1. Introduction
Molecularly imprinted polymers (MIPs) with antibody-like binding properties have been extensively utilized as synthetic receptors for affinity separation, sensing, catalysis, and drug design
[1],
[2],
[3]. Recognition sites formed by MIPs have a specific matching relationship with the template molecule, similar to a lock-and-key concept
[4],
[5]. To date, various synthetic methods have been employed for the fabrication of MIPs, including surface imprinting
[6], bulk polymerization
[7], and precipitation polymerization
[8]. Among these methods, the surface imprinting strategy can overcome the influence of steric hindrance, allowing template molecules to easily access the recognition sites of MIPs. Moreover, the imprinted template molecules can be effectively eluted, leading to a significant increase in the number of highly active recognition sites
[9]. Nanomaterials such as nanospheres
[10], nanotubes
[11], and nanosheets
[12] are commonly used as substrates for surface imprinting. Among these, nanosheets are highly intriguing because of their high aspect ratio, large specific surface area, and excellent mechanical properties. It must be emphasized that precise control of the imprinting process through multiple complementary self-assembly steps is crucial to enhance the recognition efficiency of MIPs. However, there are few reports about imprinted nanocavities based on nanosheet substrates. The main challenge lies in achieving spatially precise orientation and fixation of the template molecules around functional monomers. Great efforts have been made to develop bioinspired dual receptors and post-translational modifications (PMs) to address the aforementioned challenges
[5].
Dual receptors are physiological or pharmacological mechanisms that are typically used as dual sites to enhance specific binding. For example, multiple amino acids can act simultaneously on the same neurotransmitter, hormone, drug, or toxin
[13],
[14],
[15],
[16]. Recently, the concept of dual receptors was simulated and applied during the molecular imprinting process to significantly enhance the affinity and selectivity of MIPs
[17]. Koide et al.
[18] developed an abiotic protein affinity reagent through the copolymerization of monomers with functionality that complemented the heparin and vascular endothelial growth factor receptor (VEGFR)-2 binding domains of the protein. Shinde et al.
[17] synthesized urea-based imprinted polymer hosts capable of selectively binding switchable anions through multiple non-covalent bonds. Thus far, self-assembly processes based on dual receptors have consisted of non-covalent/non-covalent or non-covalent/covalent monomers
[19],
[20],
[21]. Compared with covalent bonding, non-covalent interactions such as hydrogen bonding and electrostatic interactions are easily influenced by solvent polarity, ion disturbances, solution pH, and temperature during the polymerization process. These factors can have a negative impact on the imprinting efficiency. Nevertheless, dual covalent receptors that utilize multiple complementary bonds for the precise spatial fixation of template molecules are less commonly reported, mainly because of the challenge of eluting via the breaking of covalent bonds.
PM is a physiological process in which histones undergo spontaneous changes in their binding groups in various tissues during biosynthesis to accommodate the requirements of matrix reactions
[22]. Davis
[23] and Takeuchi et al.
[24] simulated the PM process and devised a new approach for post-imprinting modifications (PIMs) using small molecules or proteins as template molecules. This PIM strategy enables site-specific modifications within the imprinting cavity that enable the accurate recognition and discrimination of molecular orientations
[25],
[26],
[27]. Other researchers have conducted relevant studies using the PIM strategy to enhance the selectivity of imprinted sites. For example, Zhao et al.
[28] developed a protein-imprinted polymer (protein-IP) using PIMs in a single-step click reaction to create a highly specific fluorescence detection biosensor for glycoproteins. Hao et al.
[29] developed an imprinted monolith to enhance the detection of N-myristoylated peptides by combining MIPs with the PIM strategy and a trypsin-immobilized enzyme reactor (IMER). A disulfide-bonded functional monomer was designed to covalently assemble with the template molecule. Subsequently, the disulfide linker was transformed into a free thiol inside the binding cavities after the removal of the template molecules. This transformation allowed for the site-specific introduction of new affinity groups to enhance the recognition ability
[30]. In this context, regulation of the specific binding activity of MIPs by incorporating dual covalent receptors through PIMs can be expected to be an important approach for the tailor-made formation of complementary binding sites.
Adenosine 5′-monophosphate (AMP) is a typical nucleoside compound and an important raw material for biochemical drugs and genetic engineering. It is also considered an intermediate for anti-tumor and anti-viral drugs
[9],
[31]. AMP contains an adenine ring and a ribose unit. These functional groups can undergo boron affinity interactions and complementary purine–pyrimidine base pairing. These units inevitably form various bonds with dual non-covalent monomers during the self-assembly process, leading to mutual interferences and imprecise arrangements of the monomers that are complementary to the template molecule
[31]. As a result, AMP fails to form a stable and precisely fixed orientation around monomers through multiple-point interactions. Therefore, the current molecular imprinting strategy for AMP recognition is hindered by defective sites in the self-designed polymers, resulting in low specific adsorption. To achieve optimal recognition properties for AMP molecules, imprinting approaches and controlled engineering of imprinted sites generally need to be carefully designed.
