Immobilized Sinirhodobacter sp. 1C5-22 for Multi-Metal Bioremediation: Molecular Resistance Mechanisms and Operational Validation in Industrial Wastewater Systems

Abstract

A novel heavy metal-resistant bacterium with significant bioremediation capabilities, Sinirhodobacter sp. 1C5-22 was isolated from moderately polluted Shenzhen Futian mangrove rhizosphere sediments. This strain showed exceptional tolerance (MIC ≥ 600 mg/L for Cu/Zn; > 500 mg/L for Ni). Analyses revealed distinct metal-specific distribution strategies: Cd and Ni were predominantly bound extracellularly (>80%); Cu was bound intracellularly (~60%); and Zn exhibited balanced partitioning. Integrated omics analysis identified a molecular defense mechanism coordinated by the CreB transcriptional regulator. This Adsorption–Sequestration–Efflux (ASE) system integrates extracellular polymer binding, periplasmic sequestration via stable metal-binding proteins, and efflux pump activity, resolving the apparent adsorption-tolerance paradox at elevated concentrations. For bioremediation applications, we developed a polyvinyl alcohol–sodium alginate immobilized consortium (PVA-SA 1C5-22). The engineered agent displayed significantly enhanced biosorption capacity compared to free cells and effectively mitigated heavy metal-induced oxidative damage, evidenced by stabilized malondialdehyde levels. It demonstrated robust reusability, maintaining high metal enrichment across five adsorption–desorption cycles in multi-metal wastewater with efficient HCl-driven desorption (55–70%). Critically, it achieved stable nickel removal performance (~20% adsorption, >50% desorption) from authentic electroplating wastewater (1850 mg/L Ni²⁺) through successive multiple cycles. Our integrated approach bridges microbial ecology and environmental biotechnology, establishing this immobilized system as a highly sustainable strategy for complex industrial effluent remediation.

1. Introduction

Mangrove ecosystems harbor unique microbial communities exhibiting high tolerance to environmental stressors, underpinning their significant potential for bioremediation applications [1,2]. Particularly in the rhizosphere, microbes have evolved sophisticated strategies, such as extracellular adsorption, intracellular sequestration, and efflux systems, enabling survival and activity in metal-contaminated sediments [3,4]. Crucially, this adaptive potential is intricately linked to pollution regimes. Environments subjected to moderate chronic pollution, as exemplified by Shenzhen’s Futian Mangrove Reserve [5], uniquely balance selective pressure through two key attributes: substantial microbial biodiversity maintenance [6], as validated across diverse mangrove ecosystems [7,8], and enhanced adaptive evolution and functional specialization [7,8]. This synergistic effect, also documented in estuarine sediments globally, contrasts sharply with severely contaminated industrial sites, where microbial communities become depleted in diversity and functional capacity [9,10]. Thus, moderately polluted mangrove sediments represent key ecological niches for discovering microorganisms with specialized bioremediation traits [11]. However, heavy metals discharged into aquatic systems (e.g., Cd²⁺, Ni²⁺, Cu²⁺, Zn²⁺) pose severe ecological and health risks due to their persistence, bioaccumulation potential, and multi-organ toxicity [12].

The unique ecological context of moderately polluted mangrove ecosystems has yielded taxonomically diverse microbial resources with applied potential. For instance, distinct heavy metal-responsive traits were characterized in rhizosphere fungi [11], while the novel yeast Geotrichum sp. CS-67 (isolated from Shenzhen’s intertidal sediments) demonstrated a remarkable multi-metal accumulation hierarchy (Zn²⁺ > Ni²⁺ > Cu²⁺) through extracellular and intracellular strategies [11]. Similarly, the bacterium Aestuariibaculum sp. JKB11 (isolated from rhizosphere soils in Shenzhen mangroves) exhibited extracellular-dominant adsorption (97–99%) underpinned by 53 metal-resistance genes (e.g., CzcD, FieF), achieving unprecedented removal capacities for Cu²⁺ (280 mg/g) and Ni²⁺ (495 mg/g) when immobilized in the SA/PVA hydrogel through matrix synergy [13]. These examples collectively highlight the untapped bioremediation potential residing in mangrove microbiota, particularly for developing bio-based solutions targeting complex contamination scenarios. Developing such solutions is imperative given the escalating global burden of metal-contaminated industrial wastewater, particularly from electroplating, mining, and electronics sectors, where conventional physicochemical treatments face cost and sustainability limitations [14].

Experimental evidence further confirms the critical link between sediment conditions and functional microbial diversity. Despite documented heavy metal suppression effects on overall bacterial diversity (e.g., chromium toxicity [15], zinc stress [16]), moderately impacted mangroves maintain robust microbial communities through environmental compensation mechanisms [17]. Notably, mangrove rhizospheres yield exceptional microbial richness, as demonstrated by (1) Hao et al. [18] isolating 113 strains (predominantly Streptomyces/Bacillus) from Mai Po mangrove and (2) Shi et al. [19] recovering seven distinct fungal taxa from Kandelia obovata roots. Such diversity inherently enriches metal-adapted taxa, precisely the ecological niche, facilitating strains like our target organism, Sinirhodobacter sp. 1C5-22.

While evidence supports the remediation capabilities of mangrove microbes, translating laboratory findings to engineered applications requires bridging critical knowledge gaps. Critically, heavy metal toxicity fundamentally threatens bioremediation efficacy. Metals inhibit cell division, disrupt enzymatic functions, denature proteins, damage DNA integrity, and induce oxidative stress via reactive oxygen species accumulation [20]. Chronic exposure compromises metabolic homeostasis and triggers membrane lipid peroxidation, which is a key damage mechanism causing structural/functional deterioration [21,22]. Quantifying membrane impairment (e.g., via malondialdehyde [MDA], an established biomarker [23]) is thus essential to assess protective strategies.

The escalating heavy metal burden in coastal megacities like Shenzhen [5,24] demands sustainable solutions beyond costly physicochemical methods [25]. Microbial immobilization technology is critical for field-scale bioremediation because it overcomes inherent limitations of free bacterial cells, including vulnerability to hydrodynamic stresses, fluctuations in toxicity, and impractical recovery, by providing mechanical stability, metabolic protection, and reusability through matrix encapsulation (e.g., PVA-SA) [26]. Thus, while immobilization enhances biosorption stability and operational efficiency, its integration with understudied mangrove bacteria remains largely unexplored. Here, we address this gap by functionally characterizing a novel heavy metal-resistant bacterium, Sinirhodobacter sp. 1C5-22 isolated from the Shenzhen Futian Mangrove Reserve and pioneering its polyvinyl alcohol–sodium alginate immobilized consortium (PVA-SA 1C5-22) for industrial wastewater remediation. Our study focuses on (1) physiological tolerance profiling and adsorption dynamics of strain 1C5-22 across Cd²⁺, Ni²⁺, Cu²⁺ and Zn²⁺; (2) development and optimization of PVA-SA 1C5-22 for enhanced metal sequestration; and (3) validation of the consortium’s performance in real electroplating wastewater systems.

