An enzyme reactor based on amorphous porous organic polymers (POPs), its preparation method and application

Abstract

This invention relates to the construction of novel enzyme reactors using amorphous porous organic polymers (POPs) for in-situ immobilization of enzyme molecules under room temperature aqueous conditions, thereby obtaining a highly efficient and inexpensive heterogeneous catalyst. This invention is the first attempt to immobilize enzymes in situ using amorphous porous organic polymer materials, broadening the scope of enzyme immobilization systems and providing valuable guidance for the industrial application of immobilization systems. The various azo-linked POPs involved in this invention are composed of a series of organic monomers linked by covalent bonds, exhibiting excellent stability and enabling high enzyme loading capacity and effective protection.

Description

Technical Field

[0001] This invention belongs to the field of preparation and application of biocomposite materials. Specifically, it utilizes amorphous porous organic polymers (POPs) to immobilize enzyme molecules in situ under room temperature aqueous conditions to construct a novel enzyme reactor, thereby obtaining a highly efficient and inexpensive heterogeneous catalyst. Background Technology

[0002] Enzymes, as efficient, environmentally friendly, and sustainable biocatalysts, have enormous application potential in pharmaceuticals, energy, food, and chemical industries. However, the inherent fragility of enzymes limits their widespread application; inappropriate temperatures, pH values, and even certain mechanical stresses can damage the enzyme structure, ultimately leading to inactivation. Immobilized enzyme methods can overcome the structural fragility of enzymes and offer advantages such as simple operation, high efficiency, good stability, reusability, and cost reduction.

[0003] Traditional enzyme immobilization methods, including physical adsorption, cross-linking, and covalent bonding, generally suffer from drawbacks such as low loading capacity, poor stability, and easy leakage, resulting in low enzyme utilization and efficiency. To overcome these problems, in-situ encapsulation has become a research hotspot, offering advantages such as simplicity, speed, high loading capacity, low leakage risk, and wide applicability to various large-sized enzymes. Current research mainly focuses on metal-organic frameworks (MOFs) and hydrogen-bonded organic frameworks (HOFs) due to their high porosity, high specific surface area, and well-defined pore structure. However, the coordination bonds of MOFs and the hydrogen bonds of HOFs are extremely unstable in certain situations, making their synthesis difficult and expensive, thus limiting their application in in-situ enzyme loading. Therefore, there is an urgent need to find efficient, stable, simple to synthesize, and inexpensive in-situ systems to address this issue.

[0004] Polymers with organic pores (POPs) are highly cross-linked networks formed by one or two organic units linked by covalent interactions. They typically have non-uniform pore sizes and possess a hierarchical microporous and mesoporous structure, allowing for the selection of monomers based on specific needs. POPs materials possess large specific surface areas, stable physicochemical properties, abundant modifiable sites, and a wide variety of raw material sources, leading to their widespread application in gas storage and separation, drug release, catalysis, sensing, energy storage, and biomolecule immobilization. However, due to their amorphous nature and network interpenetration, the pores of POPs are relatively dispersed, making direct physical adsorption of enzymes through POP pores almost impossible. Therefore, research on enzyme immobilization using POPs is relatively limited. In-situ encapsulation can synthesize catalysts from POPs and enzymes in a single step, representing an effective method for enzyme immobilization using POPs.

[0005] We explored and successfully implemented the in-situ immobilization of enzymes of different sizes using porous organic polymers (POPs) under mild conditions. This approach not only maintained enzyme activity but also improved enzyme stability and reusability. Furthermore, POPs are inexpensive, simple to synthesize, and can be effectively applied in industrial applications. Due to the amorphous structure and relatively dispersed pore size distribution of POPs, traditional adsorption methods cannot be used for enzyme immobilization. We are the first to attempt in-situ enzyme immobilization using amorphous porous organic polymer materials, broadening the scope of enzyme immobilization systems and providing valuable guidance for the industrial application of immobilization systems. Summary of the Invention

[0006] This invention addresses the shortcomings of in-situ immobilized enzyme synthesis methods, such as complexity, demanding requirements, high raw material costs, and weak immobilization capabilities, by designing and synthesizing a series of room-temperature aqueous POPs. Through the control of synthesis conditions, high loading capacity and high stability of enzymes of different types and sizes can be achieved, providing a new and effective method for enzyme immobilization.

[0007] This synthesis method is simple, easy to operate, and uses inexpensive raw materials. It can achieve large-scale synthesis through simple scale-up, and the synthesis conditions are mild and aqueous, which can effectively maintain the enzyme activity.

[0008] Preferably, the present invention has screened a series of azo bond-linked amorphous porous organic polymer materials for the immobilization of various enzymes, which effectively maintains the enzyme activity.

[0009] Specifically, this invention provides a method for immobilizing enzymes in amorphous porous organic polymers (POPs) linked by nitrogen-nitrogen bonds, the specific synthesis steps of which are as follows: (1) Organic monomer 1 containing an amino group is dissolved in an acidic reagent and then reacted for a certain time under the action of a diazotizing reagent. (2) Dissolve the organic monomer 2 of phenol in an inorganic salt solution. (3) After adding a solution of biomolecules to organic monomer 1, add organic monomer 2 to carry out a coupling reaction. (4) After mixing for a certain period of time, enzyme@POPs biocatalyst is obtained, and then the final product is obtained by centrifugation and washing.

[0010] Preferably, the structure of organic monomer 1 is as follows: Figure 1 As shown, the monomer types are di-, tri-, and tetra-linked monomers containing amino functional groups. In the figure, X can be either N or C, Y can be either C or O, and M is Fe. 2+ Cu 2+ Mg 2+ Mn 3+ Zr 4+ Pd 2+ Co 2+ Ni 2+ Zn2+ Metal ions, etc.

