Immobilized Pseudomonas fluorescens Lipase on Eggshell Membranes for Sustainable Lipid Structuring in Cocoa Butter Substitute

Abstract

As the supply of cocoa becomes increasingly volatile, biotechnological innovations such as lipid engineering with lipases play a crucial role in supporting more stable, ethical, and sustainable chocolate production systems. This study explores the potential of Pseudomonas fluorescens lipase immobilized on eggshell membrane-based carriers for the synthesis of a cocoa butter substitute (CBS). The carriers were prepared by treating eggshells with different acids to generate chemically distinct support materials. Lipase immobilization was performed using both adsorption and covalent binding techniques. All resulting biocatalysts were characterized and compared to the free enzyme with respect to pH and temperature optima, as well as thermal and solvent stability. Immobilization caused shifts in the enzyme’s optimal operating conditions and significantly improved its stability at elevated temperatures and in the presence of organic solvents. Among the tested systems, the lipase immobilized by adsorption onto a hydrochloric acid-treated carrier exhibited the best performance. Using this biocatalyst, a CBS containing 93.54 ± 0.16% of the target triacylglycerols (POP, POS, and SOS) was successfully synthesized and reused over five consecutive synthesis cycles without significant loss of activity. These findings demonstrate the potential of waste-derived biomaterials for the development of efficient, stable, and reusable biocatalysts in the enzymatic production of functional lipids.

1. Introduction

The rising cost and fluctuating availability of cocoa butter (CB) have raised significant concerns in the global confectionery industry [1,2] due to climate variability, pest infestation, and socio-economic instability in key producing regions [3,4,5]. This growing problem has driven the development of cocoa butter alternatives that can fully replace CB but require specific processing conditions [6,7]. These alternatives are generally divided into three categories: cocoa butter equivalents (CBEs), cocoa butter substitutes (CBSs) and cocoa butter replacers (CBRs) [1,8,9,10]. CBEs are designed to closely mimic the triacylglycerol (TAG) profile of cocoa butter—particularly its high contents of POP (1,3-dipalmitoyl-2-oleoylglycerol), POS (1-palmitoyl-2-oleoyl-3-stearoylglycerol), and SOS (1,3-distearoyl-2-oleoylglycerol)—and are typically compatible with CB in any proportion [8,11,12]. CBSs, which are often derived from lauric oils such as palm kernel oil or coconut oil, can replace CB entirely; however, they are incompatible in blending due to their different polymorphic behavior [6,7]. CBRs, meanwhile, are non-lauric fats that resemble CB in physical properties but differ in chemical composition and are mainly used in low-fat or compound chocolate applications [13,14]. Although CBEs are generally considered the most desirable alternative due to their compositional and functional similarity to cocoa butter, this study focused on the synthesis of a CBS, primarily because of the availability of lauric-rich raw materials and the catalytic specificity of the selected lipase, which is more suitable for processing such substrates. However, it is important to recognize that CBS and CBE have different advantages and limitations that affect their industrial applicability. CBEs are very similar to the triacylglycerol profile of CB and offer better compatibility in blending and final product texture, so they are favored for formulations that require the partial replacement of CB. On the other hand, while CBSs can fully replace CB, they often exhibit polymorphic behaviour, limiting their compatibility with natural CB and potentially affecting the crystallization and sensory properties of the final product. In addition, CBSs derived from lauric oils benefit from the abundant availability and lower cost of the raw materials; however, they can cause problems in processing due to these physicochemical differences. Thus, while CBEs are favored for their compositional fidelity and functionality, CBSs remain a practical and economically viable alternative, especially when substrate availability and enzymatic specificity are considered. A promising strategy for producing high-quality CBEs is enzymatic interesterification using lipases, especially sn-1,3-regioselective variants such as Pseudomonas fluorescens (PFL). This method restructures TAG molecules in plant oils to mimic the composition and functionality of CB [15,16]. Immobilized enzymes offer advantages such as improved stability, specificity, and reusability, which reduce costs and environmental impact [17,18]. Despite challenges related to process scalability, raw material availability, enzyme cost, and regulatory acceptance, the enzymatic production of CB alternatives using immobilized lipases represents a sustainable and viable long-term solution for the confectionery industry.

To support the efficiency and sustainability of such biocatalytic processes, especially those involving the lipase-catalyzed synthesis of cocoa butter alternatives, increasing attention is being directed toward the development of low-cost, eco-friendly immobilization systems based on waste-derived materials.

