allowing the sequential visualization of biochromes and of skeletal macromolecules in gels, our protocol provides a novel and exhaustive picture of organic molecules contained in a calcified tissue. The novelty of our approach lies in two aspects: firstly, a non-standard use of existing techniques and secondly, three methodological optimizations performed at different steps of the analysis.

The first aspect is the use of a commercial gel electrophoresis system (analytical or preparative). To our knowledge, analytical gels have been rarely used in simple in vitro systems with porphyrins (Kowalska et al., 2003) but never for the fractionation of free biochromes in natural extracts containing a complex mixture of macromolecules. In addition, for the first time, preparative gel electrophoresis was used for the fractionation of luminescent biochromes from natural samples. Another non-standard application is the improved detection of luminophores using a multispectral imaging allowing optimizing the selection of the spectral bands of detection in accordance with the emission properties of the different chemical moieties. The adjustment of a commercial spectrofluorometer to perform remote excitation emission measurements outside of the original sample compartment dedicated to the study of small samples or solutions in a cuvette is also a significant improvement to apply the techniques to a wider range of materials (see Section 2). A special accessory is composed of two mirrors and a bifurcated fibre of which output was inserted into a custom-made dark compartment to avoid the collection of stray light. This configuration allows collecting excitation/emission spectra on areas of 2 mm diameter; this size is compatible with the dimensions of electrophoresis bands— even weak ones—on a mini-gel. Finally, the most innovative aspect of the non-standard protocol consists in coupling electrophoresis to this optimized imaging system. Our methodology is fully complementary to the existing analytical procedures used for characterizing porphyrins in calcified tissues, as shown in Table S1.

Three key methodological optimizations were implemented. Firstly, the use of a low cut-off (1 kDa) for the ultrafiltration system

to recover most of the organic components of the skeletal tissue, especially those of low molecular weight, as already used for the extraction of sea urchin naphthoquinone pigments (Albéric et al., 2023). Secondly, the use of Tris-tricine gels of high acrylamide content (15, 16%), to obtain a clear separation of unbound biochromes and very low molecular weight constituents from skeletal matrix macromolecules (Albéric et al., 2023). Thirdly, the PL imaging was performed immediately after gel migration and demoulding but before any type of gel staining. This aspect is a key point in our analytical procedure, as all the protocols found in the literature describe the first staining step immediately after gel has been removed from the casting plates. We therefore recommend that electrophoretic gels, whatever they are, should always be checked with a UV flashlight, to reveal invisible and unstainable molecules that are potentially present (as discrete bands or smears) in the fractionated extracts.

## 4.3 | Advantages

The combination of mild extraction, non-standard gel electrophoresis and spectral imaging represents a breakthrough on the way to accessing and exploiting the information contained in electrophoretic gels. The multiple advantages offered by our protocols are twofold: the typologies of information that can be retrieved and the compatibility of the protocol with standard analytical procedures.

It is therefore possible to visualize biochromes and calcifying matrix macromolecules on the same gel: the molecular association (either covalent binding or non-covalent binding) between biochromes and matrix macromolecules can be accurately depicted. The example of *Phorcus* shows that the main luminescent but invisible biochromes are free (i.e. not bound to macromolecules), while the dark ones, visible under natural light, are likely to be associated with molecules of higher molecular weight.

Moreover, due to the combination of the mild extraction and the absence of staining, spectroscopic measurements directly on the gel are possible and give access to the spectral signature of porphyrins in a very simple way, with a fast acquisition time. Other biochromes, such as carotenoids, can also be analysed in the same manner. Remarkably, when different biochromes are mixed in an extract, the electrophoretic fractionation allows the deconvolution of this complex signal into several discrete bands, which can be studied separately with PL spectral imaging. The use of PL as a probe allows revealing extremely low amounts of biochromes that are not detectable under natural light: the spectral image, by allowing extremely sensitive detection, is therefore particularly suited to the study of traces of biochromes, such as those found in fossils.

