Novel NTA-Ni2+ Agarose-Based Microspheres: Structural Features and Chromatographic Capacity

TL;DR

This study develops new agarose-based microspheres for protein purification, showing high efficiency and structural stability.

Contribution

A novel one-step method for preparing NTA-Ni2+ agarose-based microspheres with optimized structural and chromatographic properties.

Findings

Optimal preparation conditions achieved a span value of 0.50684 for uniform microsphere size.

NTA-Ni2+ ABM showed high binding capacity for His-tagged proteins (15.2 ± 0.8 mg/mL).

SEM, AFM, DSC, and FTIR confirmed structural stability and uniform cross-linking network.

Abstract

The design and optimization of immobilized metal affinity chromatography (IMAC) media are crucial to enhancing the purification efficiency of recombinant proteins. In this study, the agarose-based microspheres are prepared by using a three-factorial Box–Behnken design followed by NTA-Ni2+ agarose-based microspheres (ABM) preparation by the “one-step” crosslinking of epichlorohydrin (ECH)–nitrilotriacetic acid (NTA) to efficiently couple the NTA ligand to the surface of the matrix. After preparation, various sophisticated techniques, including SEM, AFM, DSC, FTIR, and SDS-PAGE, were used to analyze the morphological structure, thermal stability, and chemical composition of NTA-Ni2+ ABM. The optimal conditions are identified as an emulsifier PP concentration of 8.12 wt%, a stirring speed of 1624.46 rpm, and an oil-phase temperature of 53.86 °C, giving a span value (Y) of 0.50684. SEM, AFM, DSC, and FTIR results showed that the fabricated NTA-Ni2+ ABM were structurally stable and had a uniform cross-linking network for up to 8 h of coupling reaction time. The performance results showed that the beads had a high binding capacity for His-tagged proteins (15.2 ± 0.8 mg/mL), and SDS-PAGE results demonstrated the efficient purification ability for target proteins. These findings provide the theoretical basis and a practical solution for the rational design and application of IMAC medium.

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1. Introduction

The rapid development of recombinant protein technology has increased the demand for efficient, specific downstream purification processes [1,2,3]. Among various purification techniques, Immobilized Metal Affinity Chromatography (IMAC) is a widely applied method for the selective purification of recombinant proteins bearing histidine tags (His-tag) due to its simple operation, high loading capacity, and versatility [4]. Selective adsorption and separation are achieved via the coordination between transition metal ions (e.g., Ni^2+^, Co^2+^, and Cu^2+^) immobilized on solid-phase matrices and the imidazole moieties of histidine residues in target proteins [5,6]. Among transition metal ions, Ni^2+^ is widely used in the laboratory and in industrial-scale production due to its strong affinity for His-tag, good stability, and relatively low cost [7,8,9].

In IMAC systems, the characteristics of the solid-phase matrix directly affect the purification efficiency [10,11]. An effective matrix should have good biocompatibility, a suitable pore-size distribution, sufficient mechanical strength, and provide adequate functional groups for ligand attachment [12,13,14]. Agarose gels are commonly used as ideal matrices for IMAC due to their hydrophilicity, favorable pore structure, and easy modification [15]. However, natural agarose gels have low mechanical strength, which can restrict flow rates and increase column pressure during chromatography [16]. Chemical cross-linking is a practical approach to improve the physicochemical stability of the agarose matrix [17]. Epichlorohydrin (ECH), a bifunctional cross-linking agent, can effectively enhance the mechanical strength and adjust the pore structure of the agarose matrix [18]. The degree of cross-linking not only affects mechanical stability but also swelling behavior, mass-transfer efficiency, and the final protein-binding capacity [19].

The selection of chelating ligands is a crucial element for the IMAC system, as it controls the stability of metal immobilization and affects the purity of protein separation [20,21]. Among various chelating ligands, nitrilotriacetic acid (NTA) can form a stable bivalent chelating complex with Ni^2+^, leaving two sites available for specific binding to His-tagged proteins [22]. Compared with conventional tridentate ligands (e.g., iminodiacetic acid, IDA), the NTA-Ni^2+^ complex effectively reduces the detachment of metal ions during the process, thereby enhancing product safety and purity [23]. Previous studies have shown that the metal ion shedding rate of NTA-Ni^2+^ media is significantly lower than that of IDA-Ni^2+^ media, making them especially suitable for applications requiring high-purity proteins [24].

Although IMAC is a well-established technology, current commercial IMAC media have some limitations. High costs and limited flexibility restrict the customization of commercial media, affecting usage and deep optimization for specific purification tasks [25]. In addition, the preparation process of commercial media is usually not publicly available, making performance optimization and mechanism analysis challenging [26]. To address the complexities of high-yield recombinant protein expression systems, purification processes must achieve a balance among efficiency, specificity, and cost [27]. Therefore, the development of a controllable, reproducible, and cost-effective autonomous preparation method, along with the systematic study of its “preparation-structure-property” correlation, has clear scientific value and application potential.