2. Materials and Methods

2.1. Collection of Rhizosphere Soil from Mangrove Plants

Soil samples were collected from the mangrove sediments in the tidal flat of Futian Mangrove Ecological Park, Shenzhen (22°31′ N, 114°00′ E). Plant materials, including leaves, roots and other debris, were carefully removed using sterilized shovels. All collected samples were immediately transported to the laboratory in sterile sealed bags and stored at 4 °C for subsequent analysis.

2.2. Isolation and Identification of Culturable Microorganisms from Rhizosphere Soil

2.2.1. Microbial Isolation and Cultivation

Standard serial dilution protocols were implemented for bacterial isolation. Surface soil and rhizosphere samples (1.0 g each) were vortexed with 9 mL sterile water in centrifuge tubes to create primary 10–1 suspensions. Subsequent sterile dilutions yielded concentration gradients from 10-2 to 10-6. Aliquots of 100 μL from each dilution series were uniformly plated on R2A agar (BD, San Diego, CA, USA) and maintained at room temperature for colony development with regular observation.

2.2.2. Colony Purification

Monoclonal cultures were obtained through sequential quadrant streaking. Representative colonies from initial isolation plates underwent 2–3 cycles of transfer onto fresh R2A medium until achieving axenic purity.

2.2.3. Molecular Characterization

Genomic DNA extraction from purified strains utilized the TIANamp Bacteria DNA Kit (Tiangen Biochemical Technology (Beijing) Co., Ltd., Beijing, China). Amplified 16S rDNA regions underwent electrophoretic validation on 1% agarose gels prior to bidirectional sequencing conducted by Qingke Biotechnology. Consensus sequences were analyzed through BLAST (https://www.ncbi.nlm.nih.gov/) comparisons via the NCBI platform.

2.2.4. Comprehensive Strain Profiling

Selected isolates underwent multiparametric analysis combining phylogenetic reconstruction and phenotypic characterization. MEGA7 software generated evolutionary trees using Sanger sequencing data cross-referenced with NCBI database entries. Morphological traits were systematically documented through microscopic examination. Growth optimization studies measured OD600 values across controlled temperature gradients, salinity variations (% NaCl), and pH ranges to establish ideal cultivation parameters.

2.3. Determination of Microbial Tolerance to Metal Ions

2.3.1. Initial Tolerance Screening

Microbial metal resistance was assessed using concentration-gradient plates. Purified strains were aseptically streaked onto R2A agar for individual heavy metal tolerance assessment under single-metal conditions, with cadmium tolerance tested using 0, 10, and 20 mg/L Cd²⁺ concentrations, nickel tolerance evaluated at 0, 50, 100, 200, 300, 400, 500, and 600 mg/L Ni²⁺, copper tolerance measured at 0, 50, 100, 200, 300, 400, 500, 600, and 700 mg/L Cu²⁺, and zinc tolerance analyzed at identical 0, 50, 100, 200, 300, 400, 500, 600, and 700 mg/L Zn²⁺ levels. Following seven-day incubation at ambient temperature, growth responses were methodically analyzed to identify tolerance limits. Visible colony development on metal-amended media constituted resistance verification. To ascertain maximum tolerance concentrations (MTCs), iterative culturing was performed with sequentially increased metal ion concentrations until achieving complete microbial growth suppression.

2.3.2. Assessment of Heavy Metal-Induced Oxidative Damage via Malondialdehyde (MDA) Quantification

Cellular oxidative damage induced by heavy metal stress was assessed by quantifying malondialdehyde (MDA) levels, a biomarker for lipid peroxidation. Experiments were conducted using both free Sinirhodobacter sp. 1C5-22 free cells and PVA-SA immobilized cells (PVA-SA 1C5-22) as described in Section 2.6.

Batch cultures were exposed to single-ion solutions (50 mg/L each of Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺). Samples were collected at time points (t = 0, 1, 5, 10, 15, 30, 60, 120, 180, 240 min) post-exposure for processing:

Free Cells: Cell pellets harvested by centrifugation (8000× g for 10 min at 4 °C) were resuspended in ice-cold extraction buffer provided in the Malondialdehyde (MDA) Assay Kit (Solarbio, Beijing, China). Bacterial cells were disrupted by sonication (200 W output power; 3 s pulse, 10 s pause, 30 cycles) on ice. The lysate was clarified by centrifugation (8000× g, 10 min, 4 °C) and the supernatant collected for assay.

Immobilized PVA-SA 1C5-22: Immobilized beads were collected by filtration/sieving, immediately frozen at −80 °C, and lyophilized for 24–48 h until constant weight was achieved. Dried beads were pulverized using a laboratory homogenizer/grinder. A known weight of powder was suspended in ice-cold extraction buffer, homogenized thoroughly on ice, and centrifuged (8000× g, 10 min, 4 °C). The supernatant was collected for assay.

MDA levels were quantified strictly according to the manufacturer’s instructions (Solarbio Malondialdehyde (MDA) Assay Kit; Beijing, China). Briefly, aliquot volumes of supernatant were mixed with assay reagents. Reaction mixtures were incubated at 100 °C for 60 min (tubes sealed tightly to prevent evaporation). Samples were then cooled on ice and centrifuged (10,000× g, 10 min, room temperature) to remove precipitate. Absorbance of the supernatant was measured at 532 nm and 600 nm using a microplate reader.

MDA concentration was calculated using the formula provided by the kit protocol:

MDA (nmol/g) = [ΔA × V_{_TotalReaction} ÷ (ε × d) × 109] ÷ (W × V_{_Sample} ÷ V_{_Extracted}) × F = 53.763 × ΔA ÷ W

where

ΔA532 = Δ532_{_sample}—Δ532_{_blank};

Δ600 = Δ600_{_sample}—Δ600_{_blank};

V_{_TotalReaction} = Total assay reaction volume (L);

ε = MDA molar extinction coefficient (1.55 × 105 L·mol⁻¹·cm⁻¹);

d = Light path length (cm)—specify plate type (e.g., standard 96-well plate, d≈0.6 cm);

W = Sample mass (g wet weight for free cells, dry weight for immobilized cells);

V_Sample = Volume of extract supernatant added to the reaction (L);

V_Extracted = Total extraction volume used per sample (L).