As far as the analytical procedure is concerned, the protocol does not involve any major changes and is fully compatible with subsequent standard gel stains such as CBB, silver nitrate, alcian blue, carbocyanine; in other words, the dyes mostly used in biomineralization research (Immel et al., 2016; Marin & Luquet, 2007). The spectral analysis must be performed prior to staining but does not alter the staining procedure in any way. Alternately, the unstained gels can
be partly dried, hermetically sealed between two plastic foils and stored flat in a cold and dark environment, if necessary. This allows preserving biochromes in the gels. Luminescent signal can consequently be retrieved months or years later. If required, the upscaling of the experiment to preparative fractionation is implemented to obtain large amounts of purified biochromes or protein-biochrome complexes. The application of the approach can be extended to all kinds of composite materials made of organic macromolecules and biochromes. The protocol applies also to non-mineralized systems. Plants or archaeological stained leather, or any sample containing luminescent or non-luminescent biochromes can be analysed in this manner. Most of the chemicals used are environmentally friendly and, apart from acrylamide (which, when polymerized, is harmless), none of the chemicals used for the analysis on gel are harmful. At last, the whole protocol as described here from the extraction to the spectral analysis is not costly, and implementing the mini-gel electrophoretic system in a lab is easier and less demanding and expensive than setting up other types of fractionation techniques (HPLC or spectrometric techniques).

## 4.4 | Limitations

Beside the fact that our protocol destroys the sample (as listed in the Table S1), there are few other limitations, mostly related to the duration of the complete analytical procedure and the range of biochromes that can be studied.

The whole experiment, including preparative purification of biochromes, is time-consuming: the matrix extraction step, from decalcification to freeze-drying of the extracts, lasts 1 week or more; a particular bottleneck is the ultrafiltration on a 1 kDa cut-off membrane, which is tedious but mandatory if one wants to collect most of the organics from skeletal tissues; the preparative electrophoresis (step 3) takes 2 days. In addition, when step 3 is performed, the collected tubes with biochromes also contain Tris and glycine (elution buffer), which must be removed by dialysis in very low cut-off tubes (<500 Da). The dialysis process takes a few more days. In addition, the protocol described here, from the extraction to the analysis on gel, is adapted to water or acid-soluble biochromes. Biochromes that are soluble only in organic solvents, for example ether or chloroform, cannot be treated with our method.

## 4.5 | Further developments: other objects, methodologies and applications

Our protocol is aimed firstly at the lively scientific community of biomineralization that studies calcified tissues in general and not only mollusc shells: eggshells in vertebrates, exoskeletons in corals, bryozoans and annelids, spines and plates in echinoderms, carapaces in crustaceans, but also stromatolites and microbialites made by cyanobacteria. These two latter biomineralized structures of microbial origin contain also biochromes (Lepot et al., 2014). Mineral-associated

biochromes were primarily considered from an evolutionary viewpoint, as adaptive responses to the environment (i.e. 'camouflage') or as part of the arsenal to attract sexual partners in the case of the gastropod shell (Saenko & Schilthuizen, 2021). Today, we are witnessing a resurgence of interest in their study. Some biomineral-associated biochromes are thought to be part of the immune response (via the tyrosinase enzymatic pathway) prior to their occlusion in the mineral phase (Liu et al., 2021; Trinkler et al., 2011). They are also suspected to be involved in the calcification process itself and their function in biomineralization needs to be urgently revisited (Parvizi et al., 2023; Zito et al., 2015). From applied perspectives, their usage as natural substances should be further explored: anti-microbial/anti-tumoral agents or eco-friendly dyes are, among others, compelling applications (Albéric et al., 2023; Hou et al., 2016, 2019; Roncoroni et al., 2024).

Our protocol opens up new avenues to the development of other methodological investigations. When a biochrome of interest is suspected to be associated with a protein, its visualization on an unstained gel allows a direct manual sampling of the band/spot, its subsequent in-gel enzymatic digestion and the analysis of the resulting peptides by proteomics. This provides a formal identification of the protein-biochrome complex. More broadly, the methodological framework presented here can be extended by coupling electrophoresis with other types of imaging techniques: Raman spectroscopy is a promising perspective that can provide additional information on the chemistry of the biochromes of interest.

Finally, other applications arise from our innovative protocol: since biochromes can be fractionated on gel and chemically characterized by PL spectral imaging even at trace concentration, our approach is particularly suitable for the characterization of organic (macro)molecules in ancient materials and should interest conservation science, archaeology, palaeontology, ecology and evolution. Many remarkable fossil deposits contain shells that have retained their biochromes for millions of years (Caze et al., 2015; Tahoun et al., 2021). In particular, porphyrins can be detected in samples as old as the Jurassic (Caze et al., 2011; Wolkenstein et al., 2024). Their exceptional diagenetic preservation raises questions on the multiple causes of this stability: molecular structure, affinity with the mineral phase, tight or loose association with the organic macromolecules. This last point is particularly critical as the presence of traces of native biochromes in fossil shells may be a proxy for the preservation of the associated matrix macromolecules: if so, a quick check with UV light may be a prerequisite step to the selection of suitable precious fossils before any shell matrix extraction.