In recent years, statistical design of experiments (DOE) techniques have become increasingly crucial for bioprocess optimization. Response surface methodology (RSM), a powerful tool for experimental design, can effectively investigate complex interactions among independent variables and accurately determine optimal process parameters by building mathematical models [28]. RSM has been successfully applied to improve the preparation of a variety of biomaterials, including optimizing the cross-linking process for microspheres and improving protein purification conditions. In chromatography, RSM also helps researchers to maximize the length of the NaCl gradient, flow rate, and sample uptake in heparin affinity chromatography to maximize the yield, purity, and productivity of mono-PEGylated lysozyme [29].

Although several studies have reported the effects of different ligands on IMAC media performance [10,20,30], and also how cross-linking conditions affect the agarose matrix properties. Still, limited data is available on how the degree of epichlorohydrin (ECH) cross-linking correlates with the performance of NTA-Ni^2+^ media. In particular, detailed investigations into the constitutive relationships between the level of cross-linking, the media’s microstructure and surface properties, and protein purification performance are still rare. In addition, most existing studies focus on evaluating a single variable and lack a comprehensive evaluation from material preparation to final application. Based on the above research background and technical challenges, the current study aims to independently prepare high-performance NTA-Ni^2+^ agarose-based microspheres (ABM) via epichlorohydrin (ECH) cross-linking and NTA coupling. Additionally, we systematically investigate the intrinsic correlation between the preparation process, structural properties, and protein purification performance. Finally, the binding capacity of NTA-Ni^2+^ ABM to His-tagged proteins was evaluated to verify the practical application value.

Compared with conventional agarose activation and ligand-coupling procedures, the novelty of this study lies in its preparation strategy and process optimization. RSM was employed to systematically optimize ABM preparation conditions, enabling precise control of particle size distribution. In addition, a one-step ECH-NTA coupling strategy was adopted to simplify functionalization. The relationships among preparation parameters, particle structure, and chromatographic performance were further investigated to guide the rational design of polysaccharide-based IMAC media.

2. Materials and Methods

2.1. Materials and Reagents

Agarose was purchased from Yeasen (Yeasen Biotechnology, Shanghai, China; CAS: 9012-36-6). Polyglyceryl-6 pentastearate (PP) was purchased from Lullaby (Wuhan Lullaby Pharmaceutical Chemical, Wuhan, China; CAS: 99734-30-2). Liquid paraffin (LP) (CAS: 8042-47-5) and epichlorohydrin (ECH) (CAS: 67843-74-7) were purchased from Macklin (Macklin Biochemical, Shanghai, China). Petroleum ether (PE) (CAS: 101316-46-5) and NTA-3Na (CAS: 5064-31-3) were purchased from Sigma-Aldrich (Sigma-Aldrich Trading, Shanghai, China). The BCA Protein Assay Kit (Cat. No. P0010S) and SDS-PAGE gel preparation kit (Cat. No. P0697S) were purchased from Beyotime (Beyotime Biotechnology, Shanghai, China). The other chemicals and reagents used in this study were commercially sourced and of analytical grade.

2.2. Preparation of ABM

2.2.1. Selection of ABM Preparation Conditions

ABM were prepared following the method of Li [31] with minor modifications. The oil phase was obtained by mixing LP and PE at a 7:5 volume ratio and adding 8 wt% PP, followed by stirring at 800 rpm and heating to 53 °C to ensure complete dissolution and uniform blending. The water phase was prepared by dissolving 3.5 wt% agarose and 0.5 wt% NaCl in deionised water. Subsequently, the water and oil phases were combined at a 1:15 volume ratio. A peristaltic pump was used to disperse the water phase into the oil phase, and the emulsion was stirred at 1500 rpm to generate uniform droplets. As the system cooled, the stirring speed was gradually reduced, allowing the droplets to solidify into ABM. The resulting beads were stored in 20% ethanol at 4 °C.

Three factors were investigated in this study: PP concentration (A), stirring speed (B), and oil-phase temperature (C). Particle size uniformity (Span, Y) was used as the response variable. Pre-experiments indicated that the highest Span values were obtained at 8 wt% PP, 1500 rpm stirring speed, and 50 °C oil-phase temperature.

2.2.2. Response Surface Methodology (RSM)

A three-factor, three-level Box–Behnken design (BBD) was adopted for optimizing the preparation conditions of ABM, as detailed in Section 2.2.1, with the levels of each factor presented in Table 1 [32].

2.2.3. Span Value

Span is an important parameter for characterizing particle size distributions and describing their range. Span is calculated by Equation (1) [33]:

$$[eqn]$$

where D_90_, D_10_, and D_50_ represent the particle sizes that reach 90%, 10% and 50% in the cumulative distribution, respectively. Smaller Span values indicate a more homogeneous particle size and better size consistency [34]. Particle size was measured using ImageJ software (ImageJ 2.0, National Institutes of Health, Bethesda, MD, USA).