2.4. Multi-Omics Profiling of Heavy Metal Adsorption in Strain Sinirhodobacter sp. 1C5-22

To systematically investigate the heavy metal enrichment mechanisms of strain 1C5-22, a differential metal stress experimental design was implemented. Cultures in exponential growth phase were individually exposed to discrete metal solutions: 10 mg/L Cd²⁺, 50 mg/L Ni²⁺, 50 mg/L Cu²⁺, and 50 mg/L Zn²⁺. Exposure was performed in 250 mL Erlenmeyer flasks agitated at 180 rpm (28 °C orbital incubator) during two-hour metal enrichment stages. Pre-chilled PBS buffer was used for washing before collecting bacterial pellets through high-speed refrigerated centrifugation (Hitachi CR22GIII, Tokyo, Japan) at 8000× g for 15 min under 4 °C conditions. The experimental design comprised four treatment groups, including a blank control, each with three biological replicates. Cryopreserved samples stored at −80 °C were subjected to paired-end 150 bp transcriptome sequencing using the Illumina NovaSeq 6000 platform by Novogene Co., Ltd. (Beijing, China).

Raw sequencing data underwent quality assessment via FastQC v0.11.9 followed by rigorous filtering with Trimmomatic v0.39 (criteria Phred score ≥ 30, read length ≥ 50 bp). High-quality clean reads were assembled de novo using Trinity v2.13.2 with parameters min_kmer_cov = 2 and min_contig_length = 200. Assembly completeness was evaluated through BUSCO v5.2.2 against the bacteria_odb10 dataset, retaining unigenes with core gene coverage exceeding 80%. Functional annotation involved sequential BLASTn alignment against the NT database (E-value < 1 × 10⁻⁵), DIAMOND searches in the NR protein database (E-value < 1 × 10⁻⁵), and KEGG pathway annotation via KAAS using the SBH method (E-value < 1 × 10⁻¹⁰). Differentially expressed genes were identified through DESeq2 analysis (thresholds |log2FC| ≥ 1 and FDR < 0.05), followed by GO/KEGG enrichment analysis on the Novomagic platform employing Fisher’s exact test with adjusted p-values < 0.01.

Proteomic characterization was performed using the ExPASy ProtParam tool v1.0 to analyze amino acid composition features, including residue type distribution, total atomic count, molecular formula derivation, and relative molecular mass calculation based on isotopic averaging. Physicochemical properties encompassed isoelectric point prediction via the Bjellqvist algorithm, classification of charged residues (acidic/basic amino acids), and stability assessment (instability index > 40 indicating unstable proteins). Secondary structure prediction utilized the PSIPRED 4.0 workflow with a window size of 15 and a confidence threshold ≥ 70%, cross-validated by SOPMA for α-helix, β-sheet, and random coil proportions. Subcellular localization employed a hybrid strategy combining DeepLoc-1.0 deep learning models with CELLO v2.5 multi-classifier predictions. Transmembrane topology was analyzed using TMHMM 2.0 (confidence > 0.8) against the UniProtKB/Swiss-Prot database, while secretory potential was evaluated via SignalP-6.0 (D-score > 0.5 defining secretory proteins). Hierarchical three-dimensional structure modeling integrated Phyre2 homology modeling (coverage > 90%) with AlphaFold2 deep neural network predictions (pLDDT score > 70), followed by RosettaDock refinement and PROCHECK validation (Ramachandran plot favored regions > 85%).

2.5. Determine the Ability of Microorganisms to Remove Metal Ions

2.5.1. Biosorption Capacity Evaluation Protocol for Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺

Metal biosorption capacity evaluation followed the established methodology [9]. Cellular suspensions were prepared by culturing purified strains in SG broth under ambient conditions. Biomass separation involved processing cultures through centrifugation (4 °C, 8000 rpm, 8 min) followed by dual rinsing with ultrapure water to eliminate culture medium remnants. Washed pellets were individually exposed to separate 50 mg/L solutions of Cd²⁺, Ni²⁺, Cu²⁺, or Zn²⁺ under standardized agitation (200 rpm, 25 °C) for 120 min. Post-exposure, cell-free supernatants obtained via secondary centrifugation were analyzed for metal ion concentrations using ICP-OES (OPTIMA7000, Perkin Elmer, Hopkinton, MA, USA) to quantify remaining dissolved species. Biomass was quantified as dry weight after lyophilization.

The metal removal efficiency and biosorption capacity were quantitatively evaluated through two key parameters: removal rate (R) and biosorption quantity (Q). These metrics were calculated using the following equations:

R = (1 − C_t/C₀) × 100%

Q = (C₀ − C_t) × V/M

where

C₀ = Initial metal ion concentration (mg/L);

C_t = Equilibrium concentration after treatment (mg/L);

V = Solution volume (L);

M = Biomass weight (g).

2.5.2. Metal Uptake Equilibrium and Kinetic Analysis

Metal uptake dynamics were examined through equilibrium phase studies. Experimental systems contained standardized single-ion solutions of Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺ prepared at graduated concentrations (10, 25, 50, 100, and 150 mg/L) for adsorption isotherm analysis. Reaction vessels were maintained under orbital agitation (200 rpm) at 35 °C until equilibrium stabilization (120 min). Centrifugal separation yielded cell-free supernatants for elemental quantification through ICP-OES (OPTIMA7000, Perkin Elmer). Equilibrium binding profiles were derived via computational regression modeling of concentration-dependent adsorption data.

The Langmuir and the Freundlich isotherm models were employed to analyze the adsorption behavior. The equations are expressed as follows:

Langmuir equation [27]:

$${\displaystyle \frac{{\mathrm{C}}_{\mathrm{e}}}{{\mathrm{Q}}_{\mathrm{e}}}}={\displaystyle \frac{1}{\mathrm{b}{\mathrm{Q}}_{\max }}}+{\displaystyle \frac{{\mathrm{C}}_{\mathrm{e}}}{{\mathrm{Q}}_{\max }}}$$

Freundlich equation [28]:

$$\ln {\mathrm{Q}}_{\mathrm{e}}=\ln {\mathrm{k}}_{\mathrm{f}}+{{\displaystyle \frac{1}{\mathrm{n}}}\mathrm{C}}_{\mathrm{e}}$$

where

Q_e = Adsorption capacity at equilibrium (mg/g);

C_e = Equilibrium metal ion concentration in solution (mg/L);

Q_max = Maximum adsorption capacity (mg/g);

b = Langmuir equilibrium constant (L/mg);

k_f,n = Freundlich empirical constants.

Temporal variation in metal capture dynamics was examined using parallel experimental systems, each containing a separate 50 mg/L reference solution of Cd²⁺, Ni²⁺, Cu²⁺, or Zn²⁺. All reactions occurred under controlled rotary agitation (200 rpm) at 35 °C, incorporating scheduled sample collection points. Retrieved specimens underwent solid–liquid partitioning via centrifugal processing (8 min at 8000 rpm, 4 °C), with dissolved metal levels determined through inductively coupled plasma analysis (Perkin Elmer OPTIMA7000). Temporal binding curves were generated by applying algorithmic modeling approaches to time-series concentration measurements.