Herein, bioinspired surface engineering was used to incorporate dual covalent receptors for precise modification onto mesoporous silica nanosheets (mSiO
2-NS), addressing the key issue. The resulting sorbents (denoted as “D-PMIPs”) exhibited an induced fit for AMP recognition and separation. More specifically, 3-(methacryloyloxyethyldithio)propionic acid sulfo-
N-hydroxysuccinimide (NHS) ester (abbreviated as DS-FM1) was first synthesized to covalently interact with the amine group of the adenine ring on AMP via the reaction of the NHS ester. A boric-acid-containing monomer, 4-(2-acrylamidoethylcarbamoyl)-3-fluorophenylboronic acid (abbreviated as “PBA-FM2”), with a low p
Ka of 7.2, was synthesized for the precise immobilization of the
cis-diol units of the ribose on AMP. This resulted in a stable pre-assembled complex before the imprinting polymerization. These dual covalent receptors facilitated the effective fixation of the molecular orientation and arrangement of AMP during the imprinting process. They also inhibited non-specific interactions between the adenine ring and ribose part on AMP, ultimately enhancing the specific recognition of AMP molecules. Subsequently, activators regenerated by electron transfer atom transfer radical polymerization (ARGET ATRP) was performed to create surface-imprinting polymers with dual receptors (denoted as “D-FMIPs”) on the surface of the mSiO
2-NS. After cleaving the disulfide bond using tris(2-carboxyethyl) phosphine (TCEP)
[32] and breaking the covalent interaction between the boric acid and AMP in an acidic environment, the template molecules (i.e., AMP) were completely removed. This process left the boric acid and thiol groups in the imprinted cavities of the D-FMIPs. Furthermore, a functional monomer containing pyrimidine (1-[2-(2-pyridyldisulfanyl) ethyl] pyrimidine-2,4-dione, abbreviated as “DS-FM3”) was synthesized to react with the exposed thiol group. The position of the introduced pyrimidine groups corresponded to the adenine groups from the imprinted AMP. By replacing the functional group within the cavity with PIMs, post-surface imprinting modification polymers (i.e., the D-PMIPs) were fabricated. Finally, we investigated the equilibrium, rebinding kinetics, selectivity, actual sample application, and regeneration of the resulting D-PMIPs toward AMP.
2. Experimental section
2.1. Materials and equipment
Hexadecyl trimethyl ammonium bromide (CTAB), tetraethylorthosilicate (TEOS), 3-chloropropyltriethoxysilane (CPTES), 2′-deoxyguanosine (dG), 2′-deoxycytidine (dC), 2′-deoxyadenosine (dA), AMP monohydrate, dimethyl sulfoxide (DMSO), N,N,N′,N″,N‴-pentamethyldiethylenetriamine (PMDETA), TCEP hydrochloride, ascorbic acid, N,N′-methylenebisacrylamide (MBAA), and alizarin red S (ARS) were purchased from Aladdin (China). Adenosine triphosphate (ATP), hydrochloric acid (HCl), acetonitrile, cupric chloride (CuCl2), cuprous chloride (CuCl), ethanol, tetrahydrofuran (THF), deproteinated calf serum injection, potassium dihydrogen phosphate (KH2PO4), and dibasic sodium phosphate (Na2HPO4) were purchased from Sinopharm Chemical Reagent Co., Ltd. (China). The physiologically active boronic acid monomer PBA-FM2 was purchased from Shanghai Macklin Biochemical Technology Co., Ltd. (China). Deionized water was obtained through a Milli-Q water-purification system, and none of the reagents were purified before use. Details regarding the equipment used in this work are provided in Appendix A.
2.2. Preparation of D-FMIPs
mSiO
2-NS were synthesized according to the method previously reported in Ref.
[33]; the synthetic procedure is described in Appendix A. Three functional monomers (Fig. S1 in Appendix A) were designed and synthesized, and their chemical structures were confirmed using proton nuclear magnetic resonance (
1H NMR) (Fig. S2 in Appendix A). DS-FM1 was synthesized according to a previous publication
[34], and the synthesis of DS-FM3 is described in Appendix A Fig. S3. The D-FMIPs were prepared as follows: AMP (0.0347 g, 0.1 mmol) was dissolved in a mixed solution of DMSO (10 mL) and phosphate-buffered saline (PBS; 10 mL, pH = 7.4, 50 mmol∙L
−1); DS-FM1 (0.0351 g, 0.1 mmol) was then added. After purging with nitrogen (N
2) for 30 min, the mixture was incubated for 12 h in the dark at 35 °C for the first covalent assembly via an ester aminolysis reaction. Next, PBA-FM2 (0.028 g, 0.1 mmol) was added to the mixture, followed by purging with N
2 for 30 min. The mixture was then incubated for 12 h in the dark at 35 °C for the second covalent assembly via boronate affinity. Subsequently, MBAA (0.123 g, 0.8 mmol) was used as crosslinking agent, and 0.05 g of mSiO
2-NS and surface chlorine groups as the substrate and initiators for ARGET ATRP were added to the above mixture. After purging with N
2 for 30 min, 0.006 g of CuCl, 0.008 g of CuCl
2, and 0.025 mL of PMDETA were sequentially added to the mixture. The mixture was purged with N
2 for 15 min, followed by the addition of 0.012 g of ascorbic acid and purging with N
2 for 30 min; then, the reaction was carried out under stirring at 35 °C for 12 h in the dark. Finally, the product was centrifuged and washed several times with methanol to remove the unreacted monomers. In order to remove the imprinted template, AMP, 10 mL of TCEP aqueous solution (20 mmol∙L
−1) and HCl solution (pH = 3.3) were used to react with the product until no AMP was detected via ultraviolet–visible spectroscopy in the eluent. The collected particles were named “D-FMIPs.” As a control, non-imprinted polymers with dual receptors (denoted as “D-FNIPs”) were also fabricated in parallel, without the addition of AMP. Importantly, MIPs integrated with a single receptor—that is, PBA-FM2 or DS-FM1—were also prepared to demonstrate the advantages of synergistic multiple binding; these were respectively named “S-BMIPs” and “S-FMIPs.”