The growing demand for sustainable biocatalytic processes has increased interest in cost-effective, environmentally friendly materials for immobilizing enzymes. In this context, waste from the agri-food industry is increasingly viewed not as waste but as a valuable secondary raw material [19]. Its valorization is fully in line with the principles of the circular bioeconomy, which prioritize resource efficiency, the reduction of environmental impact, and the replacement of traditional, often costly immobilization carriers. Carriers derived from agricultural and industrial waste have proven to be a promising alternative to traditional immobilization materials. These waste-derived carriers offer a sustainable, cost-effective, and circular solution for producing biocatalysts, especially for large-scale applications. Their significantly lower cost compared to commercial carriers reduces the overall cost of immobilised enzyme systems and thus improves the economic feasibility of biotechnological processes. Apart from the cost benefits, the use of waste materials contributes to environmental sustainability, as organic residues do not have to be disposed of in landfills and the associated pollution is reduced. Most of these materials are inherently biodegradable and biocompatible, minimizing toxicity concerns and simplifying end-of-life disposal. In addition, their inherent structural and chemical properties often allow for higher enzyme loading, improved mass transfer, and in some cases, increased protection of enzyme activity under operating conditions.

Various types of immobilization carriers derived from agri-food waste can be classified according to their origin—lignocellulosic materials [20,21,22,23], fruit and vegetable wastes [24], mycelium waste [25], and eggshells and eggshell membranes. These calcium carbonate-rich eggshell residues contain a collagenous inner membrane with favorable mechanical and biochemical properties. Their fibrous matrix offers a biologically compatible surface suitable for enzyme immobilization, particularly when modified through acid or thermal treatment to enhance binding efficiency [19].

The valorization of waste-derived materials for enzyme immobilization not only reduces processing costs but also supports a more sustainable and circular approach in industrial biotechnology. However, the use of such materials is also associated with some challenges. Their properties can vary significantly depending on their origin and previous treatment, which can affect the reproducibility and performance of immobilized enzymes. In many cases, physical or chemical pre-treatment, such as alkaline activation, lignin removal, or acid washing, is required to expose functional groups, increase surface porosity, or remove inhibitory substances. In addition, certain immobilization techniques or carrier properties can negatively affect enzyme activity due to steric hindrances, restricted diffusion, or the creation of unfavorable microenvironments around the active site.

Among the various waste-based carriers investigated, acid-treated eggshell membrane (ESMC) was given special attention in this study due to its favorable biochemical composition and availability as a by-product of the food industry. However, despite these promising properties, its suitability as a carrier for immobilizing lipase has not yet been sufficiently researched. This study aims to evaluate the efficacy of ESMC as a carrier for immobilizing sn-1,3-regioselective lipase from Pseudomonas fluorescens (PFL). In particular, adsorption and covalent binding immobilization methods will be compared to identify the optimal approach for this carrier. Furthermore, this study investigates the applicability of the immobilized lipase in the enzymatic synthesis of cocoa butter substitutes (CBS), as well as the operational stability and reusability of the resulting biocatalysts over several cycles. By considering these aspects, this study provides a comprehensive assessment of the potential of ESMC as a practical and sustainable support for industrial biocatalysis and highlights both its advantages and limitations.

2. Materials and Methods

2.1. Materials

The industrial eggshell waste was generously supplied by Elcon Nutritional Products Ltd. (Zlatar Bistrica, Croatia). It was processed with 5% (w/v) hydrochloric acid, 10% (w/v) acetic acid, and 15% (w/v) o-phosphoric acid. These acids were prepared from the following commercially available reagents: hydrochloric acid (37% w/v) purchased from Carlo Erba (Emmendingen, Germany), glacial acetic acid (99–100%) from LabExpert (Ljubljana, Slovenia), and o-phosphoric acid (85%) from Fisher Chemical (Shanghai, China). The obtained eggshell membranes were washed with acetone, which was acquired from LabExpert (Ljubljana, Slovenia). For carrier activation and flexible arm attachment, glutaraldehyde and polyethyleneimine were purchased from Sigma-Aldrich Chemicals (Saint Louis, MO, USA). For immobilization onto the prepared eggshell membranes, lipase from Pseudomonas fluorescens (Sigma-Aldrich, product number: 534730), which was purchased from Sigma-Aldrich Chemicals (Saint Louis, MO, USA), was used. Lipase activity was assessed using olive oil as a substrate, which was also obtained from Sigma-Aldrich Chemicals (Saint Louis, MO, USA). This study utilized various vegetable oils, including virgin olive oil (Trenton, Croatia), sunflower oil (Zvijezda, Croatia), vegetable oil (Omegol, Croatia), rapeseed oil (S-BUDGET, Austria), coconut oil (Encian, Croatia), and lard (PIK Vrbovec), all of which were purchased from the local market. Additionally, waste cooking oil was obtained from a nearby restaurant in Osijek, Croatia. A cocoa butter substitute was synthesized using palm oil (Virgin chef, Romania) and methyl stearate purchased from Sigma-Aldrich Chemicals (Saint Louis, MO, USA). All other chemicals used in this research were of analytical-grade purity.