Complementing the equilibrium studies, kinetic analyses were conducted using 50 mg/L solutions with sampling at timed intervals. The temporal adsorption patterns were modeled using the following:

Pseudo-first-order kinetic model [29]:

$$\log ({\mathrm{Q}}_{\mathrm{e}}-{\mathrm{Q}}_{\mathrm{t}})=\log {\mathrm{Q}}_{\mathrm{e}}-{\displaystyle \frac{{\mathrm{k}}_1\mathrm{t}}{2.303}}$$

Pseudo-second-order kinetic model [30]:

$${\displaystyle \frac{\mathrm{t}}{{\mathrm{Q}}_{\mathrm{t}}}}={\displaystyle \frac{\mathrm{t}}{{\mathrm{Q}}_{\mathrm{e}}}}+{\displaystyle \frac{1}{{\mathrm{k}}_2{\mathrm{Q}}_{\mathrm{e}}^2}}$$

where

Q_e = Equilibrium adsorption capacity (mg/g);

Q_t = Adsorption capacity at time t (mg/g);

K₁ = Pseudo-first-order rate constant (min⁻¹);

K₂ = Pseudo-second-order rate constant (g·mg⁻¹·min⁻¹).

While these models are classically established, their contemporary formulation aligns with recent mechanistic interpretations [31,32].

2.5.3. Cellular Domain-Specific Metal Partitioning (Extracellular Adsorption, Intracellular Absorption, Structural Integration)

To clarify the binding processes within the microbial system, intracellular metal localization was examined following the experimental framework established in He et al. [11]. Single-ion solutions (Cd²⁺, Ni²⁺, Cu²⁺, Zn²⁺) with multiple concentration gradients were individually processed. Harvested bacterial biomass was treated with 25 mL of 0.1 mol/L EDTA-2Na solution under rotary agitation (30 min), with subsequent phase separation isolating supernatant containing surface-associated ions. Cellular pellets were reconstituted in fresh eluent and disrupted via ultrasonication-mediated membrane disintegration (30 min agitation post-treatment), yielding intracellularly retained metals in centrifuged supernatants. Residual cellular matrices were mineralized using 4 mL aqua regia (3:1 HCl:HNO₃ mixture), with dissolved metals representing structurally integrated fractions. This tripartite protocol systematically delineated metal distribution across cellular domains, in which the EDTA-elutable part was extracellular adsorption, the ultrasonic-released part was intracellular absorption, and the acid-digested part was a stable combination.

2.6. Preparation of Immobilized Bacterial Agents

The encapsulation of strain 1C5-22 was implemented via a hybrid embedding strategy combining sodium alginate (SA) and polyvinyl alcohol (PVA), following methodological optimizations reported by Dong et al. [33]. SA, a hydrophilic polysaccharide, served as the primary matrix owing to its biomaterial compatibility and ecological viability [34], characteristics critical for sustaining microbial viability in aqueous systems. To mitigate SA’s structural deficiencies, PVA—a cost-efficient polyhydroxy polymer—was integrated to improve mechanical resilience and microbial adhesion [35]. This dual-component system promoted intermolecular interactions that established a reinforced porous architecture, effectively eliminating issues like matrix brittleness and colloidal aggregation observed in single-component carriers. The resultant biocomposite achieved balanced bioactivity and operational stability under environmental conditions.

The protocol involved sequential steps: (1) Sterilization (121 °C, 20 min) of a 100 mL blend containing 14% PVA and 1% SA, alongside 500 mL of a saturated boric acid-5% CaCl₂ gelling solution; (2) Centrifugal concentration (4 °C, 8000 rpm, 10 min) of 400 mL bacterial broth to 35 mL, with 30 mL supernatant homogenized into the sterilized polymer mixture to prepare the composite inoculum; (3) Controlled extrusion (0.1 mL/min) of the inoculum into the agitated gelling medium (300 rpm), producing monodisperse hydrogel beads; (4) Post-synthesis processing via 4 h ice bath curing, filtration, and biomass quantification based on wet/dry weight ratios. Final products were refrigerated at 4 °C for deployment.

2.7. Determination of Desorption Performance of Immobilized Microbial Agents

2.7.1. Cyclic Adsorption–Desorption in Model Solutions with Immobilized Cells

To evaluate the reusability potential of immobilized beads, a five-cycle adsorption–desorption experiment was designed to specifically assess the desorption performance of PVA-SA immobilized strain 1C5-22. Based on previous findings demonstrating optimal desorption efficiency with 0.1 mol/L HCl (1 h reaction time), the experimental procedure initially prepared single-ion solutions containing Cd²⁺, Ni²⁺, Cu²⁺, or Zn²⁺ (each at 50 mg/L) for adsorption. After adsorption, the metal-loaded beads were rinsed with deionized water, then immersed in 25 mL of 0.1 mol/L HCl and shaken for 1 h. The resulting eluates were collected for heavy metal quantification by ICP-OES. This adsorption–desorption cycle was repeated five times to assess performance degradation. The desorption ratio (D) was calculated using:

D (%) = C_e/(C₀ − C_t) × 100%

where C_e represents the metal concentration in the eluate, and C₀ and C_t denote the initial and post-adsorption metal concentrations in the solution, respectively.

2.7.2. Validation via Electroplating Wastewater

Subsequently, industrial electroplating wastewater from Shenzhen Guihua Jiayi Industrial Park (with full-component analysis by ICP-OES) was tested to validate desorption efficiency under practical conditions. Pretreated immobilized microbial agents and blank PVA-SA beads were added to 25 mL of wastewater (triplicate groups) and subjected to five adsorption–desorption cycles. Variations in metal concentrations before and after adsorption (quantified identically) were compared to calculate enrichment ratios and capacities.