[0011] Preferably, organic monomer 1 is a monomer containing an amino functional group. More preferably, the monomer is 1,4-phenylenediamine, 2,5-pyridinediamine, 1,5-diaminonaphthalene, biphenyldiamine, 2,2'-bipyridine-5,5'-diamine, 3,8-diaminophenanthroline, 4,4'-diaminodimethyl ketone, 4,4'-diaminodimethylethane, 4,4'-diaminodiphenyl ether, 4,4'-diaminodiphenyl sulfone, 4,4'-diaminodiphenyl disulfide, 4,4'-diaminoterphenyl, 4,4'-(1,4-phenylenebis(acetylene-2,1-diyl))diphenylamine, 1,3,5-triaminobenzene, melamine, 1,3,5-tris(4- Any one of the following: (aminophenyl)benzene, 2,4,6-tris(4-aniline)-1,3,5-triazine, 4,4',4”-triaminotriphenylmethane, 4,4',4”-triaminotriphenylamine, tetra(4-aminophenyl)methane, N,N,N',N'-tetra(p-aminophenyl)p-phenylenediamine, 4,4',4”,4”'-silanetetraethyltetraphenylamine, tetra-(4-aminophenyl)ethylene, 4,4',4”,4”'-(pyrene-1,3,6,8-tetraphenyl)tetraphenylamine, and 4,4',4”,4”-(porphyrin-5,10,15,20-tetraphenyl)tetraphenylamine. The monomer containing the amino functional group may optionally be substituted with a substituent selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. More preferably, organic monomer 1 is 1,3,5-tris(4-aminophenyl)benzene.

[0012] Preferably, the structure of organic monomer 2 is as follows: Figure 2 As shown, the monomer type is a di-, tri-, or tetra-linked monomer containing a phenolic hydroxyl functional group. More preferably, organic monomer 2 is any one of hydroquinone, resorcinol, 1,2,4,5-tetrahydroxybenzene, 1,5-dihydroxynaphthalene, 2,3,6,7-tetrahydroxynaphthalene, 4,4'-biphenyl, 2,3,6,7-tetraol-anthracene, phloroglucinol, and 2,3,6,7,10,11-hexahydroxytriphenyl. The monomer containing the phenolic hydroxyl functional group may optionally be substituted with a substituent selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. More preferably, organic monomer 2 is hydroquinone.

[0013] Preferably, the acidic reagent is a solution containing one or more inorganic acids such as hydrochloric acid, sulfuric acid, perchloric acid, and fluoroboric acid. The inorganic salt solution is a solution of inorganic salts such as sodium carbonate, sodium sulfate, sodium phosphate, potassium carbonate, potassium sulfate, and potassium phosphate. The diazotizing reagent is a solution containing sodium nitrite. More preferably, the molar ratio of the organic monomer 1 to the acidic reagent is 0.2-2:1, and the molar ratio of the organic monomer 1 to the diazotizing reagent is 0.2-1:1.

[0014] Preferably, the reaction solvent is a mixture of water and a small amount of organic solvent. More preferably, the reaction solvent is water.

[0015] Preferably, the reaction temperature is 0℃-50℃. More preferably, the reaction temperature is 4℃-25℃. Most preferably, the reaction temperature is room temperature.

[0016] Preferably, the reaction time is 1 min to 120 min. More preferably, the reaction time is 30 min to 60 min.

[0017] Preferably, the ratio of organic monomer 1 to organic monomer 2 is 0.25-4:1. More preferably, the ratio is 1:1.5.

[0018] Preferably, the concentration of the acidic reagent is 0.1-12 mol / L. More preferably, it is 0.2-0.5 mol / L.

[0019] Preferably, the concentration of the sodium nitrite solution is 0.1-1 mol / L. More preferably, it is 0.1-0.2 mol / L.

[0020] Preferably, the concentration of the selected monomer is 0.001-0.1 mol / L. More preferably, it is 0.01-0.05 mol / L.

[0021] Preferably, the amorphous porous organic polymer materials include, but are not limited to, NKPOP-1, NKPOP-2, NKPOP-3, NKPOP-4, and NKPOP-5, the specific structures of which are described in [reference needed]. Figure 3 .

[0022] Preferably, the biomolecule is selected from antibodies, nucleic acids, proteolytic enzymes, oxidoreductases, transferases, isomerases, lyases, or synthases. More preferably, the biomolecule is selected from one or more of cellulase, lipase, glucose oxidase, pepsin, laccase, amylase, lysozyme, cytochrome C, and bovine serum albumin.

[0023] Preferably, the enzyme@POPs composite catalytic material includes, but is not limited to, cellulase@NKPOP-1, cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, cellulase@NKPOP-5, lipase@NKPOP-1, glucose oxidase@NKPOP-1, glucose oxidase@NKPOP-1, cytochrome c@NKPOP-1, bovine serum albumin@NKPOP-1, and lysozyme@NKPOP-1.

[0024] Compared with the adsorption system, the in-situ enzymatic encapsulation system of POPs has a higher loading capacity and lower leakage.

[0025] Compared with other in-situ immobilized enzyme systems using porous organic materials, the POPs immobilized enzyme system is simpler to operate, operates under milder conditions, and uses cheaper raw materials.

[0026] The novel enzyme immobilization system provided by this invention broadens the scope of enzyme immobilization and industrial applications.

[0027] This invention provides an application of the aforementioned biocomposite material, characterized in that the enzyme@POPs composite catalytic material, obtained by in-situ encapsulation and immobilization of enzymes using POPs materials under room temperature aqueous conditions, maintains the catalytic activity of the enzyme molecules and can be efficiently applied as a heterogeneous catalyst for the catalysis of various reactions. This catalyst exhibits good catalytic activity, stability, and recyclability.

[0028] Preferably, the catalytic reaction includes, but is not limited to, cellulase-based hydrolysis, lipase-based hydrolysis and transesterification, glucose oxidase-based oxidation, laccase-based oxidation, and amylase-based hydrolysis.

[0029] Further preferred embodiments include, but are not limited to, the hydrolysis of sodium carboxymethyl cellulose and microcrystalline cellulose based on cellulase.

[0030] Preferably, the catalyst exhibits good catalytic activity, including but not limited to hydrolysis activity of sodium carboxymethyl cellulose and microcrystalline cellulose reaching more than 80% of that of free enzymes.

[0031] More preferably, the catalyst exhibits excellent catalytic activity, including but not limited to an activity of over 180% of the free enzyme activity in the hydrolysis of microcrystalline cellulose in an ionic liquid. More preferably, the ionic liquid is one of 1-ethyl-3-methylimidazolium acetate, 1-ethyl-3-methylimidazolium chloride, 1-butyl-3-methylimidazolium chloride, and 1-allyl-3-methylimidazolium chloride.