2.3. Preparation of NTA-Ni2+ ABM

2.3.1. ECH Crosslinking

Agarose beads were crosslinked with epichlorohydrin (ECH) under alkaline conditions to enhance their mechanical and chemical stability, following the method of Matsumoto et al., with slight modifications [35]. ABM with a dry mass of 0.46 g were washed thoroughly with deionized water, then placed in 100 mL NaOH (1 M), and stirred continuously for 30 min at 0–4 °C with the pH maintained between 10 and 12. At the end of stirring, 0.75 mL of ECH (98%, ρ ≈ 1.18 g/mL) corresponding to 0.867 g (9.38 mmol) of effective reagent was slowly added dropwise to the suspension over a period of 5–10 min. The cross-linking reaction was continued for 1 h at 0–4 °C.

2.3.2. NTA Coupling

The NTA coupling reaction was carried out under alkaline conditions, following the approach reported by Hochuli et al., with appropriate adaptations for agarose-based matrices [36]. After the ECH cross-linking reaction lasted for 1 h, NTA-3Na (30.38 g, 87.5 mmol) solution was slowly added to the reaction system, and the reaction was continued at 0–4 °C. Samples were taken at 4 h, 8 h, 10 h, and 12 h for subsequent structural characterization. The reaction was terminated by adjusting the pH to 6–7 with HCl (1 M), after which the samples were sequentially washed with deionized water and Tris-HCl buffer (pH 7–8, 0.3 M NaCl) to obtain NTA-coupled ABM. The NTA-coupled ABM were preserved in 20% ethanol at 4 °C.

2.3.3. Coordination and Immobilization of Ni2+

The NTA-coupled ABM with a dry mass of 20 g were resuspended in 100 mL Tris-HCl buffer (pH 7–8, 0.3 M NaCl), and a 1 M NiSO_4_ solution at a volume of 50 mL was introduced, followed by incubation at room temperature with continuous stirring for 2 h [4,37]. After completion of the reaction, the ABM were washed 3–5 times with Tris-HCl buffer (pH 7–8, 0.3 M NaCl) to remove all unbound free nickel ions. The final NTA-Ni^2+^ ABM were placed in PBS and stored at 4 °C.

A portion of the NTA-Ni^2+^ ABM were fixed in a 2.5% glutaraldehyde solution at 4 °C for 2–4 h. The beads were subsequently rinsed three times with phosphate-buffered saline (PBS) to remove residual glutaraldehyde. Subsequently, dehydration was performed using a series of gradient solutions of anhydrous ethanol (30%, 50%, 70%, 90%, 100%), with each concentration immersed for 15–20 min. After dehydration, the NTA-Ni^2+^ ABM were frozen at −80 °C for 12 h and then lyophilized for 48 h using a lyophilizer (DGJ-56L12NP, So Euro-Sino, Beijing, China). This protocol was employed to optimally preserve the hydrated surface ultrastructure and structural integrity of the NTA-Ni^2+^ ABM for subsequent characterization.

2.4. Structural Characterization and Application Properties

2.4.1. Electron Microscope (SEM)

The microstructure of ABM was observed under different cross-linking times using a scanning electron microscope (Zeiss Gemini Sigma 300 VP SEM, Carl Zeiss AG, Jena, Thuringia, Germany), following the method of Li et al., with minor modifications [31]. Before imaging, the samples were fixed to sample disks with conductive adhesive, and gold was sprayed onto the sample surface. The magnification was set at 100× and 200×.

2.4.2. Atomic Force Microscope (AFM)

The surface morphology and mechanical properties of ABM were characterized using an atomic force microscope (Bruker Dimension Icon AFM, Bruker Corporation, Billerica, MA, USA), following the approach described by Pernodet et al. [38]. AFM measurements were performed using freeze-dried ABM samples. Two-dimensional and three-dimensional topographies of the sample surface were obtained in tap mode; force curves were collected in peak force quantitative nanomechanics mode; and DMT modulus values of the samples were obtained after analysis. The obtained DMT modulus values represent the apparent surface modulus of the dried materials rather than the intrinsic bulk mechanical properties of hydrated gels.

2.4.3. Differential Scanning Calorimetry (DSC)

The thermal stability of ABM was evaluated by using a differential scanning calorimeter (DSC) (Discovery DSC 2500, TA Instruments, New Castle, DE, USA) according to the method reported by Aslam et al., with slight modifications [39]. The measurements were performed over a temperature range of 30–200 °C at a heating rate of 10 °C/min, and the heating was carried out under a nitrogen atmosphere (purity ≥ 99.9%).

