Assessment of the Impact of Cadmium on the Redox Status and Metabolic Profile of the Sea Urchin Paracentrotus lividus

Summary

This protocol highlights the methods of extraction and characterization of bioactive compounds from the sea urchin Paracentrotus lividus (Echinoderm) as well as determination of oxidative stress parameters after cadmium exposure.

Abstract

Cadmium (Cd) is one of the most harmful heavy metals to aquatic organisms and a widespread pollutant in marine biota. Because of its high bioavailability, persistence, and ability to bioaccumulate, cadmium (Cd), a toxic heavy metal discharged by mining runoff, industrial discharge, and agricultural inputs, is one of the most detrimental pollutants to aquatic organisms and a common pollutant in marine biota. The current study assessed the potentially toxic effects of cadmium on oxidative stress biomarkers and metabolite profiles in Paracentrotuslividus gonads exposed to a range of concentrations (0.0183, 0.183, and 1.83 mg/L) over 96 h. The results showed that Cd induced oxidative stress by increasing malondialdehyde, hydrogen peroxide, and lipid hydroperoxide levels compared to the control group, as well as increasing antioxidant activities (reduced glutathione and catalase). In addition, based on the gas chromatography-mass spectrometry analysis, 74 metabolites were identified in the gonadal extracts of both treated sea urchins and the control. The Partial Least Squares Discriminant Analysis revealed a clear class separation and discriminated nine relevant metabolites through the VIP scores. The performed heatmap analysis highlighted disturbance in the levels of sterols and fatty acid compounds such as Cholest-5-en-3-ol(3.beta.)-, Desmosterol, and (23S)-ethylcholest-5-en-3.beta.-ol and eicosapentaenoic acid (EPA), as well as the antioxidant defenses such as retinoic acid, and steroid regulation such as Pregna-5-en-20-one, 3-hydroxy-,(3β). Overall, our data revealed the high sensitivity of gonads to Cd, even at relatively low concentrations.

Introduction

The marine environment can also be considered an important sink for several toxic chemicals, such as heavy metals. Due to their lipophilicity, these elements have a tendency to bind with suspended particulate matter and progressively build up in marine sediments, where they show decreased mobility and bioavailability. As a result, marine sediments serve as a major heavy metal sink¹. Among these contaminants, Cadmium (Cd) has long been acknowledged as a major global water pollutant that threatens ecosystems and their related biota². One-third of the Cd present in aquatic ecosystems comes from the manufacturing and use of phosphate fertilizers². Other sources of Cd release into the environment include the mining sector, electroplating companies, and natural processes of the Earth's crust³. This pollutant is highly widespread owing to its ability to be easily transported through aqueous media, and its comparatively great mobility in contrast to other heavy metals². According to the literature, concentrations of Cd vary between 12 and 16 x 10⁻³ µg/L in the Baltic Sea⁴, and between 13.4 x 10⁻³ and 1.49 µg/L in a very contaminated site along the Galician coasts (NW Iberian Peninsula)⁵. Much more elevated levels of Cd were recorded in some polluted areas in the Mediterranean Sea, where concentrations may reach 80.4 x 10^-3 µg/L near Sicily Island (Italy)⁶, and 48 µg/L in the Gulf of Gabès (Tunisia)⁷. This persistent, bio-accumulative, and nonessential transition metal is recognized for its ability to cause serious health problems for both human and aquatic organisms⁸. Cd can directly exert its toxicity by binding to proteins, inhibiting enzyme activities, and interfering with calcium homeostasis⁹. It can also act indirectly through the generation of reactive oxygen species (ROS), leading to oxidative damage to biomolecules such as lipids, proteins, and DNA¹⁰. Several scientific studies have reported that Cd contamination may trigger structural and functional disorders in aquatic organisms¹¹.

As with the other marine benthic invertebrates, sea urchins are particularly exposed to marine pollution. Despite their capacity to accumulate high levels of heavy metals, sea urchins can cope with metal contamination and endure in polluted environments¹². Thus, they are commonly recognized as sentinel organisms and are widely used as models for ecotoxicological and environmental studies¹³. Numerous studies have investigated the impact of Cd exposure on several sea urchin species at different life stages, including adult organisms, embryos, and larvae¹³. It was found that Cd exposure may result in a variety of adverse effects ranging from reduction of egg fertilization rate, abnormalities in embryo and larva development, oxidative stress, to apoptosis and autophagy¹³. Most of the available literature focuses on the effect of Cd on sea urchin developing embryos^13,14; however, only a few studies¹⁵have been performed to understand the effects of this metal on the metabolic profile of adult sea urchins, and no information is available on lipid metabolism.