[0032] Preferably, the catalyst exhibits good stability, including but not limited to stability in organic solvents, high temperatures, acids, alkalis, ionic liquids, and storage stability.

[0033] Preferably, the good cycling stability of the catalyst includes, but is not limited to, cellulase@NKPOP-1, cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, and cellulase@NKPOP-5 maintaining more than 85% catalytic activity for the hydrolysis of sodium carboxymethyl cellulose after 10 cycles. Attached Figure Description

[0034] Figure 1 : The structure of organic monomer 1 for synthesizing POPs.

[0035] Figure 2 : The structure of organic monomer 2 for synthesizing POPs.

[0036] Figure 3 Schematic diagrams of the structures of several POPs.

[0037] Figure 4 Structural and morphological characterization of NKPOP-1 and cellulase@NKPOP-1.

[0038] Figure 5 Effect of cellulase loading on cellulase activity in cellulase@NKPOP-1.

[0039] Figure 6 Effect of acidity in the synthesis system of cellulase@NKPOP-1 on cellulase activity.

[0040] Figure 7 Effect of reaction temperature on cellulase activity in cellulase@NKPOP-1.

[0041] Figure 8 Effect of pH on cellulase activity in the reaction system of cellulase@NKPOP-1.

[0042] Figure 9 Catalytic activity of free cellulase and cellulase@NKPOP-1 in ionic liquids for the hydrolysis of microcrystalline cellulose.

[0043] Figure 10 FT-IR results of cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, cellulase@NKPOP-5 and their respective monomers.

[0044] Figure 11Nitrogen adsorption results for cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, and cellulase@NKPOP-5.

[0045] Figure 12 SEM and TEM results for cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, and cellulase@NKPOP-5.

[0046] Figure 13 Relative activity data for cellulase @NKPOP-1, cellulase @NKPOP-2, cellulase @NKPOP-3, cellulase @NKPOP-4, and cellulase @NKPOP-5.

[0047] Figure 14 Electron micrographs of lysozyme@NKPOP-1 and cytochrome c@NKPOP-1.

[0048] Figure 15 Electron micrographs of lipase@NKPOP-1 and β-mannanase@NKPOP-1.

[0049] Figure 16 Infrared spectra of lysozyme@NKPOP-1, lipase@NKPOP-1, cytochrome c@NKPOP-1, and β-mannanase@NKPOP-1.

[0050] Figure 17 Nitrogen adsorption results for lysozyme@NKPOP-1, lipase@NKPOP-1, cytochrome c@NKPOP-1, and β-mannanase@NKPOP-1.

[0051] Figure 18 Relative activity data for lysozyme@NKPOP-1, lipase@NKPOP-1, cytochrome c@NKPOP-1, and β-mannanase@NKPOP-1.

[0052] Figure 19 Adsorption capacity of each material for cellulase. (1: Cellulase @NKPOP-1, 2: Cellulase @NKPOP-2, 3: Cellulase @NKPOP-3, 4: Cellulase @NKPOP-4, 5: Cellulase @NKPOP-5, 6: Cellulase @COF-42-B, 7: Cellulase @NKCOF-98, 8: Cellulase @NKCOF-99, 9: Cellulase @ZIF-8, 10: Cellulase @ZIF-90, 11: Cellulase @ZPF-2)

[0053] Figure 20Relative catalytic activities of each material. (1: Cellulase @NKPOP-1, 2: Cellulase @NKPOP-2, 3: Cellulase @NKPOP-3, 4: Cellulase @NKPOP-4, 5: Cellulase @NKPOP-5, 6: Cellulase @COF-42-B, 7: Cellulase @NKCOF-98, 8: Cellulase @NKCOF-99, 9: Cellulase @ZIF-8, 10: Cellulase @ZIF-90, 11: Cellulase @ZPF-2)

[0054] Figure 21 Cellulase@NKPOP-1 cycling stability.

[0055] Figure 22 Cellulase@NKPOP-2 cycling stability.

[0056] Figure 23 Cellulase@NKPOP-3 cycling stability.

[0057] Figure 24 Cellulase@NKPOP-4 cycling stability.

[0058] Figure 25 Cellulase@NKPOP-5 cycling stability.

[0059] Figure 26 Cellulase@COF-42-B cycling stability.

[0060] Figure 27 Cellulase@NKCOF-98 cycling stability.

[0061] Figure 28 Cellulase@NKCOF-99 cycling stability.

[0062] Figure 29 Cellulase@ZIF-8 cycling stability.

[0063] Figure 30 Cellulase@ZIF-90 cycling stability.

[0064] Figure 31 Cellulase@ZPF-2 cycling stability.

[0065] Figure 32 Stability of free cellulase and cellulase@NKPOP-1 in organic solvents, acids, alkalis, and high temperatures.

[0066] Figure 33 Stability of free cellulase and cellulase@NKPOP-1 in ionic liquids.

[0067] Figure 34Storage stability of free cellulase, cellulase@NKPOP-1, cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, and cellulase@NKPOP-5.

[0068] Figure 35 : Reactor and product for large-scale preparation of cellulase@NKPOP-1. Detailed Implementation

[0069] Unless otherwise stated in the context of this application, the technical terms and abbreviations used herein have the conventional meanings known to those skilled in the art; unless otherwise stated, the raw material compounds used in the following examples are all commercially available.

[0070] The specific implementation methods for the synthesis, characterization, catalytic activity detection, and stability testing of the biocomposite materials mentioned in this invention are as follows. The following examples are for further explanation and illustration of this invention only and should not be considered as limiting the scope of the invention; the invention is limited only by the claims.

[0071] Examples 1-9 illustrate the synthesis of novel catalysts, optimization of their catalytic performance, and practical applications.

[0072] Example 1:

[0073] The synthesis of NKPOP-1 was carried out as follows: 13.2 mg of tapioca starch (TAPB) was weighed and added to a 10 mL centrifuge tube. 1.25 mL of pre-prepared 0.4 M hydrochloric acid was added. After the tapioca starch was completely dissolved, 0.75 mL of pre-prepared 0.15 M sodium nitrite solution was added. The mixture was stirred at 0-5 °C for 30 min. 6.2 mg of hydrogen hydrate (HQ) was weighed and added to 0.75 mL of pre-prepared sodium carbonate solution. The temperature was maintained at 0-5 °C. This solution was then added dropwise to the tapioca starch solution to maintain a neutral pH. After stirring for 1 h, the mixture was filtered and washed with deionized water to obtain a brown solid, NKPOP-1. Figure 4 As shown, this indicates that the material synthesis was successful.