3. Results and Discussion

3.1. Adsorption and Molecular Mechanisms of Heavy Metal Resistance in Sinirhodobacter sp. 1C5-22

3.1.1. Heavy Metal Tolerance of Strain 1C5-22

A novel heavy metal-tolerant strain, Sinirhodobacter sp. 1C5-22 was isolated from rhizosphere sediments of Shenzhen Futian Mangrove Nature Reserve. This strain exhibits significant bioremediation potential as confirmed by: (1) 16S rRNA gene sequence analysis (99.11% similarity to Sinirhodobacter sp. B57, GenBank accession: MZ573293); (2) Phylogenetic classification within Rhodobacteraceae (Figure 1). The genus Sinirhodobacter, first described by Yang et al. [36], encompasses nine validated species with diverse metabolic functions (e.g., iron reduction [36], denitrification [37]), but crucially lacks reported heavy metal resistance, which is the knowledge gap addressed in this study. Physiological characterization determined optimal growth conditions for strain 1C5-22 as 35 °C, pH 7.6, and 3.8–4‰ salinity. Colony morphology analysis (Figure 1) revealed milky-white, slightly convex, smooth-surfaced circular colonies (<1 mm diameter) with entire margins. Microscopic examination identified Gram-negative, non-sporulating, aerobic, and motile cells exhibiting ovoid to short rod-shaped morphology. Following the systematic taxonomic identification and physiological characterization of strain 1C5-22, this study determined its heavy metal tolerance thresholds through gradient concentration experiments. The minimum inhibitory concentration (MIC) results demonstrated significant differences in the tolerance of the strain to Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺. The MIC for Cd²⁺ was 20 mg/L, while those for Ni²⁺, Cu²⁺, and Zn²⁺ exceeded 500 mg/L, 600 mg/L, and 600 mg/L, respectively. Notably, compared to previously reported non-metal functions of Sinirhodobacter (e.g., nitrogen fixation, Fe³⁺ reduction [36,37]), this work provides the first evidence of extreme tolerance to Cu²⁺/Zn²⁺ (≥600 mg/L) and Ni²⁺ (>500 mg/L), establishing Cd²⁺ << Ni²⁺ < Cu²⁺ ≈ Zn²⁺ as its metal resistance hierarchy. Furthermore, the tolerance levels substantially surpassed typical heavy metal background values in mangrove sediments, indicating promising bioremediation applications in anthropogenically highly polluted environments, such as industrial wastewater systems. These findings establish critical concentration parameters for subsequent investigations into the strain’s heavy metal adsorption dynamics and molecular resistance pathways.

3.1.2. Extracellular/Intracellular Metal Distribution in Strain 1C5-22

The practical efficacy of bacterial strains in remediation scenarios depends not only on their tolerance thresholds but also on dynamic adsorption capacities for heavy metal ions. Building upon previous tolerance studies, this work systematically evaluated the heavy metal enrichment characteristics through domain-specific fractionation (Section 2.5.3). Experimental results revealed significant differences among the extracellular (EDTA-elutable), intracellular (ultrasonically released), and structural (acid-digested) phases (Figure 2). Among them, Cd²⁺ and Ni²⁺ primarily rely on extracellular binding/sequestration (accounting for 84% and 81% of their total content, respectively), effectively avoiding their intracellular toxicity. This finding is consistent with the results of Zhang et al. [38], who observed a similar phenomenon: under heavy metal stress, Vibrio cholerae reduced cell membrane fluidity and enhanced surface hydrophobicity, thereby restricting the entry of heavy metals into the cells. In contrast, Cu²⁺ tends to penetrate and accumulate intracellularly (accounting for nearly 60% of its total content). This pattern aligns with the findings of Mergeay et al. [39], namely that Ralstonia metallidurans actively transports Cu²⁺ into cells via plasmid-encoded metal resistance genes (such as the cnr gene) or immobilizes it within cells through chelation mechanisms, thereby enhancing tolerance. Conversely, the distribution of Zn²⁺ demonstrates another strategy—a balance between extracellular interception and intracellular maintenance (extracellular distribution approximately 61%, intracellular distribution approximately 39%). This point is also supported by the research of El-Deeb et al. [40], who found that Enterobacter sp. can enhance its tolerance by secreting metal-binding proteins/ion transporters to restrict Zn²⁺ extracellularly or by modulating cell membrane permeability to reduce its entry into cells. These quantified distribution differences provide crucial evidence for gaining an in-depth understanding of the specific resistance mechanisms of this strain to different heavy metals.

3.1.3. Molecular Architecture of Heavy Metal Resistance in Sinirhodobacter sp. 1C5-22

Strain 1C5-22 exhibits a paradoxical phenotype: its adsorption efficiency for heavy metals declines at elevated concentrations, yet it displays exceptional tolerance (e.g., MIC_Cu²⁺ ≥ 600 mg/L). This disparity strongly suggests the evolution of specialized, multi-layered molecular resistance mechanisms beyond mere passive adsorption. To elucidate the molecular basis underlying this concentration-dependent attenuation in adsorption and the extreme tolerance, we employed an integrated analysis of transcriptomics (under Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺ stress) and protein structure prediction. Building upon our previous high-quality genomic data (>96% mapping rate), our primary aim was to identify the core regulatory network and functional adaptations responsible.

Transcriptomic analysis of 50 million clean reads per replicate (BUSCO completeness >92%) identified CreB as a central regulator. Structural validation showed key proteins possessed predicted Cd²⁺-binding domains: PROKKA_02958 as secretory (DeepLoc: cytoplasmic score = 0.11), PROKKA_03231 as stable extracellular (instability index = 32.7), and CreB transmembrane topology (TMHMM: 4 domains).

Transcriptomic profiling revealed distinct metal-specific responses, with Cu²⁺ exposure inducing the most substantial upregulation (228 genes), correlating molecularly with its extreme copper tolerance (MIC ≥ 600 mg/L, Figure S1E). Crucially, cross-metal comparative analysis identified CreB (encoded by PROKKA_02312) as a central regulatory hub, consistently activated under all tested metal stresses. Functional characterization indicates that CreB integrates three major defense pathways (Table S1): (1) Directing RND efflux pumps and metallothionein synthesis (efflux/sequestration); (2) Mediating ribosomal repair via rRNA modification and compensating for two-component signaling (cellular maintenance); (3) Activating antioxidant defense enzymes (oxidative stress mitigation). This multifaceted role parallels global regulators like Spo0A, underscoring CreB’s pivotal position.

Integrated analysis validated the Adsorption–Sequestration–Efflux (ASE) defense paradigm. GO/KEGG enrichment (Figures S2 and S3) confirmed activated pathways critical to metal resistance–including ribosomal repair, quorum sensing, two-component systems, and oxidative stress responses–aligning with CreB’s regulatory functions and extending established networks [41,42]. Crucially, structural predictions identified key proteins underpinning each ASE layer (Table S1): PROKKA_02958 and PROKKA_01213 were characterized as unstable secretory proteins, likely mediating rapid extracellular polymer adsorption (A); PROKKA_03231 emerged as a stable periplasmic protein with mixed α/β metal-binding domains, enabling periplasmic sequestration/buffering (S); while CreB’s RND efflux pump regulation substantiated transmembrane efflux capacity (E). This collective evidence solidifies the ASE tripartite architecture as the molecular foundation for sustained extreme metal tolerance amid concentration-dependent adsorption decline, the latter potentially exacerbated by metal-induced structural damage (e.g., carboxyl group interactions [43]).

Collectively, this study defines the core molecular architecture of heavy metal resistance in Sinirhodobacter sp. 1C5-22: a CreB-coordinated, tri-layered ASE defense system (Adsorption–Sequestration–Efflux). This integrated architecture explains the strain’s unique tolerance–adsorption paradox and provides a mechanistic understanding beyond previously described systems. This framework directly informs future efforts: experimentally confirming protein localization (e.g., signal peptides), identifying precise metal-binding sites via docking, and targeting CreB networks or protein modification for engineered bio-remediation applications.