2.4.4. Fourier Transform Infrared Spectroscopy (FTIR)

A Fourier transform infrared spectrometer (FTIR) (Nicolet iS20, Thermo Fisher Scientific, Waltham, MA, USA) was employed to analyze the secondary structure of ABM, following the method of Hu et al. with slight modifications [40]. The infrared spectral signal was accumulated from 32 scans of the sample using an attenuated total reflection (ATR) accessory with air as the initial background. The test frequency range was 4000 cm^−1^ to 400 cm^−1^, with a resolution of 2 cm^−1^. Finally, the data were processed using OMNIC software (OMNIC 8.2, Thermo Fisher Scientific, Waltham, MA, USA).

2.4.5. Contact Angle

According to the method of Li et al. [31], the contact angle between the aqueous and oil phases was measured using a hanging drop method with a video optical contact angle meter (Dataphysics OCA20, DataPhysics Instruments, Filderstadt, Baden-Württemberg, Germany). Five measurements were taken for each sample using ImageJ (ImageJ 2.0, National Institutes of Health, Bethesda, MD, USA), and the average was used as the final contact angle.

2.4.6. Swelling Degree (SD)

To assess the physical stability of the prepared ABM in applications, its swelling behavior was characterized. Referring to the method of Popov S et al. [41], with minor adjustments, the dried ABM were submerged in PBS buffer (pH 7.4) and incubated at 25 °C with constant-temperature oscillation. Photographs were taken using an optical microscope (Olympus BX51, Olympus Corporation, Tokyo, Japan), and the diameters of the ABM were measured using ImageJ software (ImageJ 2.0, National Institutes of Health, Bethesda, MD, USA). The projected areas of the ABM were calculated. The projected area was used instead of direct diameter measurement to reduce directional bias and to improve accuracy in cases where microspheres exhibited slight deviations from perfect circularity in microscopy images. SD was calculated using Equation (2):

$$[eqn]$$

where S_1_ is the surface area of the ABM after soaking in PBS for a period of time, and S_0_ is the initial surface area.

2.5. Functional Characterization of Protein Purification

2.5.1. Preparation of Protein Mixture Solution

To evaluate the purification performance of NTA-Ni^2+^ ABM in a complex protein system, a protein mixture solution was prepared. Lyophilized His-tagged protein (22 kDa) was reconstituted with phosphate-buffered saline (PBS). PBS was added to the vial, vortexed thoroughly to ensure complete dissolution, and allowed to stand for 5 min. The resulting solution was transferred into a 50 mL centrifuge tube. The original vial was rinsed several times with a small volume of PBS, and the rinsate was combined with the protein solution. Finally, the volume was adjusted to 50 mL with PBS. The final concentration of the protein in the obtained solution is 0.14 mg/mL.

An equal volume (50 mL) of whole-protein bacterial lysate, which was obtained by lysing and centrifuging blank bacterial cells, was mixed with the 50 mL His-tagged protein solution. The mixture was gently stirred to achieve homogeneity, yielding a 100 mL protein mixture for subsequent purification experiments.

2.5.2. Isolation and Purification of His-Tagged Proteins by NTA-Ni2+ ABM

A defined volume of NTA–Ni^2+^ ABM was packed into a chromatography column (EZ50, Xinhu Experimental Equipment, Shanghai, China) and equilibrated with 5 column volumes (CV) of PBS. The experiments were performed using a column with an inner diameter of 25 mm, a bed height of 5 cm, and a total column volume of 24.5 mL. The prepared crude protein mixture was then loaded onto the equilibrated column at a flow rate of 1 mL/min to facilitate sufficient binding of the His-tagged protein. The flow-through fraction was collected, and UV absorbance at 280 nm was recorded to monitor protein elution profiles, including flow-through, wash, and elution peaks.

The column was rinsed with 10 CV of wash buffer (20 mM Tris-HCl, 300 mM NaCl, 40 mM imidazole, pH 7.4) to remove nonspecifically adsorbed proteins, and the corresponding wash fractions were collected. The specifically bound His-tagged target protein was subsequently eluted using 10 CV of elution buffer (20 mM Tris-HCl, 300 mM NaCl, 250 mM imidazole, pH 7.4), and the eluted fractions were collected.

The protein concentrations in all collected fractions were determined using a bicinchoninic acid (BCA) protein assay kit. The overall purification procedure was conducted following the protocol described by Bornhorst et al., with slight modifications [5].

The binding capacity was calculated from the volume of wet-packed beads (mg/mL), a commonly used metric for evaluating chromatographic media.

The protein loading of NTA-Ni^2+^ ABM was calculated according to Equation (3) [42]:

$$[eqn]$$

where C_1_,…,n is the concentration of eluted protein, V_1_,…,n is the volume of eluted protein, and V_bed_ is the volume of wet-packed beads.