[0074] The specific steps for synthesizing cellulase @NKPOP-1 are as follows: Weigh 13.2 mg of tapioca protein (TAPB) and add it to a 10 mL centrifuge tube. Add 1.25 mL of pre-prepared 0.4 M hydrochloric acid. After the TAPB is completely dissolved, add 0.75 mL of pre-prepared 0.15 M sodium nitrite solution and stir at 0-5 °C for 30 min. Weigh 6.2 mg of HQ and add it to 0.75 mL of pre-prepared sodium carbonate solution, maintaining the temperature at 0-5 °C. Weigh 50 mg of cellulase and dissolve it in 1 mL of deionized water to prepare a 50 mg / mL cellulase aqueous solution. Pour this solution into the TAPB solution and quickly add HQ dropwise. Stir for 1 h, then filter and wash with deionized water to obtain brown powdered cellulase @NKPOP-1. Figure 4 As shown, this indicates that the composite material was successfully synthesized.

[0075] Example 2:

[0076] Characterization of cellulase@NKPOP-1: NKPOP-1 and cellulase@NKPOP-1 were characterized by Fourier transform infrared spectroscopy (FT-IR), SEM, TEM, gas adsorption, and confocal fluorescence. These multiple characterizations corroborated each other, confirming the successful construction of NKPOP-1 and cellulase@NKPOP-1. The characterization results are shown below. Figure 4 The FT-IR spectra of HQ and NKPOP-1 are visible at ~3048 cm⁻¹. -1 The same -OH vibration peak was observed at ~1643 cm⁻¹ for NKPOP-1. -1 A new stretching vibration peak representing -N=N- was observed at ~1180 cm⁻¹, indicating the formation of a diazo bond, and the peak was observed at ~1180 cm⁻¹. -1 Characteristic peaks of the same enzyme as cellulase and cellulase@NKPOP-1 were observed, indicating that NKPOP-1 was successfully synthesized and loaded with cellulase. Nitrogen adsorption data showed that the specific surface area of ​​cellulase@NKPOP-1 was significantly lower than that of NKPOP-1, indicating that NKPOP-1 successfully encapsulated and loaded the cellulase using defect sites. SEM and TEM revealed that cellulase@NKPOP-1 exhibited aggregated granular morphology, consistent with the typical morphology of amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-1. Cellulase was labeled with fluorescein isothiocyanate (FITC), and the in-situ synthesized material was characterized by confocal laser microscopy (CLSM). Green fluorescence was clearly observed on the material after FITC labeling, indicating that the cellulase was firmly and uniformly immobilized within NKPOP-1.

[0077] Example 3:

[0078] The specific steps for detecting the activities of cellulase and cellulase@NKPOP-1 are as follows: Free cellulase: Take 1.5 mL of a prepared sodium carboxymethyl cellulose solution (2 g of sodium carboxymethyl cellulose dissolved in 300 mL of 0.1 M pH 4.5 acetate-sodium acetate buffer solution, heated to dissolve, and stored at 4℃ for later use) and add it to a 2 mL centrifuge tube. Weigh 5 mg of cellulase and add it to 0.5 mL of pH 4.5 acetate-sodium acetate buffer solution, then add this solution to a centrifuge tube containing 1.5 mL of sodium carboxymethyl cellulose. React at 50℃ for 1 h. Take 100 μL of the supernatant, add 200 μL of DNS reagent, heat in a boiling water bath for 5 min, and measure the absorbance at 540 nm using a multi-functional microplate reader. Calculate the amount of reducing sugar generated based on the standard curve.

[0079] Cellulase@NKPOP-1: Add 1.5 mL of prepared sodium carboxymethyl cellulose solution to a 2 mL centrifuge tube. Weigh 5 mg of cellulase@NKPOP-1 and add it to 0.5 mL of acetate-sodium acetate buffer solution (pH 4.5). Add this solution to the centrifuge tube containing 1.5 mL of sodium carboxymethyl cellulose and react at 50 °C for 1 h. Take 100 μL of the supernatant, add 200 μL of DNS reagent, heat in a boiling water bath for 5 min, and measure the absorbance at 540 nm using a multi-functional microplate reader. Calculate the amount of reducing sugar generated based on the standard curve.

[0080] Example 4:

[0081] The specific steps for regulating the activity of cellulase@NKPOP-1 are as follows: The study found that the cellulase loading and the amount of acid added have a significant impact on the catalyst activity during the construction of cellulase@NKPOP-1; the catalytic reaction temperature and pH also have a significant impact on the activity during the catalytic process.

[0082] Effect of cellulase loading in cellulase@NKPOP-1 on the activity of the composite catalyst: The cellulase loading of the constructed cellulase@NKPOP-1 was adjusted by controlling the cellulase concentration in the synthesis system. When the loading was 2.1 g... -1 At that time, the catalytic activity of cellulase@NKPOP-1 remained at 78% of that of free cellulase. With increasing loading, the catalytic activity of Cellulase@NKPOP-1 initially increased slightly and then rapidly decreased, reaching a low of 5.32 gg. -1 It reaches its peak at 83% of the free enzyme. Figure 5 The effect of cellulase loading on catalytic activity in cellulase@NKPOP-1.

[0083] Effect of hydrochloric acid concentration in the synthesis system on the activity of the composite catalyst: The catalytic activity of the constructed cellulase@NKPOP-1 was adjusted by controlling the concentration of hydrochloric acid in the synthesis system. The hydrochloric acid concentration was adjusted from 0.12 to 0.96 mol / L. With the increase of hydrochloric acid concentration, the catalytic activity showed a trend of first increasing and then decreasing. The highest catalytic activity (85%) was achieved when the hydrochloric acid concentration was 0.48 mol / L. Figure 6 The effect of hydrochloric acid concentration on the catalytic activity of cellulase@NKPOP-1.