Our protocol makes it possible to study the relationship between the preservation of biochromes and that of macromolecules in the matrix. It will undoubtedly advance the understanding of molecular diagenesis. In addition, because coupling electrophoresis and luminescent imaging can deconvolute complex biochrome signals, this will allow the reconstruction of past mollusc communities by identifying, similarly to a barcoding, peculiar biochrome complexes contained in fossil shells, which may be related to sexual dimorphism, populations, species, geographical localities, specific environments
<figure><img src="image_7.png" /></figure>

or ecological interactions between individuals. Molecular palaeo-
ontology, ecology and evolution will benefit greatly from such ad-
vances, which will open up new opportunities to trace the origin and
evolution of biochrome complexes in the geological record and to
revisit entangled ecological interactions from the past.

AUTHOR CONTRIBUTIONS

The study was designed by Mathieu Thoury and Frédéric Marin
and performed by Luca Polacchi, Mathieu Thoury and Frédéric
Marin, with the assistance of Camille Lutet-Toti, Hélène Pasco,
Jérôme Thomas and Marie Albéric. The initial draft was sketched
by Luca Polacchi, and the successive draft versions were writ-
ten by Frédéric Marin and Mathieu Thoury with the assistance of
Pierre Gueriau. The three figures and Figure S1 were designed by
Mathieu Thoury with inputs from Luca Polacchi, Frédéric Marin
and Didier Merle. The Table S1 was designed by Frédéric Marin.
The samples were provided by Olivier Basuyaux and Didier Merle.
The expertise on porphyrins was provided by Mathieu Thoury,
Marie Albéric, Benoît Habermeyer and Roger Guilard. Data were
deposited on the Dryad platform by Mathieu Thoury. Financial
support was provided by Mathieu Thoury and Frédéric Marin. All
authors contributed critically to the drafts and gave final approval
for publication.

ACKNOWLEDGEMENTS

The postdoctoral work of Luca Polacchi was performed within the
PRESERV project (Photoluminescence of porphyrins to REveal pre-
Served colour markER in post-paleozoic Vetigastropoda), which
was supported by the Paris Ile-de-France Region–DIM PAMIR
(PAtrimoines Matériels-Innovation, expérimentation et Résilience).
We would like to thank Rémi Métivier of the PPSM laboratory
for providing access to the Fluorolog spectofluorometer. The
work of Frédéric Marin and Camille Lutet-Toti was supported
by the MAELSTROM project from the TELLUS-INTERRVIE pro-
gram (CNRS, 2022) and the PRELUDE project (2023) financed by
Observatoire des Sciences de l'Univers (OSU) Theta, Besançon.
Camille Lutet-Toti is the recipient of a PhD fellowship under joint
supervision between the University Burgundy Europe (UBE)
and Università di Bologna Alma Mater Studiorum (UniBo), in the
frame of the International PhD Program 'Innovative Technologies
and Sustainable Use of Mediterranean Sea Fishery and Biological
Resources' (www.FishMed-PhD.org). This study represents partial
fulfilment of the requirements for the Ph.D. thesis of Camille Lutet-
Toti in the FishMed-PhD Program.

CONFLICT OF INTEREST STATEMENT

All authors of this paper declare that they have no competing
interests.

PEER REVIEW

The peer review history for this article is available at https://www.
webofscience.com/api/gateway/wos/peer-review/10.1111/2041-
210X.70176.

DATA AVAILABILITY STATEMENT

Data available via Dryad Digital Repository https://doi.org/10.5061/
dryad.rr4xgxdjc (Thoury et al., 2025).

ORCID

Frédéric Marin https://orcid.org/0000-0001-8319-1735

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SUPPORTING INFORMATION

Additional supporting information can be found online in the
Supporting Information section at the end of this article.

**Figure S1.** Multispectral imaging system.

**Table S1.** Comparison of the most current approaches used to identify and analyse porphyrins in calcified tissues.

**How to cite this article:** Polacchi, L., Lutet-Toti, C., Basuyaux, O., Gueriau, P., Albéric, M., Pasco, H., Thomas, J., Merle, D., Habermeyer, B., Guilard, R., Thoury, M., & Marin, F. (2025). Coupling gel electrophoresis with photoluminescence reveals biochrome complexes in modern and fossil shells. *Methods in Ecology and Evolution, 00*, 1-12. https://doi.org/10.1111/2041-210X.70176