2.5.3. SDS-PAGE

A rapid preparation kit was used to cast 12% SDS–PAGE gels. Each gel well was loaded with 20 μL of protein sample and 5 μL of molecular weight marker (8–195 kDa; Servicebio Technology, Wuhan, China; Cat. No. G2087). Protein concentrations were determined using a BCA Protein Assay Kit before electrophoresis. For SDS-PAGE analysis, a fixed sample volume (20 μL) was loaded per lane. Band intensities were analyzed using ImageJ software for semi-quantitative evaluation and estimation of protein loading capacity. Electrophoresis was initiated at 80 V, and the voltage was increased to 120 V after the samples had migrated into the resolving gel [32]. After electrophoresis, the samples were stained with Caulmers Brilliant Blue (Sigma-Aldrich Trading, Shanghai, China; CAS: 6104-59-2) for 30 min. After the staining, the sample was eluted with elution solution (ethanol: glacial acetic acid: distilled water = 250 mL: 80 mL: 670 mL) for 30 min until the electrophoretic bands were clear.

SDS-PAGE images were analyzed using ImageJ software. The purity of His-tagged proteins in the eluate was evaluated by the ratio of the signal intensity of His-tagged protein bands to the total protein signal intensity in their lanes.

2.6. Statistical Analysis

Data are presented as the mean ± standard deviation (SD) from three independent experiments. Statistical analyses were conducted using SPSS software (version 17.0, IBM Corporation, Armonk, NY, USA). Differences among multiple groups were evaluated by one-way analysis of variance (ANOVA), followed by a Tukey post hoc test. Differences were considered significant at p < 0.05. RSM analyses were performed using Design Expert (Version 13, Stat-Ease, Minneapolis, MN, USA) with Origin (Version 2025, OriginLab Corporation, Northampton, MA, USA) and Graphpad Prism (Version 10.1, GraphPad Software, San Diego, CA, USA) for plotting.

3. Results and Discussion

3.1. Optimal Preparation Conditions of ABM

The preparation conditions of ABM were optimized in a simulated system containing agarose, emulsifier, and oil phase based on response surface methodology (RSM). A Box–Behnken design (BBD) was used to prepare the reaction system with emulsifier concentration, magnetic stirrer speed, and oil phase temperature to obtain the response span values. The RSM experimental results are presented in Table 2.

The dependent responses exhibited span value (2.19) against the independent variables at run 1. The fitted second-order polynomial equation for span value is as follows:

$$[eqn]$$

The ANOVA results are presented in Table 2. For a better and well-fitted model, the model should be significant, with the lack-of-fit term remaining statistically non-significant. In this study, the BBD model is significant, while the lack-of-fit term is not significant (Table 3). Similarly, the regression coefficient R^2^ = 0.9761 is also close to 1, exhibiting higher R^2^ values, which reflects that our model fits the second-order polynomial equation well. Additional parameters, including the coefficient of variation (CV), F-value, adjusted R2, and adequate precision, further validate the significance of the model. In the present study, F-values (5.19), Adj R^2^ (0.94), adequate precision (17.53), and CV (10.09) exhibited the fitness of the RSM model. This observation is in accordance with Javed et al. [43], as they determined the parameters (including R^2^, CV, and adeq. Precision) during the recovery of the total polyphenolic compounds in the sargassum fusiforme using RSM and ANN models.

Contour plots were constructed to examine in more detail how the independent variables affect the outcome response, as shown in Figure 1. As shown in Table 2, the linear effects of factors A, B, and C, along with the quadratic effects of A^2^, B^2^, and C^2^, were significant in the model. The optimal conditions for the preparation of ABM were an emulsifier PP concentration of 8.12 wt%, a magnetic stirrer speed of 1624.46 rpm, and an oil phase temperature of 53.86 °C. To validate the experiment, we adjusted the parameters to 8.12 wt%, 1620 rpm, and 53.9 °C, respectively. It is worth noting that the span value predicted by the model is 0.5056, which is very close to the actual value of 0.507, verifying the reliability of the model. The reported value was rounded to three significant figures to reflect the practical accuracy of image analysis. Particle size distribution influences chromatographic performance by affecting mass transfer efficiency and protein diffusion within the porous matrix. A narrower particle size distribution provides more uniform packing, shorter diffusion paths, and a larger accessible surface area, which facilitates protein transport and interaction with immobilized affinity sites, thereby improving binding performance [44,45]. In addition, smaller, more uniformly distributed particles can reduce band broadening and enhance mass transfer kinetics, thereby improving separation efficiency [46].

3.2. Structural Characterization and Application Properties of ABM

3.2.1. The Surface Feature Analysis of ABM by Scanning Electron Microscopy

After optimizing the ABM preparation conditions, ABM were synthesized and further characterized using scanning electron microscopy (SEM), as shown in Figure 2. At 100× magnification, all samples maintained a precise spherical shape, indicating that the one-step ECH–NTA coupling conditions, particularly the pH and overall reaction environment, avoided the deformation of the bead structure. However, as the coupling time gradually increased, notable changes in surface morphology were detected. At 0 h, ABM exhibited a relatively smooth and compact surface. After 4 h of reaction, the surface became noticeably rougher, with small protrusions and micropores, which might be due to the combined effects of ECH crosslinking and the initial grafting of NTA moieties.