2.3. Preparation of D-PMIPs
DS-FM3 (0.028 g, 0.1 mmol) was first dissolved in a mixed solution containing 2.0 mL of DMSO and 10 mL of PBS (pH = 7.4, 50 mmol∙L−1). Subsequently, the as-prepared D-FMIPs were added to the mixed solution, and the reaction system was stirred continuously at 35 °C for 5.0 h. Finally, the product obtained after the PIMs process (i.e. the D-PMIPs) was separated by centrifugation, washed three times with methanol, and dried in a vacuum overnight at 45 °C. Meanwhile, the D-FNIPs and S-FMIPs were also treated with the PIM process, and the binding properties of the products (i.e., D-PNIPs and S-PMIPs) were compared with those of D-PMIPs. The products (i.e., mSiO2-NS, D-PNIPs, D-PMIPs, S-BMIPs, and S-PMIPs) were then stored at room temperature, under dry, vacuum conditions.
2.4. Batch mode adsorption and selectivity experiments
Batch-mode AMP adsorption experiments were conducted with the D-PMIPs, D-PNIPs, S-PMIPs, S-BMIPs, and mSiO2-NS to investigate their adsorption kinetics, equilibrium, and regeneration. The selectivity of the D-PMIPs was evaluated with structural analogs (dA and ATP) and other typical nucleoside compounds (dC and dG). The detailed process of the adsorption experiments is provided in Appendix A.
2.5. Actual sample adsorption experiments
Actual serum solutions from three health volunteers were first treated with centrifugation and then filtered through a Millipore cellulose nitrate membrane (pore size: 0.22 μm)
[35]. Deproteinized calf blood serum injection was also selected as an actual sample, since AMP is its main active ingredient. A volume of 2.0 mL of serum solution or deproteinized calf blood serum injection was dispersed into 16 mL of PBS solution (pH = 7.4, 50 mmol∙L
−1), and then 2.0 mL of AMP solution (700 μmol∙L
–1) was added to form the spiked serum or deproteinized calf blood serum samples. Measurement of the sorbents’ ability to adsorb AMP from the spiked samples was conducted as follows
[33]: 10 mg of sorbent (i.e., D-PMIPs, D-PNIPs, S-PMIPs, S-BMIPs, and mSiO
2-NS) was added to 2.0 mL of spiked sample. The mixture was stirred in a thermostatic water bath oscillation at 25 °C for 4.0 h. Subsequently, this mixture was immediately filtered through a Millipore cellulose nitrate membrane (0.22 μm) to remove suspended impurities. The filtrate was analyzed by means of XB-C18 (5.0 μm, 4.6 mm × 200 mm) at 45 °C. The mobile phase, delivered at 0.8 mL∙min
–1, consisted of PBS solution (pH = 7.4, 50 mmol∙L
−1) and methanol in a volume ratio of 85:15. The detecting wavelength was 259 nm, and the injection volume was 10 μL.
3. Results and discussion
3.1. Design and preparation of D-PMIPs
Considering the complementarity of the monomers to the given template molecule in terms of their chemical and topological characteristics, three functional monomers (Fig. S1) were first synthesized to fix the spatial orientation and arrangement of the AMP molecule via multiple cooperative interactions and to provide for precise site-specific functionalization through the PIM process. Functional monomer 1 (DS-FM1) had an amine-reactive ester moiety for a covalent link with the amino groups of the AMP molecule; it also possessed a disulfide linkage for the later introduction of a pyrimidine group (
Fig. 1(a))
[26]. Functional monomer 2 (PBA-FM2) bore a boric acid group, which was expected to immobilize the
cis-diol moiety of AMP via boronate affinity. In addition, PBA-FM2 exhibited sufficient 1,2-
cis-diol sensitivity under physiological conditions (pH = 7.4), due to the introduction of a strongly electron-withdrawing substituent (fluorine) on the phenyl ring of the phenylboronic acid (
Fig. 1(a))
[34]. Functional monomer 3 (DS-FM3), which had a pyridyl disulfide moiety, reacted with the thiol groups in the tailor-made imprinted sites, introducing a pyrimidine group corresponding to that of the adenine group in AMP (
Fig. 1(b))
[35].