2.2. Methods

2.2.1. Preparation of ESMC

Our research group has already published a number of papers describing the transformation processes of waste eggshells into various high-value products and carriers for the immobilization of enzymes based on the eggshell membrane [26,27,28]. According to the mentioned papers, preparation involved washing the eggshells with water before treatment with 5% hydrochloric acid (HCl), 10% acetic acid (HAc), or 15% o-phosphoric acid (H₃PO₄). After separation, washing, drying, and milling, three forms of ESMC were produced: ESMC-HCl, ESMC-HAc, and ESMC-H₃PO₄.

2.2.2. Lipase Immobilization

The immobilization of PFL onto ESMC was carried out via adsorption [29,30] and covalent binding [31,32]. In both approaches, the initial lipase activities used for immobilization ranged from 320 to 3200 U.

For adsorption, the carriers were incubated with the enzyme solution in 50 mM phosphate buffer (pH 7.5) at room temperature under constant rotation using a multi-rotator (Multi RS-60, BioSan, Riga, Latvia) set at 17 rpm. The immobilization was performed over a period of 1 to 6 h, maintaining a carrier-to-solution ratio of 1:20 (w/v), which was selected based on the water holding capacity of the carrier, as reported in our previous study [26].

Covalent binding was performed either directly or indirectly, depending on the activation procedure. For direct binding, the carriers were activated with glutaraldehyde at concentrations of 0.5% or 1% for 2 h. For indirect binding, glutaraldehyde-activated carriers were first modified with 0.2% polyethyleneimine for 2 h, followed by a second activation with 0.5% glutaraldehyde for an additional 2 h. Lipase was then covalently attached under the same conditions as for adsorption—room temperature, 17 rpm, and at a 1:20 carrier-to-solution ratio, over a period of 1 to 3 h.

After immobilization, all samples were filtered (Whatman No. 1 filter paper, pore size ~11 µm), washed to remove non-covalently bound enzyme, and lyophilized (ALPHA 2-4 LSC PLUS, Martin Christ Gefriertrocknungsanlagen GmgH, Germany, Osterode am Harz) for further use. Immobilized lipase activity was measured and expressed as U/g of wet carrier, while immobilization yield was calculated based on the difference between the applied and residual enzyme activities.

2.2.3. Characterization of Free and Immobilized PFL

To characterize the lipases, the optimal pH and temperature, stability (pH, temperature, organic solvents), substrate specificity, and reusability were determined. Therefore, lipase activity was measured according to Mustranta et al. [33], using a titrimetric assay with olive oil as the substrate. The activity assay was performed using 1 mL free lipase (1 mg/mL), 100 mg wet immobilized lipase, or 25 mg lyophilized immobilized lipase. pH optimum was determined to be within a pH range of 6 to 10 (phosphate buffer pH 6, 7, and 8, Tris-HCl buffer pH 8 and 9, and glycine-NaOH buffer pH 9 and 10), while the optimal temperature was determined to be within a temperature range of 30 to 70 °C. The pH stability was evaluated over a period of 6 h at the optimal temperature by dissolving the lipases in buffers of varying pH, spanning a pH range of 6 to 9. Thermal stability was assessed over 6 h at the optimal pH and at different temperatures (40, 50, 60, and 70 °C). The stability in organic solvents (30% methanol and 30% ethanol) was evaluated for three hours at the optimal temperature. For substrate specificity, the tested oils included standard olive oil, commercial virgin olive oil, sunflower oil, vegetable oil, rapeseed oil, coconut oil, fresh lard, and waste cooking oil. The properties of reusability were assessed by repeating the activity of the immobilized lipase in the hydrolysis of p-nitrophenyl palmitate (pNPP) ten consecutive times [34].

2.2.4. Synthesis of Cocoa Butter Substitute

The enzymatic synthesis of cocoa butter substitute was carried out following the procedure described by Saxena et al. [35]. The reaction was performed using palm oil and methyl stearate as lipid substrates, as well as the previously selected immobilized PFL as the biocatalyst. The reaction was conducted at 40 °C and 150 rpm for 24 h.