3.2. Application of 1C5-22 Immobilized Bacterial Agent for Heavy Metal Removal

3.2.1. Enhanced Metal Enrichment via SA/PVA Immobilization of Strain 1C5-22

To enhance the stability and operability of strain 1C5-22 in practical complex environments, this study developed a polyvinyl alcohol–sodium alginate immobilized bacterial agent system (PVA-SA 1C5-22) based on its heavy metal adsorption characteristics in the free state. The immobilization technology, through the synergistic effects of physical embedding and chemical crosslinking, effectively addresses application limitations such as the susceptibility of free cells to loss and weak resistance to environmental disturbances while enhancing the interfacial interaction efficiency between the strain and heavy metals [15,44]. This section systematically investigates the dynamic adsorption performance and reusability potential of the immobilized bacterial agent for heavy metal ions, extends the research to real industrial wastewater scenarios to evaluate its simultaneous removal efficiency for multi-metal composite pollution systems, and provides theoretical and technical support for the engineering application of strain 1C5-22.

While quantification of sorptive properties intrinsic to the carrier matrix could theoretically provide supplementary mechanistic insight, their deliberate exclusion in the present investigation adheres to established methodological conventions for immobilized microorganism systems where biological transformation constitutes the primary remediation pathway [45,46,47,48,49]. This prioritization reflects the field’s consensus, articulated in seminal frameworks and reinforced by contemporary high-impact investigations, that analyses should emphasize functional biomass preservation within synthetic matrices rather than deconvoluting passive contributions of inert structural components. Compelling precedents exist in the recent literature, including rigorous studies by Yang et al. [47] and He et al. [48], which explicitly focus on viability-sustaining capacities of encapsulation architectures as the critical determinants of biocatalytic resilience under operational stressors.

At a low concentration of 10 mg/L (Figure 3), PVA-SA 1C5-22 demonstrated a significant enrichment advantage over its free-cell counterpart: The enrichment efficiencies for Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺ reached 78.59%, 93.20%, 90.89%, and 58.46%, respectively. These values were significantly higher than those observed for the free cells (67.91%, 70.09%, 75.17%, and 40.14%, respectively), representing increases of 11.68 to 23.11 percentage points. Concurrently, the enrichment capacities of PVA-SA 1C5-22 attained 31.85 mg/g Cd, 37.31 mg/g Ni, 44.23 mg/g Cu, and 24.83 mg/g Zn, exceeding those of the free cells by 21.22 to 36.54 mg/g.

Even under high-concentration stress (50 mg/L), the enrichment superiority of PVA-SA 1C5-22 remained evident (Figure 3). The enrichment efficiencies for Cd, Ni, Cu, and Zn were 30.77%, 40.22%, 65.29%, and 46.96%, respectively. Compared to the free cells (19.27%, 23.87%, 43.32%, and 33.99%, respectively), this represented improvements of 11.50 to 21.97 percentage points. However, the enhancement in enrichment capacity relative to free cells narrowed to 6.03–8.97 mg/g at this concentration, stabilizing at final values of 17.27 mg/g Cd, 16.24 mg/g Ni, 16.34 mg/g Cu, and 12.16 mg/g Zn. It is noteworthy that the enrichment efficiency of the free cells decreased drastically with increasing concentration. For instance, the Cd enrichment efficiency plummeted from 67.91% (at 10 mg/L) to 19.27% (at 50 mg/L), a decrease of 71.6%. In contrast, the Cd enrichment efficiency within PVA-SA 1C5-22 declined more gently from 78.59% to 30.77%, a reduction of 60.8%.

3.2.2. Oxidative Damage Mitigation via Immobilization in Strain 1C5-22

The findings in this section demonstrate the protective capacity of immobilized materials for microbial strains under heavy metal stress. Experimental data revealed that free-living 1C5-22 cells exposed to 50 mg/L Cd²⁺ exhibited a rapid surge in MDA content to 38.88 nmol/g within 1 min, followed by a sharp decline to 5.81 nmol/g at 60 min and stabilization at approximately 15 nmol/g (Figure 4A). In contrast, immobilized cells maintained MDA levels below 5 nmol/g throughout the exposure period, demonstrating the efficacy of embedding materials in mitigating Cd²⁺-induced oxidative damage. Under Ni²⁺ stress (Figure 4B), free cells displayed biphasic MDA dynamics, with an initial decrease within 10 min followed by a peak of 27.6 nmol/g and subsequent stabilization at 15 nmol/g after 30 min. Although immobilized cells showed a transient increase at 15 min, they stabilized at lower levels by 30 min, indicating superior tolerance. In Cu²⁺-stressed systems (Figure 4C), free cells exhibited an initial MDA spike of 55.43 nmol/g, declining to 45.13 nmol/g at 15 min and fluctuating between 45 and 49 nmol/g thereafter. Immobilized cells consistently maintained stable MDA levels near 3 nmol/g. For Zn²⁺ exposure (Figure 4D), free cells reached a peak MDA concentration of 183.48 nmol/g at 5 min, gradually decreasing to 72.54 nmol/g after 4 h, while immobilized cells started at 70.64 nmol/g and steadily declined to 30–50 nmol/g.

A notable phenomenon was observed under Ni²⁺ stress. Immobilized cells initially exhibited higher MDA levels than free cells during the early phase, but rapidly decreased to lower values after the peak, whereas free cells sustained elevated MDA levels. This suggests that immobilization may confer long-term protection through delayed response mechanisms. Collectively, immobilization technology significantly reduced MDA fluctuations in strain 1C5-22 across all tested heavy metals, with particularly pronounced stabilization effects against Ni²⁺ and Cd²⁺. These findings imply that embedding materials alleviate direct heavy metal toxicity through dual mechanisms of physical barrier and chemical adsorption, thereby preserving microbial metabolic activity [16,17].

3.2.3. Adsorption Behavior and Sustainable Removal Performance of PVA-SA 1C5-22

Having established the significant protective role of SA/PVA immobilization against heavy metal-induced membrane damage in strain 1C5-22, its potential for robust field applications becomes evident. To advance towards practical implementation, we next assessed the operational robustness and reusability of the immobilized system (PVA-SA 1C5-22) under simulated industrial remediation scenarios, specifically targeting wastewater with extreme heavy metal loads.

To establish a baseline for assessing long-term stability, we first characterized the fundamental metal binding differences between free strain 1C5-22 and its PVA-SA immobilized counterpart (PVA-SA 1C5-22) using adsorption models. Langmuir and the Freundlich isotherm analyses defined equilibrium capacities and adsorption patterns, while pseudo-first and pseudo-second order kinetics elucidated rate-determining steps (Figure 5, Tables S2 and S3).