At 8 h and 10 h, the degree of roughening and the surface porous structure of the ABM became more pronounced, forming a clearer, more open three-dimensional network. This transition from compact morphology to loose and more porous structure is consistent with previously reported structural remodeling caused by crosslinking and ligand coupling [18]. The increase in surface area and porous structure provides more active sites for the full exposure of NTA ligands and subsequent chelation with Ni^2+^, as well as creating more favorable mass transfer channels for the diffusion and binding of protein molecules [47].

It is noteworthy that after 12 h, the surface structure of the ABM, rather than improving, showed partial contraction or collapse of the structural network under high magnification (Figure 2). This may be due to excessive ECH crosslinking caused by the long reaction time, leading to contraction of the gel network and a reduction in pore size. Similar phenomena of excessive cross-linking leading to structural tightening have been reported in other gel-based material studies [48]. Overall, SEM analysis revealed that the ECH–NTA coupling process significantly affects the surface structure of ABM. The most favorable porous morphology was obtained at 8–10 h of reaction time. A reaction time that is too short may result in insufficient modification, whereas a long reaction time may compromise the porous structure due to over-crosslinking. Therefore, controlling the coupling time at 8–10 h is critical for achieving optimal microstructure and performance.

3.2.2. The Surface Morphology and Roughness Analysis of ABM by Atomic Force Microscopy

Atomic force microscopy (AFM) is an advanced technique widely used to investigate the mechanical properties of tissues, cells, fibrous materials, and biomolecules. As depicted in Figure 3A, as the coupling time gradually increased (from 0 h to 12 h), the initial smooth surface of ABM started transforming into a rough surface with prominent ridges and uneven features, indicating ongoing surface modification during the reaction. The 3D AFM images also revealed increasing spatial heterogeneity, which may result from diffusion limitations within the gel matrix and variations in local reaction rates [49]. The DMT modulus, which responds to the elastic deformation resistance of the material, also changed during the coupling process [50]. The results showed that the surface stiffness was enhanced during the early and middle stages of the reaction, reaching its highest value (40 GPa) at 8–10 h. The relatively high modulus values observed here can be attributed to multiple factors. First, dehydration and structural densification of the polymer network during freeze-drying significantly increase the material’s stiffness. In addition, AFM nanoindentation probes local surface mechanical properties at the nanoscale, potentially yielding higher apparent modulus values. Furthermore, the increased stiffness may also be associated with a denser crosslinking network and an elevated density of NTA functional groups on the bead surface, which enhances the structural rigidity of the polymer matrix [51,52]. As the coupling time increased (10–12 h), the DMT modulus decreased to 21.2 GPa, suggesting reduced elasticity and increased susceptibility to deformation. This trend is consistent with the structural contraction or partial collapse observed in the SEM analysis. Figure 3B exhibited the quantitative analysis of surface roughness, including the root mean square roughness (Rq) and average roughness (Ra). Both Rq and Ra significantly increased with longer coupling times, further confirming the SEM outcomes that the ECH-NTA coupling reaction induces significant changes in the surface structure of the ABM.

3.2.3. The Thermal Properties and Chemical Structure Analysis of ABM by Differential Scanning Calorimetry and Fourier Transform Infrared Spectroscopy

The DSC is widely used to analyze the thermal behavior of different materials by measuring the heat absorbed or released during controlled heating, as well as by the physical characteristics of the substances. This technique has a unique capability to measure the enthalpy changes associated with specific thermal processes directly. Figure 4A revealed the significant effect of chemical modification on the thermal stability of the materials. At 0 h, experimental samples exhibited a central thermal peak at around 105 °C, which is likely associated with the thermal decomposition of the agarose gel network, reflecting the inherent thermal stability of the agarose matrix [53]. As the ECH-NTA reaction progressed, this peak gradually shifted to higher temperatures, indicating improved thermal stability. This change in thermal stability is likely due to the formation of new chemical bonds between NTA and the agarose matrix [54]. However, the peaks at 10–12 h showed a slight decrease in temperature relative to the peak at 8 h, suggesting that the system may have reached a saturation point or equilibrium. Further NTA binding did not significantly alter the thermal properties.