The D-PMIPs were synthesized according to the procedure shown in
Fig. 2. Importantly, two types of silane coupling agents (i.e., TEOS and CPTES), which respectively contain hydrophilic hydroxyl groups and hydrophobic chlorine groups, were selected to form a solid silica shell on the surface of polystyrene (PS) microspheres through the interfacial sol–gel method. The diameter of the PS microspheres was (2.3 ± 0.1) μm, as shown in
Fig. 3(a), while that of the PS/SiO
2 composite microsphere was (2.4 ± 0.1) μm (
Fig. 3(b)), indicating the successful fabrication of a PS/SiO
2 core–shell structure. After etching the PS spheres with THF and removing the surfactant CTAB with HCl, mSiO
2-NS were prepared by crushing the resulting hollow microspheres under ultrasonic treatment. The morphology of the resulting mSiO
2-NS displayed an irregular sheet-like structure with a smooth surface (
Figs. 3(c) and
(d)). To investigate the effect of the silane coupling agent ratio on the physical and chemical properties of the mSiO
2-NS, different volumes of CPTES (i.e., 0.10, 0.15, 0.20, 0.25, 0.30, and 0.56 mL) were used to generate PS/SiO
2 microspheres. As shown in Fig. S4 in Appendix A, a few self-polymerized nanoparticles appeared on the smooth surface of the mSiO
2-NS when the volume of CPTES was increased. The results of nitrogen adsorption–desorption demonstrated that the mSiO
2-NS were rich in mesoporous structure; when more CPTES was used, the pore size expanded from 23.89 to 46.56 Å (
Table 1; Fig. S5 in Appendix A). However, the Brunauer–Emmett–Teller (BET) surface area gradually decreased from 766.03 to 68.71 m
2∙g
–1, possibly because of the easy formation of micelles in the presence of large amounts of CPTES
[36]. The surface wettability and chlorine content of the mSiO
2-NS—as key factors for further grafting—were also evaluated. It was observed that more chlorine (Cl) groups were introduced along with an increase in CPTES, resulting in stronger hydrophobicity (Fig. S6 in Appendix A). This result can be attributed to co-condensation based on the basic catalytic reaction of TEOS and CPTES, where the hydrophobic surface is closely related to the mass of Cl groups. A Cl group can serve as an initiator of ARGET ATRP, but an aqueous environment is suited for binding AMP; thus, 0.15 mL of CPTES and 0.30 mL of TEOS comprised the optimal reaction ratio.
Second, sequential assembly and the PIM process were applied to fabricate the D-PMIPs. DS-FM1 was covalently assembled with the amino group of AMP through an amidation reaction, and then the
cis-diols of AMP were sequentially assembled with PBA-FM2 via boronate affinity to precisely fix the orientation and arrangement of the AMP. Accordingly, the non-specific effects between the functional groups of the adenine ring and the
cis-diol moieties were inhibited
[26],
[34]. Subsequently, MBAA was selected as the cross-linking agent, and chlorine-terminated mSiO
2-NS was used as the initiator to fabricate the D-FMIPs through ARGET ATRP. After grafting the MIPs onto the mSiO
2-NS, the surface of the D-FMIPs became rougher compared with that of the nanosheet substrate (
Figs. 3(e) and
(f)). To further confirm the presence of the imprinting polymer layer, the thicknesses of the mSiO
2-NS (
Fig. 3(g)) and D-FMIPs (
Fig. 3(h)) were probed using atomic force microscopy (AFM). The results showed thicknesses of about 82 and 93 nm, respectively, indicating the successful introduction of imprinted polymers. The thin imprinted polymer layer was able to overcome the influence of steric hindrance to allow AMP access and improve the efficiency of the identification. Then, the template molecules were removed by reducing the disulfide bond in DS-FM1 by means of TCEP and destroying the five-membered cyclic esters under an acidic environment, leaving only the boric acid and thiol groups in the recognition cavities
[25]. Finally, pyrimidine groups were modified onto the imprinted cavities through a sulfhydryl–disulfide exchange reaction, resulting in the formation of D-PMIPs.
3.2. Characterization of D-PMIPs
The chemical and physical properties of the synthetic mSiO
2-NS, functional sorbents (D-FMIPs and D-PMIPs), and D-PMIPs after the adsorption of AMP were characterized using Fourier-transform-infrared (FT-IR) spectroscopy, X-ray photoelectron spectroscopy (XPS), and fluorescence microscopy. The FT-IR spectra were measured and are shown in Fig. S7 in Appendix A. The peaks at 1072 and 700 cm
–1 can be attributed to the Si–O–Si and C–Cl stretching vibrations of the mSiO
2-NS, and the peak at 2957 cm
–1 is related to the C–H stretching vibration of CPTES
[37]. After grafting the imprinted polymers onto the mSiO
2-NS, a new broad peak around 3440 cm
–1 appears, which is attributed to the –OH and N–H stretching vibrations. Another new peak around 1670 cm
–1 is ascribed to the C=O stretching vibration. Furthermore, a strong stretching vibration at 2957 cm
–1 is assigned to the C–H bond of the –CH
2 and –CH
3 groups from the polymers. These results indicate the successful grafting and preparation of the D-FMIPs.