Continuous exposure of Paracentrotus lividus (Lamarck, 1816) embryos to subacute/sublethal cadmium concentrations causes abnormal development with defects in skeleton elongation and patterning, coupled with a general reduction of protein synthesis, and differential metallothionein expression¹⁶.

The proposed method improves the assessment of cadmium (Cd) toxicity and oxidative stress in marine organisms by combining biochemical assays with targeted and untargeted metabolomic profiling using gas chromatography-mass spectrometry (GC-MS). Compared to traditional approaches that rely solely on single biomarkers such as lipid peroxidation or antioxidant enzyme activities (e.g., SOD, CAT), the addition of GC-MS-based metabolite screening provides a comprehensive biochemical fingerprint of the organism's physiological state¹⁷. This integration enables more sensitive and systemic detection of sublethal effects, even at low Cd concentrations, and can detect early perturbations in energy metabolism, amino acid turnover, and oxidative pathways that occur before observable toxicological damage. Importantly, this method complements existing toxicological assessments by providing higher resolution data and allowing for the identification of novel biomarkers that traditional assays may miss¹⁸. It also promotes mechanistic understanding through pathway-level analysis, which enhances predictive ecotoxicology models¹⁹. This method detects cadmium exposure in the 1-100 µg/L range, which is relevant for polluted coastal and estuarine systems²⁰. The approach is most appropriate for controlled laboratory exposures or moderately complex field studies. However, interpretation can be difficult in highly contaminated matrices where multiple stressors (e.g., heavy metals, PAHs, emerging contaminants) interact and potentially confound biomarker specificity. Despite this, when properly designed, GC-MS's robustness and specificity, combined with multivariate data analysis (e.g., PCA, PLS-DA), provide strong discriminatory power to resolve overlapping effects.

This integrated biochemical-metabolomic platform is particularly useful for sentinel species monitoring and ecological risk assessments under frameworks such as the Marine Strategy Framework Directive (MSFD) and is consistent with current efforts to advance omics-based environmental monitoring²¹.

In the current study, we used the purple sea urchin P. lividus, which is one of the most exploited and economically significant species throughout the Atlantic and Mediterranean coast²², to assess the effect of Cd contamination on the redox status and the metabolomic profile of the gonads.

Protocol

1. Samples and experimental design

1. Collect a total of 40 adult sea urchins P. lividus specimens (20 males and 20 females, mean length: 45 ± 4 mm, mean age: 2.0 ± 0.5 years, mean weight: 22.54 ± 2.50 g) by divers at the depth range of 5-7 m in May 2023 from Bizerte coasts (Mediterranean Sea, Tunisia).
2. Transfer the collected sea urchins P. lividus of mixed sex, estimated age approximately 12-18 months based on size to the lab and keep in aerated natural seawater 16 ± 1 °C with natural salinity of ~38 PSU, pH of 7.8-8.0 and photoperiod of 12:12. Monitor the parameters daily throughout the exposure period to ensure environmental stability and minimize external variation.
3. Inject 1 mL of KCl 0.5 M into the peristomial region to induce spawning and visually identify gametes. Rinse individuals with filtered seawater and acclimate them for 3 days in controlled conditions with no food provision.
4. Randomly divide Sea urchins into four groups of nine individuals (three individuals per tank in triplicate) and allocate them into 50 L glass aquaria. Determine the sample size based on previously published studies^23,24 that evaluated oxidative stress biomarkers and metabolomic responses in marine invertebrates and use comparable or fewer biological replicates.
   NOTE: This level of replication ensures strong biological representation and statistically significant results.
5. Expose after acclimation, animals for 96 h to a range of cadmium chloride (CdCl₂) concentrations as follows: Group I (Ctrl): sea urchins kept in non-CdCl₂water; Group II (Tr1): sea urchins exposed to a CdCl₂concentration of 0.0183 mg/L; Group III (Tr2): sea urchins exposed to a CdCl₂concentration of 0.183 mg/L; Group IV (Tr3): sea urchins exposed to a CdCl₂concentration of 1.83 mg/L.
6. To maintain the stability of water during the experiment, change 50% of water every 24 h, remove the water carefully using a siphon, and replace it with freshly prepared water while maintaining the concentrations of CdCl₂.