[0084] Effects of temperature and pH on the catalytic activity of cellulase@NKPOP-1 in the catalytic system: The catalytic activity of the constructed composite catalyst was adjusted by controlling the reaction temperature and pH in the reaction system. The study found that the catalytic activity reached its highest level of 95% at a reaction temperature of 50℃ and a reaction pH of 4.5. Figure 7 and Figure 8 The effects of temperature and pH on the catalytic activity of cellulase@NKPOP-1 during catalysis.

[0085] Example 5:

[0086] Effects of cellulase and cellulase@NKPOP-1 on the hydrolytic activity of microcrystalline cellulose in ionic liquids: 200 mg of microcrystalline cellulose was dissolved in 5 g of ionic liquid and stored for later use. Effects of cellulase on the hydrolytic activity of microcrystalline cellulose in ionic liquids: 5 mg of cellulase was dissolved in 1 mL of acetate-sodium acetate buffer solution (pH 4.5), and 1 mL of the dissolved ionic liquid was added. The two solutions were thoroughly mixed at 50 °C and reacted for 1 h. 100 μL of the supernatant was taken, 200 μL of DNS reagent was added, and the mixture was heated in a boiling water bath for 5 min. The absorbance at 540 nm was measured using a multi-functional microplate reader.

[0087] Effect of cellulase@NKPOP-1 on the hydrolytic activity of microcrystalline cellulose in ionic liquid: Cellulase@NKPOP-1 containing 5 mg of cellulase was dissolved in 1 mL of acetate-sodium acetate buffer solution at pH 4.5. 1 mL of the dissolved ionic liquid was added, and the mixture was thoroughly combined at 50 °C and reacted for 1 h. 100 μL of the supernatant was taken, 200 μL of DNS reagent was added, and the mixture was heated in a boiling water bath for 5 min. The absorbance at 540 nm was measured using a multi-functional microplate reader.

[0088] Hydrolytic activity of cellulase and cellulase@NKPOP-1 in ionic liquids: The study found that the catalytic activity of Cellulase@NKPOP-1 increased significantly in all four ionic liquids, while the free cellulase decreased significantly in all four ionic liquids, with the maximum catalytic activity reaching 180% of that under normal conditions. Figure 9The effects of cellulase and cellulase@NKPOP-1 on the degradation ability of cellulose in different ionic liquids.

[0089] Example 6:

[0090] Synthesis of NKPOP-2: 13.2 mg of tapioca starch (TAPB) was added to a 10 mL centrifuge tube, followed by 1.25 mL of pre-prepared 0.48 M hydrochloric acid. After the TAPB was completely dissolved, 0.75 mL of pre-prepared 0.16 M sodium nitrite solution was added, and the mixture was stirred at 0-5 °C for 30 min. 4.73 mg of phloroglucinol was weighed and added to 0.75 mL of pre-prepared sodium carbonate solution. The temperature was maintained at 0-5 °C, and the solution was added dropwise to the TAPB solution to neutralize the reaction system. After stirring for 1 h, the mixture was filtered and washed to obtain black powder NKPOP-2.

[0091] Synthesizing cellulase @NKPOP-2: Add 13.2 mg TAPB to a 10 mL centrifuge tube, add 1.25 mL of pre-prepared 0.48 M hydrochloric acid, and after the TAPB is completely dissolved, add 0.75 mL of prepared 0.16 M sodium nitrite solution. Stir at 0-5 °C for 30 min. Weigh 4.73 mg of phloroglucinol and add 0.75 mL of prepared sodium carbonate solution, maintaining the temperature at 0-5 °C. Weigh 37.5 mg of cellulase and dissolve it in 1.25 mL of deionized water to prepare a 30 mg / mL cellulase aqueous solution. Pour this solution into the TAPB solution and quickly add phloroglucinol dropwise to neutralize the reaction. Stir for 1 h, then filter and wash to obtain brown powder cellulase @NKPOP-2.

[0092] Material characterization: NKPOP-2 and cellulase@NKPOP-2 were characterized by nitrogen adsorption, infrared spectroscopy, SEM, and TEM. The results of these multiple characterizations corroborated each other, proving the successful synthesis of cellulase@NKPOP-2. Characterization results are shown below. Figure 10-12 The FT-IR spectra of phloroglucinol and NKPOP-2 can be found at ~3048 cm⁻¹. -1 The same -OH vibration peak was observed at ~1643 cm⁻¹, with NKPOP-2 at ~1643 cm⁻¹. -1 A new stretching vibration peak representing -N=N- was observed at ~1180 cm⁻¹, indicating the formation of a diazo bond, and the peak was observed at ~1180 cm⁻¹. -1Characteristic peaks of the same enzyme as cellulase and cellulase@NKPOP-2 were observed, indicating that NKPOP-2 was successfully synthesized and loaded with cellulase. Nitrogen adsorption data showed that the specific surface area of ​​cellulase@NKPOP-2 was significantly lower than that of NKPOP-2, indicating that NKPOP-2 successfully encapsulated and loaded the cellulase using defect sites. SEM and TEM revealed that cellulase@NKPOP-2 exhibited aggregated granular structure, consistent with the morphology of typical amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-2.

[0093] Activity test: 1.5 mL of prepared sodium carboxymethyl cellulose solution was added to a 2 mL centrifuge tube. Cellulase@NKPOP-2 containing 5 mg of cellulase was added to 0.5 mL of acetate-sodium acetate buffer solution (pH 4.5), and then added to a centrifuge tube containing 1.5 mL of sodium carboxymethyl cellulose. The reaction was carried out at 50 °C for 1 h. 100 μL of the supernatant was taken, and 200 μL of DNS reagent was added. The mixture was heated in a boiling water bath for 5 min. The absorbance at 540 nm was measured using a multi-functional microplate reader. The amount of reducing sugar generated was calculated based on the standard curve. The results showed that the relative activity of cellulase@NKPOP-2 reached 93%. Figure 13 .

[0094] Example 7:

[0095] Synthesis of NKPOP-3 and cellulase@NKPOP-3: The synthesis method was similar to that in Example 6, with organic monomer 1 being benzidine (10.38 mg) and organic monomer 2 being phloroglucinol (7.1 mg). All other operations were as described in Example 6.