For free cells, Cd²⁺ adsorption conformed strongly to the Langmuir model (R² = 0.93), indicating homogeneous, monolayer binding. In contrast, adsorption of Ni²⁺, Cu²⁺, and Zn²⁺ better fitted the Freundlich models (R² = 0.79–0.83), suggesting heterogeneous, multilayer interactions (Figure 5A,B, Table S2). This aligns with established species-specific behaviors: Langmuir adsorption for Cd²⁺ in Pseudomonas aeruginosa N6p6 [50] and Freundlich-type Ni adsorption in Escherichia coli Ws11 [51]. Immobilization has induced distinct changes in adsorption affinities and mechanisms. Immobilized cells (PVA-SA 1C5-22) exhibited Langmuir-type behavior for both Cd²⁺ and Zn²⁺ (R² = 0.91–0.94), indicating a shift towards more homogeneous binding sites compared to free cells for Zn²⁺, while Freundlich characteristics persisted for Ni²⁺ and Cu²⁺ (R² = 0.81–0.85) (Figure 5C,D, Table S2). This transition of Cd²⁺ and Zn²⁺ to Langmuir behavior resembles the pattern observed in composite systems like Pseudomonas putida-clay for Pb²⁺/Cd²⁺ [52].

Kinetic analyses revealed further mechanistic complexity and immobilization effects (Figure 5E,F, Table S3). Adsorption onto both free and immobilized cells for Cd²⁺, Ni²⁺, and Zn⁺² was predominately governed by pseudo-second-order kinetics (R² > 0.93 for free, >0.95 for immobilized Cd²⁺/Ni²⁺; R² > 0.95 for immobilized Zn²⁺), strongly indicative of chemisorption involving valence forces, such as electron transfer or ionic bonding. The significantly enhanced fit (R² = 0.98 compared to 0.96 for free) for Cd²⁺ adsorption on immobilized cells suggests the PVA-SA matrix provides additional, potentially more accessible, chemisorption sites. For Cu²⁺, the poor pseudo-second-order fit (R² = 0.82 for free) implied multi-step adsorption or significant heterogeneity in binding sites, potentially involving initial rapid binding facilitated by the Jahn–Teller distortion characteristic of Cu²⁺ ions [53]. Zn²⁺ adsorption also showed nuances consistent with its tendency for weaker coordination complexes and stable hydration shells, indicating multi-step mechanisms potentially involving hydration shell disruption before binding [54].

Moreover, FTIR spectroscopy of free 1C5-22 and immobilized PVA-SA 1C5-22 (Figure S4) before and after metal exposure identified critical functional groups participating in chemisorption through characteristic band shifts: For Cd²⁺ adsorption, free cells exhibited strong amide I/II band changes (1617–1530 cm⁻¹, indicating protein-associated binding), while the immobilized system showed supplementary engagement of –OH and –COO⁻ groups. Ni²⁺ and Cu²⁺ adsorption in both systems demonstrated dominant –NH/–OH interactions (3300–2900 cm⁻¹ shifts) and carboxylate participation (–COO⁻ asymmetric stretch at 1600–1650 cm⁻¹). Zn²⁺ adsorption broadly involved phosphates (P–O at 1050–1100 cm⁻¹), hydroxyls, and carboxylates across both formats. These results confirm chemisorption is mediated primarily by microbial surface groups (–COO⁻, –NH₂, –OH, C = O, P–O), with immobilization enhancing accessibility/reactivity of these moieties as inferred from kinetic models—consistent with studies of bacterial metal coordination via carboxylates/amines [55] and functional group synergies in bio-immobilized systems [56,57].

These kinetic deviations align with intrinsic coordination properties of Cu²⁺ and Zn²⁺ that necessitate sequential adsorption steps: (1) Rapid surface coordination to carboxylates/amines driven by electrostatic forces, (2) Slower reorganization step—specifically Jahn-Teller distortion-driven ligand rearrangement for Cu²⁺ and displacement of Zn²⁺ hydration shells at phosphate sites, and (3) Final chemisorption stabilization (e.g., Cu²⁺-amide covalent bonding inferred from FTIR band shifts; Zn²⁺-phosphate P-O bond distortion ≈6% increase at 1050 cm⁻¹). This mechanistic framework resolves the poor fit of single-step models for Cu²⁺/Zn²⁺ by explicitly incorporating structural transitions (inferred from spectroscopy and hydration theory [53,54]), without requiring empirical fitting adjustments.

Collectively, these modeling studies highlight two key aspects. First, the PVA-SA matrix significantly alters metal adsorption behavior compared to free cells, particularly promoting Langmuir-like adsorption for Cd²⁺ and Zn²⁺ while also enhancing Cd²⁺ chemisorption kinetics. This suggests the carrier induces modifications such as creating a more uniform distribution of active sites or facilitating access due to changes in physical architecture, including porosity and surface functional groups [58]. Second, adsorption pathways are intrinsically dependent on metal-specific coordination chemistry, requiring distinct transition steps: Cd²⁺ and Ni²⁺ follow rapid chemisorption dominated by direct binding to carboxylates/amines [59]; Cu²⁺ adsorption necessitates structural reconstruction via Jahn–Teller distortion to achieve stable coordination geometries [60]; Zn²⁺ adsorption requires prior hydration layer shedding at phosphate clusters before effective ligand coordination [45]. These differences arise from distinct ionic properties, such as radius, charge density, hydration energy, and ligand preference, which govern how ions interact with critical surface functional groups like carboxylates and phosphates involved in biosorption [61].

While literature suggests hydration dynamics may influence Zn²⁺ coordination [15], our FTIR kinetics confirm functional group binding ultimately governs adsorption efficiency under operational conditions. To validate the practical operation and reusability essential for field deployment, PVA-SA 1C5-22 underwent five consecutive adsorption–desorption cycles under simulated high-load industrial wastewater conditions. This rigorous testing revealed distinct metal retention patterns, with Cd²⁺ enrichment rates showing adaptive improvement and exceeding 20% retention after the first cycle, while Ni²⁺, Cu²⁺, and Zn²⁺ enrichment stabilized at moderate recovery rates (20–25%) following an initial decline, suggesting differential metal resilience (Figure 6A–D). Efficient regeneration occurred using HCl, achieving desorption efficiencies over 70% for Cd²⁺ and Cu²⁺, and exceeding 55% for Ni²⁺ and Zn²⁺; this performance surpasses immobilized systems like Pseudomonas aeruginosa and Bacillus cereus [62,63], likely due to effective proton competition disrupting metal-site bonds within the optimized PVA-SA matrix in acidic conditions. Furthermore, PVA-SA 1C5-22 demonstrated robust reusability, with the adaptive enhancement in Cd²⁺ enrichment hinting at beneficial physiological or metabolic adaptations of the immobilized strain under metal stress, a novel insight meriting investigation for sustainable bioremediation. The synergistic combination of the tailored PVA-SA carrier and microbial properties effectively overcame specificity and durability challenges common to immobilized biosorbents.