CBS were melted before analyses in a thermoblock (AccuBlock^TM, Labnet International, USA, Detroit) at 50 °C. Samples were weighted (50 mg) in a 10 mL volumetric flask, filled with n-hexane to the mark, and then filtered through a 0.22 µm filter. The determination of 1,3-dipalmitoyl-2-oleoyl-glycerol (POP), 1-palmitoyl-2-oleyl-3-stearyl-glycerol (POS), and 1,3-stearoyl-2-oleoyl-glycerol (SOS) was conducted on a Shimadzu GC—2010 Plus gas chromatograph (Shimadzu Corp., Kyoto, Japan) equipped with a flame ionization detector (FID) (Shimadzu Corp., Kyoto, Japan) and fitted with an Zebron 5HT Inferno^TM column (30 m × 0.25 mm × 0.1 µm). The injector and detector temperatures were set at 365 °C, the split ratio was 1:40, and the injection volume was 1 µL. The temperature program was as follows: initial temperature of 50 °C (0.5 min hold); 30 °C/min up to 350 °C; and 1 °C/min up to 365 °C (hold 5 min). The total time of analysis was 30.5 min. Reference standards of POP, POS, and SOS (Sigma Aldrich, Austria, Vienna) were used for the determination of retention times. An area normalization method was used for quantification, and the results were expressed as the average percentage of identified TAG (%). Analyses were performed in duplicate.

2.2.5. Statistical Analysis

Means and standard deviations were calculated using Microsoft^® Excel^® 2016 MSO. One-way analysis of variance (ANOVA) (p < 0.05) was performed using Statistica 14.0.0.15 software to assess statistical differences between lipases and the tested characterization parameters. Fisher’s post hoc test was performed (p < 0.05) to detect statistical differences between immobilized lipases in all determined parameters.

3. Results and Discussion

3.1. Adsorption

This article investigates the immobilization of PFL on ESMC treated with different acids using a range of initial enzyme activities from 320 to 3200 U. Early investigations identified that one hour is the most suitable time for effective immobilization, either by adsorption or covalent binding (Figures S1 and S2). Beyond this period, a decrease in enzymatic activity was observed, probably due to the formation of multilayers on the carrier surface, which cause steric hindrances between enzyme molecules and negatively affect their catalytic function by restricting substrate access. As shown in Figure 1, an increase in the initial lipase concentration leads to enhanced catalytic activity of the immobilized enzyme. However, this is accompanied by a decrease in immobilization yield, which ranged from 81.74% at the lowest concentrations to 50.57% at the highest concentrations. The decrease in immobilization yield at higher enzyme concentrations is due to the saturation of available binding sites on the carrier surface. When the enzyme concentration exceeds the carrier’s binding capacity, excess enzyme remains unbound in solution, leading to a lower calculated immobilization yield. Despite this, the total amount of immobilized enzyme and corresponding catalytic activity can still increase up to an optimal point before steric hindrance or multilayer formation negatively affects performance. The increased activity can be attributed to the greater availability of enzyme molecules for interaction with the carrier material. When different acid-modified ESMC carriers were evaluated, ESMC-HCl exhibited the highest activity at 3200 U, reaching 148.38 ± 3.48 U/g. A similar trend was observed for ESMC-HAc, where the maximum activity was also reached at 3200 U, yielding 98.26 ± 3.65 U/g. In contrast, ESMC-H₃PO₄ showed its highest activity at a slightly lower enzyme load of 2770 U, with a specific activity of 84.94 ± 0.73 U/g. These values are notably higher than those in the study by Jiang et al. [36], in which BCL immobilized on an eggshell membrane showed a maximum ester hydrolysis activity of only 12 U/g. Overall, the results suggest that higher loading of enzymes improves catalytic performance up to a certain point; however, excess enzymes may lead to limitations due to carrier surface saturation and lower binding efficiency. The careful tuning of immobilization time and enzyme concentration is therefore essential to achieve optimal performance of the biocatalyst.

3.2. Covalent Binding

In the case of covalent binding, activation with a lower concentration of glutaraldehyde proved more effective. The optimal immobilization time, as observed for adsorption, was one hour (Figure S2). In addition, other trends consistent with those observed with adsorption were noted, such as an increase in immobilized lipase activity and a decrease in immobilization yield with increasing lipase activity load (Figure 2). Accordingly, the highest activities of immobilized lipases upon covalent binding were achieved at the two highest applied lipase activity loads, 2770 and 3200 U. The immobilization yield ranged from 49.09% at higher applied lipase activity loads to 94.70% at lower applied lipase activity loads. As observed for immobilization by adsorption, the highest immobilized lipase activities were obtained on ESMC-HCl, reaching 166.05 ± 5.37 U/g and 221.16 ± 6.46 U/g for direct and indirect binding, respectively. In this case, covalent binding did not lead to a loss of activity of the immobilized enzyme; on the contrary, the insertion of a flexible polyethyleneimine spacer even led to an increase in activity, confirming previous literature results [37].