[0096] Material characterization: NKPOP-3 and cellulase@NKPOP-3 were characterized by nitrogen adsorption, infrared spectroscopy, SEM, and TEM. The results of these multiple characterizations corroborated each other, proving the successful synthesis of cellulase@NKPOP-3. The characterization results are similar to those in Example 6, see [link to Example 6]. Figure 10-12 .

[0097] Activity test: The activity test method was similar to that in Example 6. The results showed that the relative activity of cellulase@NKPOP-3 could reach 95%. Figure 13 .

[0098] Example 8:

[0099] Synthesis of NKPOP-4: 14.2 mg of melamine was added to a 40 mL centrifuge tube, followed by 7.5 mL of pre-prepared 0.48 M hydrochloric acid. After the melamine was completely dissolved, 4.5 mL of a prepared 0.16 M sodium nitrite solution was added, and the mixture was stirred at 0-5 °C for 1 day. 18.6 mg of hydroquinone was weighed and added to 4.5 mL of a prepared sodium carbonate solution. The temperature was maintained at 0-5 °C, and the solution was added dropwise to the melamine solution to neutralize the reaction system. After stirring for 1 hour, the mixture was filtered and washed to obtain a white powder, NKPOP-4.

[0100] Synthesizing cellulase @NKPOP-4: 14.2 mg of melamine was added to a 40 mL centrifuge tube, followed by 7.5 mL of pre-prepared 0.48 M hydrochloric acid. After the melamine was completely dissolved, 4.5 mL of pre-prepared 0.16 M sodium nitrite solution was added, and the mixture was stirred at 0-5 °C for 1 day. 18.6 mg of hydroquinone was weighed and added to 4.5 mL of pre-prepared sodium carbonate solution, maintaining the temperature at 0-5 °C. 75 mg of cellulase was weighed and dissolved in 7.5 mL of deionized water to prepare a 10 mg / mL cellulase aqueous solution. This solution was poured into the melamine solution, and hydroquinone was quickly added dropwise to neutralize the reaction. After stirring for 1 hour, the mixture was filtered and washed to obtain a pale yellow powder, cellulase @NKPOP-4.

[0101] Material characterization: NKPOP-4 and cellulase@NKPOP-4 were characterized by nitrogen adsorption, infrared spectroscopy, SEM, and TEM. The results of these multiple characterizations corroborated each other, proving the successful synthesis of cellulase@NKPOP-4. The characterization results are similar to those in Example 6, see [link to Example 6]. Figure 10-12 .

[0102] Activity test: The activity test method was similar to that in Example 6. The results showed that the relative activity of cellulase@NKPOP-4 reached 83%. Figure 13 .

[0103] Example 9:

[0104] Synthesis of NKPOP-5 and cellulase@NKPOP-5: The synthesis method was similar to that in Example 8. Organic monomer 2 was phloroglucinol, and the amount added was 14.2 mg. 37.5 mg of cellulase was weighed and dissolved in 7.5 mL of deionized water to prepare a 5 mg / mL cellulase aqueous solution. The remaining operations were as shown in Example 8.

[0105] Material characterization: NKPOPs and cellulase@NKPOPs were characterized by nitrogen adsorption, infrared spectroscopy, SEM, and TEM. The results of these multiple characterizations corroborated each other, confirming the successful synthesis of cellulase@NKPOP-3. The characterization results are similar to those in Example 6, see [link to Example 6]. Figure 10-12 .

[0106] Activity test: The activity test method was similar to that in Example 6. The results showed that the relative activity of cellulase@NKPOP-5 reached 86%. Figure 13 .

[0107] Example 10: Synthesis of lysozyme@NKPOP-1. The specific steps are as follows: Weigh 13.2 mg of 1,3,5-tris(4-aminophenyl)benzene and add it to a 10 mL centrifuge tube. Add 1.25 mL of pre-prepared hydrochloric acid (0.48 M) and dissolve the 1,3,5-tris(4-aminophenyl)benzene completely. Then add 0.75 mL of pre-prepared sodium nitrite solution (0.135 M) and stir at 0-5 °C for 30 min. Weigh 5.6 mg of hydroquinone and add 0.75 mL of pre-prepared sodium carbonate solution (0.48 M), maintaining the temperature at 0-5 °C. Dissolve the lysozyme in distilled water to prepare an enzyme solution of 2 mg / mL. Take 1.25 mL of the enzyme solution and pour it into the 1,3,5-tris(4-aminophenyl)benzene solution. Then quickly add hydroquinone dropwise. Stir for 1 h, filter, and wash three times with deionized water to obtain brown powder lysozyme@NKPOP-1.

[0108] Material characterization: Lysozyme@NKPOP-1 was characterized by infrared spectroscopy, nitrogen adsorption, TEM, and SEM. The results of these multiple characterizations corroborated each other, jointly demonstrating the successful synthesis of lysozyme@NKPOP-1. The results are as follows: Figure 14 As shown, at ~1650cm -1 Characteristic peaks of the same enzyme as lysozyme and lysozyme@NKPOP-1 were observed, indicating that NKPOP-1 was successfully synthesized and loaded with lysozyme. Nitrogen adsorption data showed that the specific surface area of ​​lysozyme@NKPOP-1 was significantly lower than that of NKPOP-1, indicating that NKPOP-1 successfully encapsulated and loaded lysozyme using defect sites. SEM and TEM revealed that lysozyme@NKPOP-1 exhibits aggregated granular structure, consistent with the morphology of typical amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-1.

[0109] The specific steps for detecting the activity of lysozyme and lysozyme@NKPOP-1 are as follows: Free lysozyme: Add 2.8 mL of phosphate buffer (0.2 M, pH 6.2) to a cuvette, then add 100 μL of 5 mg / mL laccase solution to the cuvette, and then add 100 μL of 3 mg / mL Micrococcus lysate solution to the cuvette. After mixing, use a UV-Vis spectrophotometer to detect the change in absorbance at 450 nm in real time.

[0110] Activity assay of lysozyme@NKPOP-1: 2.9 mL of phosphate buffer (0.2 M, pH 6.2) was added to a cuvette. Then, a solution of lysozyme@NKPOP-1 containing 50 μg of lysozyme was added to the cuvette, followed by 100 μL of 3 mg / mL *Micrococcus lysinensis* solution. After mixing, the absorbance at 450 nm was monitored in real-time using a UV-Vis spectrophotometer. The results showed that the relative activity of lysozyme@NKPOP-1 reached 75%. Figure 18 .