3.2.4. Performance Validation in Real Electroplating Wastewater

Building upon the robust performance established in Section 3.2.3 under simulated multi-metal conditions, we validated the efficacy and functional stability of PVA-SA 1C5-22 under a critical real-world challenge: treating non-synthetic wastewater sourced directly from an electroplating facility. Crucially, comparative testing with authentic electroplating wastewater revealed the immobilized system’s decisive superiority: PVA-SA 1C5-22 maintained measurable Ni adsorptive function (≈20% removal) under extreme 1850 mg/L stress while free 1C5-22 cells suffered complete metabolic collapse (0% removal efficiency; Figure 6F). This effluent presented a formidable remediation hurdle characterized visually by dark green clarity and analytically by Ni²⁺, constituting 99% of its components at an extreme concentration of 1850 mg/L. While electrochemical precipitation remains optimal for reducing initial nickel loads (e.g., 1850 mg/L → ~50–200 mg/L), PVA-SA 1C5-22 delivers reliable bio-polishing at such post-precipitation concentrations: achieving > 90% efficiency at 10 mg/L (Figure 3B) and 40% efficiency at 50 mg/L, significantly outperforming free cells which collapse under these loads (Section 3.2.1). The validation at 1850 mg/L herein served to probe extreme functional stability boundaries.

The PVA-SA 1C5-22 was directly applied to this authentic industrial effluent. Crucially, it maintained stable performance through five consecutive adsorption–desorption cycles with the real electroplating effluent (Figure 6E,F). Experimental results demonstrated consistent Ni²⁺ adsorption rates of approximately 20% and desorption efficiencies exceeding 50% across all cycles. The obtained residual Ni²⁺ concentration (~1480 mg/L) reflects the adsorption capacities achievable under these acute stress conditions. Under these operational conditions with a fixed biomass dosage, the system may have approached its functional adsorption capacity threshold at the extreme Ni²⁺ concentration (1850 mg/L). Material loading optimization, such as increased bead dosage or multi-stage configurations, could potentially enhance removal efficiency for similar ultra-high concentration scenarios in future implementations. The validation goals for this extreme concentration scenario were twofold: (a) demonstration of sustained structural integrity/reusability through multiple adsorption–desorption cycles under severe Ni²⁺ stress; and (b) confirmation of functional metabolic preservation in encapsulated biomass, evidenced by stable desorption efficiencies (>50%) across cycles (Figure 6E,F), which overcomes the critical collapse of free-cell systems under comparable loads. Nevertheless, these results suggest integration potential: At industrially relevant post-precipitation concentrations (≤50 mg/L Ni²⁺), PVA-SA 1C5-22 exhibits significantly higher adsorption efficiencies (≥40–93%, Section 3.2.1, Figure 3B,D), warranting future work on sequential treatment process optimization. The sustained robustness demonstrated in both structural integrity and preserved metabolic function unequivocally establishes PVA-SA 1C5-22 as a technically viable enhancer for hybrid remediation systems facing severe nickel loads.

This sustained efficacy under the stress conditions of the actual electroplating effluent originates from synergistic protective and adsorptive mechanisms enabled by the PVA-SA immobilization matrix. The three-dimensional polymeric architecture not only substantially reduced direct Ni²⁺ toxicity to the embedded microbial strain through effective physical shielding but also synergistically enhanced volumetric Ni²⁺ adsorption capacity via complementary interactions between the carrier’s embedded sorption sites and the microbial surface binding domains. Concurrently, the stabilized microenvironment preserved essential metabolic activity, enabling coordinated extracellular biosorption and intracellular accumulation processes. Such integrated carrier–microbe coupling mechanisms effectively counter the operational instability typically exhibited by free cells in complex, high-load wastewater systems, thereby overcoming critical limitations inherent to conventional bioremediation approaches for extreme industrial contaminants like nickel.

4. Conclusions

This study establishes Sinirhodobacter sp. 1C5-22, isolated from Shenzhen Futian mangrove rhizosphere sediments, as a novel and exceptionally resilient bacterium with significant bioremediation potential for heavy metal pollution. We present the first evidence of remarkable heavy metal tolerance within the genus Sinirhodobacter, demonstrating notably high MICs for Cu²⁺, Zn²⁺ (≥600 mg/L), and Ni²⁺ (>500 mg/L). Critically, investigations revealed distinct, metal-specific intracellular vs. extracellular distribution patterns: Cd²⁺ and Ni²⁺ primarily underwent extracellular sequestration (>80%), while Cu²⁺ accumulated intracellularly (~60%); Zn²⁺ exhibited a balanced distribution.

Integrative transcriptomic and protein structural analyses unveiled the sophisticated molecular architecture underpinning this tolerance. We identified the CreB transcriptional regulator as the central hub coordinating a multi-layered defense system–the Adsorption–Sequestration–Efflux (ASE) tripartite mechanism. CreB orchestrates (1) extracellular polymer adsorption via unstable secretory proteins (e.g., PROKKA_02958), (2) periplasmic buffering/sequestration by stable metal-binding proteins (e.g., PROKKA_03231), and (3) RND efflux pump-mediated efflux, alongside ribosomal repair and antioxidant activation. This integrated ASE system resolves the apparent paradox of declining adsorption efficiency at high concentrations, simultaneously with extreme tolerance.

Translating fundamental insights into practical application, we developed a polyvinyl alcohol–sodium alginate immobilized bacterial agent (PVA-SA 1C5-22) that significantly outperformed free cells, demonstrating enhanced adsorption with superior enrichment capacity and efficiency for Cd²⁺, Ni²⁺, Cu²⁺, and Zn²⁺ (especially at environmentally relevant low concentrations, elevating Cd enrichment by ~11.7–23.1 percentage points at 10 mg/L). It demonstrated robust protection through effective mitigation of HM-induced oxidative damage (notably for Cd²⁺ and Ni²⁺), evidenced by suppressed/stabilized malondialdehyde (MDA) levels via synergistic physical and chemical barriers preserving metabolic activity. We also observed excellent operational stability featuring reusability over five adsorption–desorption cycles in simulated high-load, multi-metal wastewater with retained enrichment (20–25% for Ni, Cu, Zn) and adaptive improvement for Cd, alongside efficient desorption (>55–70% using HCl), confirming regeneration feasibility. Finally, critical field relevance was assessed through proof-of-concept efficacy in authentic electroplating wastewater (1850 mg/L Ni²⁺), maintaining ~20% adsorption and >50% desorption efficiency across multiple cycles.

Collectively, this work bridges fundamental microbial physiology with environmental biotechnology: (1) It provides unprecedented mechanistic insights into heavy metal resistance in Sinirhodobacter through the CreB-mediated ASE paradigm. (2) It also delivers a demonstrably effective, immobilized bacterial agent (PVA-SA 1C5-22) with enhanced durability, protection, adsorption performance, and reusability for real-world remediation scenarios under extreme HM stress. This study underscores mangrove ecosystems as rich reservoirs of stress-adapted microbes and positions Sinirhodobacter sp. 1C5-22 is a promising candidate for developing engineered solutions targeting complex industrial heavy metal contamination.