3.3. pH and Temperature Optimum

In our study, the immobilization of PFL resulted in a pronounced shift in both optimal pH (from 8 to 9) and optimal temperature (from 50 °C to 40 °C), indicating modifications in the enzyme’s microenvironment and structural flexibility upon attachment. Comparable phenomena have been reported in other systems due to changes in enzyme conformation caused by its binding to the support material, leading to strong attachment to the support and subsequent conformational changes in the enzyme [38,39,40]. In the case of PFL on supports containing eggshell membrane, the shift toward a more alkaline pH optimum (Figure 3) and lower optimal temperature (Figure 4) likely reflects combined effects of support charge, hydrophilicity, and structural rigidity, although the exact mechanisms require further elucidation.

3.4. pH and Temperature Stability

In contrast to many reports in the literature highlighting the enhanced pH stability of lipases following immobilization [41,42,43,44,45,46], our study did not observe such improvements. The free lipase retained the highest stability across the tested pH range, maintaining up to 98.68 ± 0.56% of its initial activity. Among immobilized preparations, the best results were obtained with directly covalently bound lipase on ESMC-HCl, which retained 86.01 ± 2.03% to 90.67 ± 2.10% of activity at pH 6 and 89.68 ± 0.47% at pH 9. Adsorption on ESMC-HCl yielded a slightly lower retention of 83.81 ± 0.58% at pH 6. Nevertheless, none of these values exceeded the stability of the free enzyme, suggesting that immobilization in this case did not confer additional resistance to pH-induced denaturation. Such enhanced behavior is typically attributed to the restricted conformational mobility of the enzyme and favorable microenvironmental effects induced by the support matrix. The absence of similar improvements in our system may be due to differences in the physicochemical properties of the support (e.g., charge, hydrophilicity), enzyme orientation, or the degree of activation and crosslinking. These results highlight that while immobilization can improve pH stability in certain systems, such benefits are not universally guaranteed and are highly dependent on the enzyme–support interaction and immobilization conditions.

In terms of thermal stability, a notable improvement following immobilization was observed only at elevated temperatures of 60 and 70 °C. At lower temperatures, the free lipase consistently exhibited higher stability, retaining up to 95.99 ± 0.73% of its initial activity. At 60 °C, the stability of the enzyme increased from 40.85% (in the free form) to as much as 66.83 ± 3.00% when adsorbed and 72.09 ± 1.12% when directly covalently immobilized. In contrast, indirect covalent immobilization resulted in negligible improvement, with stability remaining close to that of the free enzyme, around 40%. At the highest tested temperature of 70 °C, where the free enzyme completely lost its activity (0.26 ± 0.46%), the immobilized lipases retained up to 72.86 ± 1.03% of their initial activity. These findings clearly demonstrate that immobilization can significantly enhance the thermal resistance of lipases at elevated temperatures, particularly through direct covalent attachment. Improvement in thermal stability following lipase immobilization has been widely reported in the literature, particularly at elevated temperatures. In the present study, a significant increase in residual activity was observed only at 60 °C and 70 °C, where the immobilized lipases retained up to 72.86 ± 1.03% of their initial activity at 70 °C, compared to complete inactivation of the free enzyme (0.26 ± 0.46%).

Although immobilization generally enhances enzyme stability under harsh conditions by restricting conformational flexibility and protecting against denaturation, our results show that the free PFL exhibited superior stability at moderate temperatures and across the tested pH range. This may be due to immobilization-induced conformational constraints or suboptimal enzyme orientation on the support, which can reduce catalytic efficiency or stability under certain conditions. Nevertheless, immobilization provided clear benefits at elevated temperatures, significantly improving thermal resistance and enabling enzyme reuse. These findings underscore the importance of optimizing immobilization strategies to balance enhanced operational stability with preservation of enzyme activity.