The emergence of an absorption peak at 1380 cm
–1 from the –B(OH)
2 group in the D-FMIPs and D-PMIPs provides evidence of successful functionalization with the PBA-FM2 monomer
[38]. Nevertheless, the characteristic peaks of the thiol residues and the disulfide bonds do not conform to the common ranges (i.e., at 2600–2500 and 550–430 cm
–1, respectively), because the weak or indistinct peak is deeply affected by the number of groups and the intensities of adjacent peaks. Thus, an organic element analysis was further performed for the mSiO
2-NS, D-FMIPs, and D-PMIPs to confirm the imprinting process (
Table 2). After the modification of the D-FMIPs, the nitrogen (N) element content increased to 1.433%, due to the participation of PBA-FM2 and MBAA in the imprinting process. When the PIM process was conducted with DS-FM3, the N content of the D-PMIPs increased to 5.956%, confirming the changes in the functional groups within the cavity via disulfide exchange‐reaction chemistry. These phenomena can also be proved by the changes in the sulfur (S) content of the D-FMIPs (0.318%) and D-PMIPs (0.258%) that occurred based on the oxidation-reduction reaction of the sulfhydryl groups
[39].
In addition, the PIM process was checked by means of XPS (Fig. S8(a) in Appendix A). In the wide-scan XPS spectra, B 1s and S 2p could be detected but C 1s, N 1s, and O 1s could not be seen, in the case of modification with MIPs. To further analyze the chemical composition of the D-PMIPs, the C 1s spectra of the D-PMIPs were fitted to several peaks, which can be ascribed to the C–S (284.04 eV), C–C (285.79 eV), C–B (284.66 eV), C–N (285.65 eV), C–O (286.45 eV), C–H (287.01 eV), and C=O (288.05 eV) from the functional monomers and crosslinking agent, respectively (Fig. S8(b) in Appendix A). The N 1s spectrum of the D-PMIPs was resolved into three peaks at 401.11 eV (N–C=O), 400.41 eV (N–C3), and 399.53 eV (C–N–C), illustrating the emergence of DS-FM3 and PBA-FM2 (Fig. S8(c) in Appendix A). In addition, the characteristic peak of B–O–C at 530.80 eV implied the presence of boric acid groups for immobilizing the cis-1,2-diols (Fig. S8(d) in Appendix A).
To understand the differences in the chemical composition of the D-PMIPs before and after removing AMP molecules, the B 1s and S 2p peaks in the high-resolution XPS spectra of D-FMIPs–AMP and D-PMIPs were analyzed. As AMP is released, the characteristic peak of B–O–C at 198.02 eV slightly shifts to 199.79 eV, mainly due to the increase in electronegativity (Figs. S8(e) and (f) in Appendix A). Meanwhile, the peak areas of S–S in D-FMIPs–AMP (Fig. S8(g) in Appendix A, at 161.73 eV) and D-PMIPs (Fig. S8(h) in Appendix A, at 162.07 eV) are both lower than that of S–H D-FMIPs (Fig. S8(i) in Appendix A, at 174.35 eV), mainly due to the disulfide exchange reaction
[9],
[40]. Notably, the amounts of N, S, and B modification can be roughly estimated from the atomic percentages given by XPS (
Table 3). The S content of the D-FMIPs–AMP was 1.73%, while that of the D-FMIPs was 0.9%. Then, the S content of the D-PMIPs increased to 1.4% after the PIM process
[41]. This conclusion is consistent with the analysis of organic elements, as it shows that the exposed thiol group can react with a monomer containing a pyridyl disulfide moiety to reconstruct the disulfide linkage in the recognition cavity.
To ascertain the pH responsive recognition and release performance toward a
cis-diol-containing compound, ARS-labeled D-PMIPs were immersed under different pH conditions and analyzed by means of fluorescence microscopy (
Fig. 4). The ARS-labeled D-PMIPs exhibited bright green fluorescence under alkaline (pH = 9.0,
Fig. 4(a)) and physiological conditions (pH = 7.4,
Fig. 4(b)), but there was almost no fluorescence under acidic conditions (pH = 3.3,
Fig. 4(c)). After the labeled D-PMIPs were respectively washed with solutions with different pHs and then treated with a solution with a pH of 3.3, the fluorescence intensity was significantly diminished (
Figs. 4(d)–(f)). Thus, the PBA-FM2-modified sites of the D-PMIPs display good affinity to
cis-diol-containing compounds under a physiological environment; the D-PMIPs’ remarkable pH-responsive release performance is also useful for improving the specificity by controlling the elution conditions
[42],
[43].
The N
2 adsorption–desorption isotherms of the mSiO
2-NS and D-PMIPs were also measured, as shown in
Fig. 4. Typical type-IV curves were found according to the BET classification, with type-H3 hysteresis loops, and both materials were shown to be rich in mesoporous pore structures. The BET surface areas of the mSiO
2-NS and D-PMIPs were calculated to be 498.73 and 46.12 m
2∙g
–1, respectively, with respective pore diameters of 26.92 and 95.09 Å. These results indicate that the mSiO
2-NS can provide sufficient surface area and inner space to support the imprinted MIPs. The increase in pore diameter may be attributed to the disordered growth and rough microstructure of the MIPs based on the mSiO
2-NS
[44],
[45].