2. Tissue preparation

1. Perform, using the protocol described by Telahigue et al.²³, the preparation of the sea urchin tissue homogenates.
2. Dissect on ice sea urchins from each group at the end of the experiment. Use sterile scissors to make a circular cut around the mouth to open the test.
3. Remove carefully the five gonads with a spatula and keep for further analysis. Wash with cold distilled water to remove debris and surface contaminants
4. Homogenize using a probe sonicator set to 40% amplitude, in 3 bursts of 10 s each with 20 s rest between bursts, 300 mg of gonad fractions with Tris-HCl buffer (20 mM, pH 7.4) under cold conditions.
5. Divide each gonad sample into two portions: one for biochemical analysis (fresh) and one for metabolomic analysis (lyophilized).
6. Centrifuge at 10,000 x g for 20 min. Store the supernatants at - 80 °C for biochemical parameter analysis.
7. Lyophilize other gonad fractions using a freeze dryer prior to GC-MS analysis. Maintain for 48 h at -50 °C with a vacuum of less than 0.1 mbar until total drying is accomplished.
8. Utilize the Lowry method to determine the total protein content of each sample using BSA as the standard. Determine LOOH level at 560 nm and express as mmol/mg of protein.
   NOTE: This quantification was performed prior to all biochemical assays to normalize biomarker levels.

3. Reduced glutathione assay

1. Estimate reduced glutathione (GSH) content according to the method of Ellman²⁵, based on the absorbance of 2-nitro-5-mercapturic acid resulting from the reduction of 5-50-dithio-bis-2-nitrobenzoic acid (DTNB) by the thiol (SH) group of the glutathione.
2. Centrifuge at 1600 x g for 15 min a mixture of tissue homogenates in phosphate buffer and sulfosalicylic acid (4%).
3. Take a volume of 500 µL of the obtained supernatant and add it to Ellman's reagent (DNTB) with 0.025 mL. Read at 412 nm the absorbance of the resulting mixture.
4. Asses GSH content as mg per g of tissue. Express total GSH content as µg/ mg of tissue. Calculate GSH levels using a standard calibration curve based on known concentrations of reduced glutathione.
   NOTE: The objective is to quantify reduced glutathione (GSH) in gonadal homogenates as an oxidative stress marker following cadmium exposure.

4. Catalase activity

1. Carry out reaction setup according to the method of Aebi²⁶. Prepare the reaction medium consisting of 2.9 mL of 100 mM phosphate buffer (pH 7.4) and 20 µL of gonadal tissue supernatant. Pre-equilibrate the reaction medium to room temperature.
2. Start the reaction by adding 80 µL of freshly prepared 0.5 M hydrogen peroxide (H₂O₂) solution to the cuvette. Immediately mix the contents thoroughly and record the decrease in absorbance at 240 nm for 1 mi at 10 s intervals using a UV-Vis spectrophotometer.
3. Calculate CAT activity based on the rate of H₂O₂ decomposition, determined by monitoring the decrease in absorbance at 240 nm over time. Use the following formula: CAT activity= (ΔA/min x V) / (ε x l x C)
   where: ΔA/min = change in absorbance per minute at 240 nm, V = total volume of the assay (in L), ε = molar extinction coefficient of H₂O₂(43.6 M^-1/cm), l = path length of the cuvette (usually 1 cm), C = protein concentration in mg/mL.
4. Use the molar extinction coefficient of 43.6 M/cm. Express CAT activity as nmol H₂O₂ decomposed per minute per mg protein (nmol H₂O₂/min/mg protein).
   NOTE: The objective is to quantify catalase activity in gonadal homogenates of P. lividus as a measure of antioxidant enzyme response to cadmium exposure.

5. Measurement of hydrogen peroxide generation

1. Measure the hydrogen peroxide (H₂O₂) levels in P. lividus gonads using ferrous oxidation-xylenol orange assay according to the method described by Ou and Wolff²⁷.
2. Determine the concentration of H₂O₂in the supernatant using a spectrophotometer at 560 nm. Express the results as nmol/mg of protein.

6. Malondialdehyde assay

1. Prepare samples as described in the Draper and Hadley method²⁸. Mix 500 µL of gonad homogenate with 1 mL of 10% trichloroacetic acid (TCA). Centrifuge at 2500 x g for 10 min at 4 °C.
2. Transfer 1 mL of the resulting supernatant into a new tube and add 1 mL of 0.67% TBA solution. Incubate the mixture in a water bath at 100 °C for 15 min.
3. Cool the samples to room temperature. Measure absorbance at 532 nm using a spectrophotometer.
4. Subtract background absorbance using a reagent blank (TBA + TCA without tissue extract). Prepare a standard curve using 1,1,3,3-tetraethoxypropane (TEP) and express MDA concentration as nmol/mg protein.