3.5. Stability in Methanol and Ethanol

The immobilization of PFL lipase, either via adsorption or covalent attachment, demonstrated a positive impact on the enzyme’s stability in organic solvents, specifically methanol and ethanol. Organic solvents are known to disrupt the native conformation of enzymes, often leading to denaturation and loss of catalytic activity. However, immobilization can enhance structural rigidity and create a microenvironment that protects the enzyme from solvent-induced inactivation. In methanol, the free form of the enzyme retained only 18.29 ± 1.23% of its initial activity, indicating a substantial loss of function. In contrast, immobilized forms retained significantly higher residual activities: 47.40 ± 0.94% for indirect covalent binding, 74.85 ± 0.80% for direct covalent binding, and up to 84.86 ± 0.60% for adsorption-based immobilization. The superior performance of the adsorbed enzyme may be attributed to the preservation of its native structure due to milder immobilization conditions, as well as possible favorable interactions with the carrier surface. In ethanol, all enzyme forms exhibited better stability compared to methanol, consistent with the generally lower denaturing effect of ethanol on proteins [47,48]. In ethanol, all enzyme forms exhibited better stability compared to methanol, consistent with the generally lower denaturing effect of ethanol on proteins. The free enzyme retained 52.71 ± 0.97% of its initial activity. Immobilized forms again outperformed the free enzyme: indirectly covalently bound lipases retained 55.42 ± 0.92%, directly bound lipases retained up to 71.22 ± 0.60%, while adsorbed lipases maintained 70.86 ± 4.22% of their initial activity. Although slightly lower than in methanol, the stability of the adsorbed enzyme in ethanol remained high, suggesting that adsorption provides an effective stabilization strategy across different solvent systems. Overall, these findings confirm that immobilization not only improves enzyme reusability but also significantly enhances operational stability in challenging solvent environments, making it a valuable approach for biocatalysis in non-aqueous media. According to Zhang et al. [49], lipases are widely used in organic solvents due to advantages like improved substrate solubility and a favorable shift in reaction equilibrium, and Pseudomonas fluorescens JCM5963 is a promising strain that exhibits notable activity and stability in such media.

3.6. Substrate Specificity

Substrate specificity was evaluated for both free and immobilized PFL using a selection of vegetable oils and fats. Both forms of the enzyme exhibited comparable hydrolytic patterns across the tested substrates, with the highest activities consistently observed in coconut oil (Figure 5). This result suggests a pronounced affinity of PFL for medium-chain triglycerides, which dominate the composition of coconut oil. Coconut oil is rich in saturated fatty acids, primarily medium-chain triglycerides such as lauric acid (45–53%) and myristic acid (16–21%), which are more easily digested and metabolized than long-chain fatty acids found in oils like sunflower or olive. Despite lipases having low specificity for C12:0 and C14:0, their high activity on coconut oil is attributed to the easier hydrolysis of lauric acid and the favorable physical properties of the substrate, which improve enzyme accessibility [50,51].

3.7. Reusability

In the hydrolysis of pNPP, PFL proved to be highly effective over 10 reuse cycles, particularly when immobilized by adsorption, retaining up to 71.89 ± 0.24% of its initial activity (Figure 6a). This high retention can be attributed to the mild and reversible nature of adsorption, which typically preserves the native enzyme conformation and active site accessibility, thus limiting structural perturbations and maintaining catalytic efficiency across multiple cycles. In contrast, covalently bound lipases showed somewhat lower residual activities (Figure 6b,c), with the directly bound form retaining 61.61 ± 1.22% and the indirectly bound form showing the lowest value of 27.57 ± 0.92%. The covalent immobilization approach, while providing stronger enzyme attachment, may impose structural constraints or modify amino acid residues near the active site, thereby reducing enzyme flexibility and catalytic activity upon reuse. Additionally, harsher immobilization chemistries and possible steric hindrance can limit substrate access, which accelerates activity loss during repeated cycles.

These results were consistent with the literature and appeared even more favorable, considering that PFL immobilized on glyoxyl-octyl agarose beads maintained its activity over five cycles in the hydrolysis of triacetin in dioxane. In contrast, the same enzyme immobilized on nanoparticles retained 35–70% of its residual activity after seven cycles in the hydrolysis of p-nitrophenyl butyrate [52,53]. The superior performance of adsorption-based immobilization observed in our study, even over more reuse cycles, highlights the advantage of gentle binding methods in preserving enzyme function.