[0111] Example 11: Synthesis of lipase @NKPOP-1. The specific implementation steps are as follows: Weigh 13.2 mg of 1,3,5-tris(4-aminophenyl)benzene and add it to a 10 mL centrifuge tube. Add 1.25 mL of pre-prepared hydrochloric acid (0.48 M) and dissolve the 1,3,5-tris(4-aminophenyl)benzene completely. Then add 0.75 mL of prepared sodium nitrite solution (0.135 M) and stir at 0-5℃ for 30 min. Weigh 5.6 mg of hydroquinone and add 0.75 mL of prepared sodium carbonate solution (0.48 M), maintaining the temperature at 0-5℃. Dissolve the lipase in distilled water to prepare an enzyme solution of 2 mg / mL. Take 1.25 mL of the enzyme solution and pour it into the 1,3,5-tris(4-aminophenyl)benzene solution. Then quickly add hydroquinone dropwise. Stir for 1 h, filter, and wash three times with deionized water to obtain brown powder lipase @NKPOP-1.

[0112] Material characterization: Lipase@NKPOP-1 was characterized by infrared spectroscopy, nitrogen adsorption, TEM, and SEM. The results of these multiple characterizations corroborated each other, demonstrating the successful synthesis of lipase@NKPOP-1. The results are as follows: Figure 14 As shown, at ~1016cm -1 Characteristic peaks of the same enzyme as lipase and lipase@NKPOP-1 were observed, indicating that NKPOP-1 was successfully synthesized and loaded with lipase. Nitrogen adsorption data showed that the specific surface area of ​​lipase@NKPOP-1 was significantly lower than that of NKPOP-1, indicating that NKPOP-1 successfully encapsulated and loaded the lipase using defect sites. SEM and TEM revealed that lipase@NKPOP-1 exhibited aggregated granular morphology, consistent with the typical morphology of amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-1.

[0113] The specific steps for detecting the activities of lipase and lipase@NKPOP-1 are as follows: Free lipase: Add 2600 μL of phosphate buffer (10 mM, pH 7.0) to a cuvette, add 200 μL of 0.5 mg / mL lipase solution to the cuvette, and then add 200 μL of 50 mM 4-nitrophenylacetonitrile solution to the cuvette. Use a UV-Vis spectrophotometer to detect the change in absorbance at 402 nm in real time.

[0114] Lipase@NKPOP-1: 2900 μL of phosphate buffer (10 mM, pH 7.0) was added to a cuvette. Lipase@NKPOP-1 containing 100 μg of lipase was added to the cuvette and dispersed evenly. Then, 100 μL of 100 mM 4-nitrophenylacetonitrile solution was added to the cuvette. The absorbance at 402 nm was monitored in real-time using a UV-Vis spectrophotometer. The results showed that the relative activity of lipase@NKPOP-1 reached 81%. Figure 18 .

[0115] Example 12: Synthesis of cytochrome c@NKPOP-1. The specific steps are as follows: Weigh 13.2 mg of 1,3,5-tris(4-aminophenyl)benzene and add it to a 10 mL centrifuge tube. Add 1.25 mL of pre-prepared hydrochloric acid (0.48 M) and dissolve the 1,3,5-tris(4-aminophenyl)benzene completely. Then add 0.75 mL of prepared sodium nitrite solution (0.135 M) and stir at 0-5 °C for 30 min. Weigh 5.6 mg of hydroquinone and add 0.75 mL of prepared sodium carbonate solution (0.48 M), maintaining the temperature at 0-5 °C. Dissolve cytochrome c in distilled water to prepare an enzyme solution of 1 mg / mL. Take 1.25 mL of the enzyme solution and pour it into the 1,3,5-tris(4-aminophenyl)benzene solution. Then quickly add hydroquinone dropwise. Stir for 1 h, filter, and wash three times with deionized water to obtain cytochrome c@NKPOP-1.

[0116] Material characterization: Cytochrome c@NKPOP-1 was characterized by infrared spectroscopy, nitrogen adsorption, TEM, and SEM. The results of these multiple characterizations corroborated each other, jointly demonstrating the successful synthesis of cytochrome c@NKPOP-1. The results are as follows: Figure 14 As shown, at ~1650cm -1Characteristic peaks of the same enzyme were observed in both cytochrome c and cytochrome c@NKPOP-1, indicating that NKPOP-1 was successfully synthesized and loaded with cytochrome c. Nitrogen adsorption data showed that the specific surface area of ​​cytochrome c@NKPOP-1 was significantly lower than that of NKPOP-1, indicating that NKPOP-1 successfully encapsulated and loaded cytochrome c using defect sites. SEM and TEM revealed that cytochrome c@NKPOP-1 exhibited aggregated granular structures, consistent with the morphology of typical amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-1.

[0117] Assay for the activity of cytochrome c and cytochrome c@NKPOP-1: Assay for the activity of free cytochrome c enzyme: 2750 μL of potassium phosphate buffer (50 mM, pH 7.0) was added to a cuvette. 100 μL of 20 mM diammonium 2,2-azido-bis(3-ethyl-benzothiazol-6-sulfonic acid) solution was added to the cuvette. 100 μL of 0.5 mg / mL cytochrome c solution and 50 μL of 0.3% wt hydrogen peroxide were added to the cuvette. After mixing, the absorbance value at 415 nm was detected in real time using a UV-Vis spectrophotometer.

[0118] Cytochrome c@NKPOP-1 activity assay: 2850 μL of potassium phosphate buffer (50 mM, pH 7.0) was added to a cuvette. 200 μL of 20 mM 2,2-azido-bis(3-ethyl-benzothiazole-6-sulfonic acid) diammonium salt solution was added to the cuvette. Then, cytochrome c@NKPOP-1 containing 50 μg of cytochrome c and 50 μL of 0.3% wt hydrogen peroxide were added to the cuvette. After mixing, the absorbance at 415 nm was monitored in real time using a UV-Vis spectrophotometer. The results showed that the relative activity of cytochrome c@NKPOP-1 reached 77%. Figure 18 .