FTIR analyses were used to identify organic, polymeric, and, in a few cases, inorganic materials, and Figure 4B showed the structural changes in the ABM during the experiment. All samples exhibited a pronounced absorption peak around 1040 cm^−1^, which is attributed to the C-O-C stretching vibration of the agarose backbone [55]. The absorption peak at about 3340 cm^−1^ corresponds to the O-H stretching vibration of hydroxyl groups arising from hydrogen-oxygen bonding within the agarose matrix [56]. Additionally, the absorption bands at approximately 1600 cm^−1^, 1400 cm^−1^, and 2900 cm^−1^ corresponded to stretching vibrations of the carbonyl group (C=O), the carbon-oxygen bond (C-O), and the carbon-hydrogen bond (C-H), respectively [57]. As the coupling reaction progressed, the intensities of existing peaks changed, along with the formation of new peaks. In particular, the peaks within 1400–1600 cm^−1^, attributed to the asymmetric and symmetric stretching vibrations of NTA carboxylate groups (-COO^−^), became more noticeable, confirming successful attachment of NTA to the agarose matrix [58]. The absorption band at 1640 cm^−1^ showed variations in both shape and intensity, which might be due to the bending vibration of the hydroxide bond (O-H) [59]. This effect may be attributed to the formation of a crosslinked network, which alters the gel’s water-holding capacity, thereby indirectly demonstrating the modifying impact of ECH crosslinking on the gel’s microenvironment. The FTIR results were also consistent with the DSC outcomes, indicating that the NTA coupling process led to significant chemical and structural changes in the ABM. Taken together, the surface morphology and physicochemical properties of the ABM were better when the ECH-NTA coupling time was set to 8 h for further experiments, and Ni^2+^ loading was performed following the procedure outlined in Section 2.3.3 to prepare the NTA-Ni^2+^ ABM.

3.2.4. Dynamic Contact Angle Variation and Quantitative Analysis of Water–Oil Interface

Dynamic contact angle analysis determines how two immiscible liquids interact with a solid surface, revealing the surface roughness and chemical heterogeneity. As shown in Figure 5A,B, the dynamic contact angle of water droplets at the oil-phase interface decreases gradually with time from 128.7° to 78.6°, indicating improvement in wettability. The high contact angle at the initial stage reflects the limited wetting ability of water relative to the oil-phase interface. As time gradually progresses, the marked reduction in the contact angle reflects a continuous decrease in interfacial tension. This decreasing contact angle trend is consistent with classical wetting theory: a decrease in interfacial tension shifts the contact line forward, bringing the system closer to a fully or partially wetted equilibrium state and thus improving the morphological stability of water droplets in the oil phase [60].

3.2.5. Swelling Degree of NTA-Ni2+ ABM

Swelling degree analysis helps investigate a material’s ability to absorb liquid by increasing in volume, thereby determining its structural stability. Figure 5C–E illustrated the swelling behavior of NTA-Ni^2+^ ABM in PBS. As shown in Figure 5C, the ABM volume gradually increased over time. The expansion in volume, quantified by the surface area (Figure 5D), confirms that the swelling follows a diffusion-controlled pattern. As depicted in Figure 5E, the degree of swelling continues to increase rapidly up to 12 h. After 16 h, the swelling rate eventually reaches a steady state, suggesting that water diffuses into the gel network until a balance is reached between osmotic pressure and the elastic restoring force of the matrix.

This phenomenon is consistent with classical polymer-gel swelling theory, in which hydrophilic groups in the gel network absorb water, relax the polymer chains, and allow the network to expand. The NTA-Ni^2+^ coordination sites act as effective cross-linking points, helping to stabilize the structure and preventing excessive swelling [61]. Additionally, metal–ligand bonding is more stable than conventional physical cross-linking. It helps maintain the overall structural integrity of the ABM, which is also consistent with existing reports on the enhanced stability of metal–ligand structures in gel materials [62]. Because metal–ligand coordination is generally more robust than simple physical cross-links, the overall integrity of the beads is well maintained. The equilibrium swelling degree therefore reflects the combined effects of network density and hydrophilicity. In this system, the gradual leveling of the swelling curve suggests that NTA-Ni^2+^ modification preserves the three-dimensional agarose network while forming a uniform cross-linked structure with good stability and diffusion properties. Such controlled swelling is vital for subsequent protein enrichment, immobilization, and chromatographic applications, as it increases the surface area and mass-transfer efficiency while maintaining the functional integrity of the coordination structure.

3.3. Evaluation of Functional Properties of NTA-Ni2+ ABM for Protein Purification

3.3.1. Procedure for Purification of His-Tagged Proteins by NTA-Ni2+ ABM

As shown in Figure 6A, His-tagged protein purification was carried out using the prepared NTA–Ni^2+^ ABM complex. At first, a pre-prepared protein mixture solution (Section 2.5.1) was passed through a chromatography column packed with the NTA-Ni^2+^ beads. The His-tagged proteins selectively bound to Ni^2+^ chelated by NTA ligands via N- or C-terminal polyhistidine sequences. While remaining untagged, the protein did not bind and was removed during the flow-through phase [63]. Subsequently, weakly bound impurities were removed by wash buffer (20 mM Tris-HCl, 300 mM NaCl, 40 mM imidazole, pH 7.4). Finally, the target His-tagged proteins were eluted by elution buffer (20 mM Tris-HCl, 300 mM NaCl, 250 mM imidazole, pH 7.4), which is consistent with the mechanism of NTA-Ni^2+^ affinity chromatography reported in the literature [4,5]. The column volume remained stable over the applied flow rate range.