3.8. Selection of Lipases

Statistical analysis (ANOVA) confirmed significant differences (p < 0.05) in all tested factors, including lipase type, time, pH, temperature, organic solvent, and oil, affecting pH and temperature stability, solvent tolerance, and substrate specificity. To further explore differences among immobilized lipases, a post hoc Fisher test was applied, revealing statistically significant variations between PFL immobilized on different carriers for nearly all parameters (

Table 1. pH stability at 6 h of free and immobilized PFL. Results are shown as average relative enzyme activity ± standard deviation of three independent determinations.

pH Value	Average Relative Enzyme Activity [%]
	Free PFL
pH 6	84.56 ± 0.79
pH 7	95.94 ± 3.93
pH 8	98.68 ± 0.56
pH 9	95.99 ± 0.73
	PFL immobilized by adsorption
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
pH 6	83.81 ± 0.58 A	79.45 ± 2.05 B	78.28 ± 1.23 B
pH 7	71.22 ± 0.22 A	62.83 ± 1.01 B	62.53 ± 2.14 B
pH 8	66.03 ± 0.55 A	50.94 ± 0.90 B	58.18 ± 2.19 C
pH 9	66.28 ± 0.88 A	60.20 ± 0.78 B	56.30 ± 1.82 C
	PFL immobilized by direct covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
pH 6	86.01 ± 2.03 A	86.25 ± 1.35 A	90.67 ± 2.10 B
pH 7	71.78 ± 0.54 A	68.90 ± 0.32 B	69.26 ± 0.55 B
pH 8	68.16 ± 0.78 A	66.13 ± 1.29 B	63.70 ± 0.00 C
pH 9	89.68 ± 0.47 A	82.68 ± 1.43 B	83.08 ± 0.00 B
	PFL immobilized by indirect covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
pH 6	42.16 ± 2.98 A	41.33 ± 0.65 A	40.61 ± 1.22 A
pH 7	45.41 ± 0.52 A	41.83 ± 1.61 A	41.17 ± 5.75 A
pH 8	33.85 ± 0.37 A	30.35 ± 0.63 B	33.35 ± 0.16 A
pH 9	63.88 ± 0.54 A	52.17 ± 1.13 B	60.99 ± 1.11 C

Means with different superscripts within the raw data indicate significant differences; p < 0.05.

,

Table 2. Temperature stability at 6 h of free and immobilized PFL. Results are shown as average relative enzyme activity ± standard deviation of three independent determinations.

Temperature [°C]	Average Relative Enzyme Activity [%]
	Free PFL
40	87.90 ± 0.15
50	95.99 ± 0.73
60	40.85 ± 0.00
70	0.26 ± 0.46
	PFL immobilized by adsorption
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
40	66.28 ± 0.88 A	60.97 ± 0.78 B	56.30 ± 1.82 C
50	65.18 ± 1.88 A	60.98 ± 2.62 B	60.40 ± 1.91 B
60	66.83 ± 3.00 A	55.20 ± 1.67 B	57.16 ± 0.86 B
70	57.09 ± 1.14 A	30.27 ± 0.98 B	41.93 ± 0.83 C
	PFL immobilized by direct covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
40	89.68 ± 0.47 A	82.68 ± 1.43 B	83.08 ± 0.00 B
50	76.66 ± 3.55 A	75.21 ± 1.05 A	75.95 ± 0.34 A
60	87.66 ± 0.88 A	72.09 ± 1.12 B	80.75 ± 0.28 C
70	68.62 ± 1.88 A	67.37 ± 0.98 A	72.86 ± 1.03 B
	PFL immobilized by indirect covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
40	63.88 ± 0.54 A	52.17 ± 1.13 B	60.99 ± 1.11 C
50	59.69 ± 0.97 A	52.72 ± 0.98 B	56.51 ± 1.30 C
60	48.96 ± 1.38 A	42.45 ± 1.72 B	40.35 ± 0.41 B
70	32.35 ± 1.00 A	20.77 ± 1.43 B	21.66 ± 0.80 B

Means with different superscripts within the raw data indicate significant differences; p < 0.05.

,

Table 3. Stability of free and immobilized PFL at 3 h in methanol and ethanol. Results are shown as average relative enzyme activity ± standard deviation of three independent determinations.

Organic Solvent	Average Relative Enzyme Activity [%]
	Free PFL
methanol	18.29 ± 1.23
ethanol	52.71 ± 0.97
	PFL immobilized by adsorption
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
methanol	84.86 ± 0.60 A	69.27 ± 0.58 B	70.71 ± 1.58 B
ethanol	70.86 ± 4.22 A	67.00 ± 0.42 A,B	63.15 ± 1.57 B
	PFL immobilized by direct covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
methanol	72.65 ± 0.63 A	74.85 ± 0.80 B	71.13 ± 0.68 C
ethanol	70.84 ± 0.43 A	71.22 ± 0.60 A	64.34 ± 0.93 B
	PFL immobilized by indirect covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
methanol	47.40 ± 0.94 A	38.79 ± 0.68 B	45.99 ± 0.46 A
ethanol	55.42 ± 0.92 A	46.86 ± 1.87 B	52.33 ± 1.22 C

Means with different superscripts within the raw data indicate significant differences; p < 0.05.

and

Table 4. Activity of immobilized PFL. A-activity of lyophilized immobilized lipases at optimal conditions (pH 9, 40 °C). Results are shown as average relative enzyme activity ± standard deviation of three independent determinations.