3.3. Adsorption kinetics of D-PMIPs toward AMP
As shown in
Fig. 5(a), the AMP adsorption of the five sorbents was extremely rapid in the first 2.0 h and then gradually approached equilibrium. Notably, the D-PMIPs exhibited the highest adsorption rate and AMP uptake, suggesting a great affinity based on the dual covalent receptors inside the imprinted cavities. In addition, the adsorption capacity of the D-PMIPs for AMP (14.57 μmol∙g
–1) was greater than the total AMP uptakes on S-BMIPs (5.658 μmol∙g
–1) and S-PMIPs (6.845 μmol∙g
−1), revealing that there is a synergistic effect between the boronate affinity and pyrimidine base for AMP binding.
To further evaluate the effectiveness of the dual covalent receptors and PIM strategies on the adsorption efficiency, the adsorption kinetics of the mSiO
2-NS, D-PNIPs, D-PMIPs, S-BMIPs, and S-PMIPs were studied by fitting the kinetic data through several models, including pseudo-first-order and pseudo-second-order kinetic models, an Elovich model, an intra-particle diffusion model, and a Boyd plots model (Table S1 in Appendix A)
[46],
[47],
[48]. The results are shown in
Fig. 5, and the associated rate constants and linear regression values are listed in
Table 4,
Table 5. The
R2 values from the pseudo-second-order model are higher for the D-PMIPs, D-PNIPs, S-BMIPs, and S-PMIPs, indicating that the adsorption process is chemisorption. The calculated equilibrium adsorption capacity (
Qe,c, μmol∙g
–1) for the D-PMIPs from the pseudo-second-order model is closer to the experimental values (
Qe,e) (
Fig. 5(a) and
Table 4), which is consistent with the conclusion above. However, the adsorption process of the mSiO
2-NS was better described by the pseudo-first-order model (
Fig. 5(a) and
Table 4), demonstrating an adsorption mechanism based on physical adsorption (e.g., electrostatic effect).
Upon fitting with an Elovich model (
Fig. 5(b)), the related high
R2 values (> 0.95) indicated that the sorbents were covered by a superficial layer of AMP (
Table 5)
[46]. The intra-particle diffusion coefficients for the adsorption of AMP on the five sorbents were calculated from the slopes of the plot between
Qt (μmol∙g
–1) and
t1/2 (h
1/2); the fitted results of three linear regions are shown in
Fig. 5(c). For all three linear portions, the calculated slope values of the D-PMIPs are higher than those of the mSiO
2-NS, D-PNIPs, S-BMIPs, and S-PMIPs, indicating the excellent diffusion performance of the D-PMIPs. It can be concluded that the dual covalent receptors incorporated through precise PIM are effective for improving the diffusion performance. The initial instantaneous diffusion from the bulk phase into the pore surfaces—especially for the D-PMIPs—may be caused by the close contact between AMP and the affinity groups on the external surface. In addition, the
kid1 of the D-PMIPs is 11.05 g∙μmol
–1∙h
–1/2 and about 65% of saturation is achieved; therefore, intra-particle diffusion is not the rate-controlling step and the film diffusion controls the initial adsorption rate
[47]. After that, the second line segment is consistent with the intra-particle diffusion model, suggesting that AMP diffuses smoothly into the interior sites of the sorbents via the pores. However, this linear region does not go through the zero point, confirming that intra-particle diffusion is not the exclusive decisive factor for the adsorption rate. From the Boyd model fitting (
Fig. 5(d)), it can be seen that the plot of
Bt against
t is linear but does not pass through the origin, suggesting that film diffusion should be the rate-controlling step
[48].
3.4. Adsorption isotherms of D-PMIPs toward AMP
As shown in
Fig. 5(e), the amount of adsorption increases as the initial AMP concentration is raised, until the uptake equilibrium is reached. The D-PMIPs display excellent uptake ability toward AMP, although the corresponding contact time is not the shortest, which is probably related to the strong chemisorption of the imprinted sites.
In general, adsorption isotherm models are useful for describing the interaction mechanisms of a target molecule and sorbent by considering both the equilibrium data and the adsorption properties
[49]. Herein, Langmuir, Freundlich, and Temkin isotherm models were applied to fit the adsorption equilibrium data in this work (Table S2 in Appendix A). The correlation coefficients of the mSiO
2-NS, D-PNIPs, D-PMIPs, S-BMIPs, and S-PMIPs for AMP at 25 °C (pH = 7.4) are summarized in
Table 6 [50],
[51]. The adsorption data are more consistent with the Langmuir model (
Figs. 5(e) and
(f)), and the fitting of the Langmuir model (
R2 = 0.967, 0.944, 0.989, 0.982, and 0.955) is higher than those of the Freundlich model (
R2 = 0.845, 0.843, 0.875, 0.873, and 0.843) and Temkin model (
R2 = 0.951, 0.951, 0.966, 0.964, and 0.920), illustrating that the adsorption process of the five sorbents toward AMP is a monolayer adsorption. The calculated maximum adsorption capacity (
Qm, μmol∙g
–1) of the D-PMIPs is 17.47 μmol∙g
–1, which is the highest among the five studied sorbents, as shown in
Table 6. The values of the mSiO
2-NS, D-PNIPs, S-BMIPs, and S-PMIPs are 5.806, 6.972, 6.402, and 8.844 μmol∙g
–1, respectively. AMP has purine and
cis-diol moieties, and the imprinted cavities of the as-prepared D-PMIPs possess dual receptors. Thus, the strong adsorption forces based on AMP’s purine with the D-PMIPs’ pyrimidine base and AMP’s
cis-diols with the D-PMIPs’ boric acid give the D-PMIPs the highest adsorption capacity toward AMP, in what should be the primary adsorption mechanism between AMP and the D-PMIPs.