3.3.2. Analysis of SDS-PAGE Results

To further verify the specific binding ability of NTA-Ni^2+^ ABM to His-tagged proteins and also to assess the protein content of each fraction during the purification process, SDS-PAGE analysis was conducted on the flow-through, wash fractions, and elution peaks. In the flow-through samples (Figure 6B), no prominent target protein bands (22 kDa) appeared except for P130, suggesting that His-tagged proteins had bound to the NTA-Ni^2+^ matrix during loading. Notably, a faint 22 kDa band was observed in a few lanes at the end of the flow-through solution, suggesting slight saturation or insufficient localized binding sites at the end of the loading period, resulting in incomplete capture of trace amounts of the target protein. This phenomenon is common during IMAC purification, especially under high protein loading conditions [4].

In the washing solution (Figure 6C), target protein bands (22 kDa) did not appear, reflecting the effectiveness of the washing step in removing the non-specific binding proteins and further confirming that the target proteins were still stably bound to the ABM during the washing phase. While in the elution fraction (Figure 6C), the target His-tagged protein showed clear and strong bands, indicating that the elution process successfully dissociated the target protein. This result not only demonstrated the good selectivity of the NTA-Ni^2+^ ABM for His-tagged proteins but also coincided with the appearance of chromatographic elution peaks, further supporting the validity of the purification process. The washing and elution steps showed typical IMAC characteristics, which were highly consistent with those already reported in the literature [5].

3.3.3. His-Tagged Protein Loading Analysis of NTA-Ni2+ ABM

Based on the method in 2.5.1, the binding capacity of the NTA-Ni^2+^ ABM for 22 kDa His-tagged proteins was calculated to be approximately 15.2 ± 0.8 mg/mL, based on the quantitative analysis of the eluted fractions. The obtained binding capacity is comparable to that of commercially available Ni-NTA agarose resins, which typically exhibit protein-binding capacities of 10–20 mg/mL, depending on ligand density and experimental conditions [4,5]. This result indicates that the prepared material provides chromatographic performance comparable to commercial media, and the preparation process optimized by the response surface methodology is successful and efficient in preparing competitive protein purification materials. The present work focuses on establishing a controllable preparation strategy and understanding preparation–structure–performance relationships rather than exceeding the performance of commercial products. In addition, the NTA-Ni^2+^ ABM exhibited better binding ability, and its binding performance did not decrease significantly for over five to six purification cycles. Collectively, these outcomes confirm the improved operational stability, reusability, and regeneration potential of NTA-Ni^2+^ ABM, providing an essential basis for reducing their use costs in laboratory- and industrial-scale protein purification.

UV absorbance at 280 nm (A280) was monitored for collected fractions, and the values are indicated in the SDS-PAGE results (Figure 6B,C). Due to instrument limitations, continuous chromatograms could not be exported. Future work will include detailed chromatographic profiles.

In the present work, the metal-binding capacity of the prepared sorbent was not quantitatively determined, which represents a limitation of this study. Although the chromatographic performance toward His-tagged proteins was evaluated, quantitative analysis of ligand density and Ni^2+^ loading would provide a more complete characterization of the material. Future work will focus on determining metal-loading capacity and optimizing saturation conditions.

4. Conclusions

In this study, high-performance NTA-Ni^2+^ ABM was successfully prepared by ECH-NTA “one-step” coupling and response surface methodology (RSM) optimization. The optimal preparation conditions (A: 8.12 wt%, B: 1624.46 rpm, C: 53.86 °C) produced beads with a span value of 0.50684. The RSM model was validated using various other parameters, including R^2^, CV, F-value, adjusted R^2^, and adequate precision. SEM and AFM analysis showed that an 8 h reaction time yielded NTA-Ni^2+^ ABM with greater surface roughness and a DMT modulus of 40 GPa, indicating a stable three-dimensional network. Likewise, a higher DSE peak (116.3 °C) and a reduced dynamic contact angle suggested improved thermal stability and surface wettability. NTA-Ni^2+^ ABM also exhibited a high surface area and swelling degree. Functional evaluation of protein purification confirmed that the medium exhibited high loading capacity and excellent selectivity for His-tagged proteins.

In this study, we not only established a controlled, optimized process for the preparation of high-performance NTA-Ni^2+^ ABM but also revealed the relationship between structural features and performance through several sophisticated multiscale characterizations. The prepared media provide a reliable solution for efficient recombinant protein purification at the laboratory scale and also show promising applications in bioseparation. Future studies could further explore its industrial scale-up production process, long-term stability, and purification efficiency in complex biological sample systems.