7. Lipid hydroperoxide measurement

1. Measure the lipid hydroperoxide (LOOH) in gonad tissues using the ferrous oxidation in xylenol orange assay, as described by the method of Jiang et al.²⁹.
   NOTE: The objective is to assess lipid peroxidation levels in gonadal tissues by quantifying malondialdehyde (MDA), a byproduct of oxidative membrane damage.

8. Extraction of bioactive compounds from gonads and analytical screening

1. Perform methanolic extraction of lipophilic compounds in triplicate from the dried gonadal tissue of each group in triplicate by soaking the material in analytical grade methanol (≥99.8%; 1:25 w/v) on a rotary shaker at 150 rev/min at ambient temperature.
2. Centrifuge all extracts for 15 min at 700 x g, 4 °C, after evaporation of MeOH fractions by rotary evaporation at a water bath temperature of 40 °C and a rotation speed set to 100 rpm. Resuspend the crude extract in 1 mL of MeOH, then store in sealed vials at 4 °C.

9. Gas chromatography-mass spectrometry analysis of gonadal extracts

1. Make a 500 µL aliquot in a vial using the methanolic extract that has been stored.
2. Perform analysis by Gas chromatography-mass spectrometry (GC-MS) on a gas chromatograph coupled to a mass spectrometer. The GC column was 5 %-Phenyl-methylsiloxane: 30 m x 0.25 mm x 0.25 µm film thickness.
3. Use helium as carrier gas at a constant flow rate of 1 mL/min. Set the injection volume to 1 µL in the split mode at a ratio of 100:1. Program the oven temperature from 4-280 °C at 5 °C/min.
4. Operate the spectrometer in electron-impact (EI) mode, ionization energy was 72 eV, ion source temperature was 250 °C, scan time 1 s, and mass range 45-550 amu.
5. Perform identification of compounds by comparison of their mass spectra with those from the NIST98MS Library and Willy Library275 Database.

10. Statistical analysis

1. Evaluate the dataset first for normality using the Shapiro-Wilk test and for homogeneity of variances using Levene's test.
2. Preprocess the data with MetaboAnalyst 6.0³⁰, using standard normalization procedures, log transformation, and Pareto scaling to meet ANOVA assumptions such as normal distribution and variance homogeneity.
3. Perform a multivariate statistical analysis using Metaboanalyst, use the Benjamini-Hochberg false discovery rate (FDR) method for multiple testing correction to account for Type I errors related to numerous biochemical comparisons, and guarantee statistical robustness.
4. Use the one-way analysis of variance (ANOVA) to compare results between groups. Differences between means were considered significant when p <0.05.
5. Use Partial Least Squares Discriminant Analysis (PLS-DA) to discriminate between comparable groups. Determine the variable importance in the projection (VIP) for all compounds.
6. Calculate p-values using the paired Student's t-test for single-dimensional statistical analysis of standardized variables.

Results

The validity and efficacy of our methods were ensured by the use of biological and technical replicates, which enabled consistent and reliable measurements. The assays demonstrated low variability, with standard deviations typically less than 10%, indicating high repeatability. To ensure accurate metabolite quantification, GC-MS analyses included calibration curves and internal standards. All techniques used are based on widely accepted protocols that have previously been validated in mar...

Discussion

This study was planned to evaluate the toxic effect of Cd on the sea urchin P. lividus gonads by analyzing biochemical biomarkers and active metabolites. The exposure of P. lividus to different concentrations of Cd produced an oxidative stress response. These findings are similar to previous reports indicating close correlations between the increased antioxidant levels in the sea urchin grounds and contaminant levels¹³.

Our results revealed a significa...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Freeze dryer	Biobase	BK-FD12PT freeze dryer	Lyophilization
Gas chromatograph	Agilent Technologies	HP 6890 gas chromatograph	Flame Ionization Detector (FID)
GC column	Agilent Technologies	HP-5 (5%-Phenyl)-methylsiloxane column	Nonpolar column that delivers superb performance in a wide range of applications
Mass spectrometer	Agilent Technologies	HP 5973 mass spectrometer	Quadrupole MS