The lower adsorption capacities of the S-BMIPs and S-PMIPs may be ascribed to the single adsorption force from the pyrimidine base or
cis-diols, confirming that the synergistic effect of dual receptors is an effective strategy for enhancing adsorption capacity. Also, the adsorption capacity of the D-PNIPs is low due to the lack of imprinting recognition sites. The low adsorption of the mSiO
2-NS toward AMP is mainly attributed to physical effects (e.g., electrostatic effect). At the same time, the separation factors (
RL) were calculated according to the Langmuir model (Table S2). As shown in
Table 6, the
RL for the five sorbents ranges from 0 to 1, where the lower
RL value for the D-PMIPs (i.e., 0.083) demonstrates the D-PMIPs’ more favorable adsorption of AMP.
The maximum number (
Nmax, μmol∙g
–1) of high-/low-affinity binding sites and the binding association constant (
Ka, μmol∙L
–1) were also evaluated based on the Scatchard model (Table S2). As shown in
Fig. 5(g), two straight lines were observed for all imprinted sorbents, which likely suggests that two types of recognition sites (i.e., high- and low-affinity binding sites) are involved in the adsorption process
[52]. For imprinted sorbents, specific recognition and non-specific recognition sites may be considered for describing for two types of sites. The
Nmax of the D-PMIPs for high-affinity binding sites is 39.99 μmol∙g
–1, which corresponds to about 70.9% of all binding sites. By calculation, the number of high-affinity sites per unit area of the D-PMIPs is about 0.867 μmol∙m
–2. The total
Nmax value of the D-PMIPs (56.40 μmol∙g
–1) is 3.953 times higher than that of the S-BMIPs (14.27 μmol∙g
–1) and 4.258 times higher than that of the S-PMIPs (13.25 μmol∙g
–1), which is consistent with the results from the Langmuir model (
Table 7). It should be noted that the
Ka of the D-PMIPs for high-affinity binding sites is 284.9 μmol∙L
–1; this remarkably high value is much higher than those of the S-BMIPs and S-PMIPs. This result indicates that the dual covalent receptors and PIMs on the substrate significantly enhance the binding force of the high-affinity binding sites, efficiently increasing the availability of the D-PMIPs for the specific separation of AMP.
3.5. Selectivity binding analysis and reusability of D-PMIPs
The selective binding performance is also an important factor for the availability of sorbents. Therefore, four structural analogs of AMP (i.e., ATP, dA, dG, and dC) were adopted to study the selectivity of the four sorbents. As shown in
Fig. 5(h), the adsorption capacity of the four sorbents for AMP follows this order: D-PMIPs > S-PMIPs > D-PNIPs > S-BMIPs. The analysis results for the different nucleoside molecules can be summarized as follows:
(1) The D-PMIPs show the highest adsorption capacity for AMP (14.76 μmol∙g
–1) in comparison with dC (4.628 μmol∙g
−1), dG (4.486 μmol∙g
–1), dA (6.308 μmol∙g
–1), and ATP (6.121 μmol∙g
–1). This is mainly because of the D-PMIPs’ inability to form a complementary pyrimidine base or have a boronate affinity with the four structural analogs. The functional base groups present in dC and dG are cytosine and guanine
[53]; moreover, although dA and ATP have the same adenine as the AMP, the relative size and shape do not match the imprinted recognition sites
[19]. The S-PMIPs show a higher adsorption capacity for AMP (7.499 μmol∙g
–1), dA (5.241 μmol∙g
–1), and ATP (5.047 μmol∙g
–1) than for dC (3.770 μmol∙g
–1) and dG (3.589 μmol∙g
–1), which is due to the uracil base grouping by the PIMs in the imprinting recognition sites. Finally, the S-BMIPs show a low adsorption capacity for dA, dC, and dG due to the lack of
cis-diols.
(2) The imprinting factors (IFs) for dC, dG, AMP, dA, and ATP by the D-PMIPs are 1.031, 0.959, 2.634, 1.136, and 1.408 (
Table 8), respectively. The value for AMP in this work is obviously higher than those in previously reported studies (Table S4 in Appendix A). These observations indicate that the different functional groups, shapes, and sizes of the template molecules all contribute to the specific selectivity of the imprinted binding sites.