	Lipases immobilized by adsorption
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
A [U/g]	659.96 ± 19.87 A	640.55 ± 21.36 A	608.37 ± 32.48 B
	Lipases immobilized by direct covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
A [U/g]	440.91 ± 4.39 A	445.23 ± 3.41 A	463.33 ± 9.37 B
	Lipases immobilized by indirect covalent binding
	ESMC-HCl-PFL	ESMC-HAc-PFL	ESMC-H3PO4-PFL
A [U/g]	392.78 ± 8.83 A	441.20 ± 3.40 B	441.09 ± 9.86 B

Means with different superscripts within the raw data indicate significant differences; p < 0.05.

), which aligns with ANOVA findings. Overall, the stability of immobilized PFL varied depending on the ESMC used, making it difficult to identify an optimal carrier. However, PFL adsorbed onto ESMC-HCl demonstrated superior activity across all tested oils and lard, as well as the best pH and temperature stability, especially at optimal conditions, as well as excellent reusability. Notably, Table 4 shows that this carrier yielded the highest activity, with significant differences compared to other ESMCs. Based on its overall performance, PFL immobilized on ESMC-HCl via adsorption was selected for further functionalization experiments, including the synthesis of CBS.

3.9. Cocoa Butter Substitute

Cocoa butter substitutes are widely used in the food industry due to the high cost and limited availability of natural cocoa butter. Their synthesis typically requires specific triacylglycerols, including POP, POS, and SOS, which impart the desired physical and sensory properties. In this study, the functionality of PFL immobilized on ESMC-HCl was evaluated in the enzymatic synthesis of CBS (Figure 7). The product obtained in the first reaction cycle was thick, viscous, and cream-colored, with a yield of 93.53 ± 0.16% of the desired triacylglycerols (POP, POS, and SOS) (Figure S3). Notably, the immobilized enzyme demonstrated excellent reusability; after five consecutive reaction cycles, the yield remained high at 90.69 ± 0.85%. These results indicate that ESMC-HCl-PFL is highly effective for synthesizing cocoa butter substitutes and retains its catalytic performance over multiple uses. Compared to previous work by Saxena et al. [35], who reported an 83.17% yield using a free Bacillus sp. RK-3 lipase at 37 °C with overnight incubation, our immobilized enzyme system demonstrates significantly improved efficiency. While reaction conditions differ slightly, both systems use the same substrates, allowing for meaningful comparison. The enhanced yield is likely due to improved catalytic stability and substrate accessibility conferred by enzyme immobilization. Furthermore, the immobilized lipase showed excellent reusability, maintaining a high CBS yield of 90.69 ± 0.85% after five consecutive reaction cycles, which highlights its potential for cost-effective industrial applications, offering both economic and practical advantages.

4. Conclusions

This study confirms the efficacy of Pseudomonas fluorescens lipase immobilized on an acid-treated eggshell membrane as a robust and reusable biocatalyst for the enzymatic production of a cocoa butter substitute (CBS). Among different immobilization strategies, adsorption on hydrochloric acid-treated carriers resulted in the best catalytic performance. The biocatalyst enabled the synthesis of CBS with 93.54 ± 0.16% of the targeted triacylglycerols (POP, POS, and SOS) and achieved high product quality and desirable physicochemical properties. In addition, the immobilized enzyme maintained its activity and yield over five synthesis cycles, demonstrating excellent operational stability and reusability.

Immobilization significantly shifted the optimal operating conditions of the enzyme and improved its stability at elevated temperatures and in the presence of organic solvents, overcoming the limitations normally associated with free lipases. The use of waste-derived carriers not only improved the performance of the enzyme but is also in line with sustainability principles, as it promotes the valorization of resources and reduces dependence on virgin materials.

These results provide strong evidence for the practical applicability of immobilized lipases in lipid engineering and support the further optimization and scale-up of CBS production processes. Overall, the integration of enzyme immobilization and waste recycling represents a promising strategy for developing stable, ethical, and environmentally friendly chocolate production systems.