(3) The kd value of the D-PMIPs for AMP is 2.155 × 10–2 L⋅g–1, which is one order of magnitude higher than that for the other molecules (i.e., dC, dG, dA, and ATP), and the k′ values for the four structural analogs are all greater than 1.8. The selective recognition of the D-PMIPs for the comparison compounds follow this order: AMP > dA > ATP > dC > dG. It is also notable that the kd × 10–3 values of the four sorbents toward AMP follow this order: D-PMIPs > S-PMIPs > D-PNIPs > S-BMIPs. These findings lead us to conclude that the dual receptors enable the effective fixation of the molecular orientation and arrangement of AMP during the imprinting process, and the PIMs can precisely adjust the affinity groups in the imprinted cavities to match the adenine, thereby synergistically enhancing the selective binding performance of the D-PMIPs.
To evaluate the reusability of the D-PMIPs, an experiment on the regeneration of the D-PMIPs were carried out using HCl solution (pH = 3.3) as the eluent to destroy the boronate affinity and pyrimidine base interaction between the imprinting recognition sites and AMP. Adsorption–desorption cycles were evaluated six times with the same sample sorbents
[54]. A scanning electron microscopy (SEM) image of the D-PMIPs after six regenerations is provided in Fig. S9(a) in Appendix A. The structure shows minimal change, and a coarse surface can still be observed, indicating that the D-PMIPs exhibit good stability and reusability over multiple desorption and regeneration processes. The adsorption capacity of the recovered D-PMIPs for AMP was nearly 80% of that in the first cycle (Fig. S9(b) in Appendix A). It is likely that a mass loss of D-PMIPs or non-renewable active sites due to an insufficient removal of AMP caused a decrease in the adsorption capacity.
3.6. Separation of AMP in a real sample via D-PMIPs
In order to validate the feasibility of D-PMIPs for real applications, a standard addition method for real sample analysis was used in this work
[55]. AMP is the main active ingredient in deproteinized calf blood injection, and an obvious AMP peak located at about 4.0 min with a peak area of about 77 mAU is present in its typical chromatogram (Fig. S10(a) in Appendix A). If the AMP peak area on a chromatogram of spiked deproteinized calf blood injection is assessed after respective extraction by the five sorbents, it is expected that the D-PMIPs will still maintain an excellent uptake performance toward AMP, with at least 52.70% AMP selectively trapped by the D-PMIPs in the spiked sample (Fig. S10 in Appendix A), ensuring a promising sorbent for enriching AMP in the presence of a complex matrix. As can be seen in the figures, the present sorbent based on D-PMIPs exhibited a better selective extraction performance than the other four sorbents for the spiked serum sample, with the same trend occurring for the spiked deproteinized calf blood injection. These results indicate that the D-PMIPs present an extremely efficient solution for the selective trapping of AMP and offer an interesting approach to the enrichment of nucleoside compounds from biological samples. Moreover, the proposed PIMs and dual covalent receptors are reliable strategies for the development of imprinted sorbents with high selectivity in terms of the controllable engineering of imprinted sites.
4. Conclusions
In summary, a novel adsorption material, D-PMIPs, was developed for the specific recognition of AMP by combining dual-covalent receptors with PIMs on the surface of mSiO2-NS. The D-PMIPs have three significant advantages: First, the surface imprinting of the mSiO2-NS can be effectively eluted, leading to a significant increase in the number of highly active recognition sites. Second, the dual covalent receptors enable the effective fixation of the spatial orientation and arrangement of AMP during the imprinting process through boronate affinity and an ester aminolysis reaction. Third, the PIMs can precisely adjust the affinity groups in the imprinted cavities through sulfhydryl–disulfide exchange reactions, thereby synergistically enhancing the effectiveness of the recognition sites. Benefiting from this unique design, the as-prepared mSiO2-NS have a large surface area of 498.73 m2∙g–1, resulting in the D-PMIPs having abundant specific recognition sites and thus a high adsorption capacity of 17.47 μmol∙g–1 and outstanding recognition ability for AMP. These high-performance features ensure the D-PMIPs’ efficiency in selectively trapping AMP from biological samples. Accordingly, this work may offer a new concept for developing imprinted sorbents with high selectivity through the controllable engineering of imprinted sites.
Acknowledgments
This work was financially supported by the National Natural Science Foundation of China (22078132, 22108103, and U22A20413), the Open Funding Project of the National Key Laboratory of Biochemical Engineering (2021KF-02), China Postdoctoral Science Foundation (2021M691301), Jiangsu Key Research and Development Program (BE2022356), the Postdoctoral Fellowship Program of China Postdoctoral Science Foundation (CPSF) (GZ20230989), and Jiangsu Agricultural Independent Innovation Fund Project (CX(21)3079).
Compliance with ethics guidelines
Pan Wang, Tao Cheng, Zhuangxin Wei, Lu Liu, Yue Wang, Xiaohua Tian, and Jianming Pan declare that they have no conflict of interest or financial conflicts to disclose.
Appendix A. Supplementary data
Supplementary data to this article can be found online at
https://doi.org/10.1016/j.eng.2024.11.015.
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