[0119] Example 13: Synthesis of β-mannanase@NKPOP-1. The specific implementation steps are as follows: Weigh 13.2 mg of 1,3,5-tris(4-aminophenyl)benzene and add it to a 10 mL centrifuge tube. Add 1.25 mL of pre-prepared hydrochloric acid (0.48 M) and dissolve the 1,3,5-tris(4-aminophenyl)benzene completely. Then add 0.75 mL of pre-prepared sodium nitrite solution (0.135 M) and stir at 0-5 °C for 30 min. Weigh 5.6 mg of hydroquinone and add 0.75 mL of pre-prepared sodium carbonate solution (0.48 M). Maintain the temperature at 0-5 °C. β-Mannanase was dissolved in distilled water to prepare an enzyme solution of 2 mg / mL. 1.25 mL of the enzyme solution was poured into a 1,3,5-tris(4-aminophenyl)benzene solution and hydroquinone was added dropwise. After stirring for 1 h, the solution was filtered and washed three times with deionized water to obtain β-mannanase@NKPOP-1.

[0120] Material characterization: β-Mannanase@NKPOP-1 was characterized by infrared spectroscopy, nitrogen adsorption, TEM, and SEM. The results of these multiple characterizations corroborated each other, demonstrating the successful synthesis of β-Mannanase@NKPOP-1. The results are as follows: Figure 14 As shown, at ~1650cm -1 Characteristic peaks of the same enzyme as β-mannanase and β-mannanase@NKPOP-1 were observed, indicating that NKPOP-1 was successfully synthesized and loaded with β-mannanase. Nitrogen adsorption data showed that the specific surface area of ​​β-mannanase@NKPOP-1 was significantly lower than that of NKPOP-1, indicating that NKPOP-1 successfully encapsulated and loaded β-mannanase using defect sites. SEM and TEM revealed that β-mannanase@NKPOP-1 exhibits aggregated granular structure, consistent with the morphology of typical amorphous porous organic polymers, and its morphology remained unchanged compared to NKPOP-1.

[0121] The specific steps for detecting the activities of β-mannanase and β-mannanase@NKPOP-1 are as follows: Prepare DNS reagent by dissolving 18.2g of potassium sodium tartrate in 50mL of distilled water and heating to 40℃. Then, add 0.03g of 3,5-dinitrosalicylic acid, 2.1g of NaOH, and 0.5g of phenol to the hot solution in sequence, stir until dissolved, cool, and dilute to 100mL with distilled water. Store in a brown bottle at room temperature.

[0122] Detection of free β-mannanase activity: 0.3% konjac flour substrate was prepared by adding 5 mL of disodium hydrogen phosphate-citrate buffer (pH=4) and preheating in a water bath at 75℃ for 10 min. Then, 200 μg of β-mannanase was added and reacted for 10 min. 1 mL of the reaction solution was taken out, and then 3 mL of DNS reagent was added and heated in a boiling water bath for 5 min. 3 mL of the solution was then transferred to a cuvette, and the absorbance value at a wavelength of 550 nm was detected in real time using a UV-Vis spectrophotometer.

[0123] Activity assay of β-mannanase@NKPOP-1: A 0.3% konjac flour substrate was prepared in 5 mL of disodium hydrogen phosphate-citrate buffer (pH=4) and preheated in a 75℃ water bath for 10 min. Then, β-mannanase@NKPOP-1 containing 200 μg of β-mannanase was added and reacted for 10 min. 1 mL of the reaction solution was then taken, followed by the addition of 3 mL of DNS reagent. After heating in a boiling water bath for 5 min, 3 mL was transferred to a cuvette. The absorbance at 550 nm was monitored in real time using a UV-Vis spectrophotometer. The results showed that the relative activity of β-mannanase@NKPOP-1 reached 55%. Figure 18 .

[0124] Example 14: The loading capacity, catalytic activity, and cycling stability of cellulase @NKPOP-1, cellulase @NKPOP-2, cellulase @NKPOP-3, cellulase @NKPOP-4, cellulase @NKPOP-5, cellulase @COF-42-B, cellulase @NKCOF-98, cellulase @NKCOF-99, cellulase @ZIF-8, cellulase @ZIF-90, and cellulase @ZPF-2 were studied. The specific implementation steps are as follows:

[0125] The specific implementation steps for studying cellulase loading are as follows: prepare cellulase of different concentrations in a certain proportion, add cellulase during the synthesis of materials, and after the reaction, take the supernatant each time and measure the absorbance using the Bradford method. Calculate the concentration of cellulase encapsulated in situ based on the cellulase standard curve, and obtain the total mass of the catalyst after drying, thereby determining the cellulase loading. Figure 19 The loading data correspond to the following: 1. Cellulase @NKPOP-1; 2. Cellulase @NKPOP-2; 3. Cellulase @NKPOP-3; 4. Cellulase @NKPOP-4; 5. Cellulase @NKPOP-5; 6. Cellulase @COF-42-B; 7. Cellulase @NKCOF-98; 8. Cellulase @NKCOF-99; 9. Cellulase @ZIF-8; 10. Cellulase @ZIF-90; 11. Cellulase @ZPF-2.

[0126] The relative activity study of cellulase was conducted using the following steps: For each cellulase loaded with different materials, 5 mg of catalyst containing cellulase was weighed and added to 1.5 mL of sodium carboxymethyl cellulose solution. The reaction was carried out at 50°C for a certain time, and the catalytic activity was calculated by measuring the absorbance at 540 nm using the DNS method. The catalytic activity data corresponded as above.

[0127] The cycling stability study was conducted using the following steps: After the reaction was complete, the material was collected by centrifugation, and 1.5 mL of sodium carboxymethyl cellulose solution was added. The reaction was continued at 50°C, and this process was repeated continuously. The study showed that the cellulase@NKPOPs loading was significantly higher than that of other comparative materials, exhibiting the highest relative enzyme activity and significantly better cycling stability. (See attached image). Figure 18-31 .

[0128] Example 15: The stability of cellulase and cellulase@NKPOP-1 was studied. The specific steps were as follows: 5 mg of cellulase and cellulase@NKPOP-1 containing 5 mg of cellulase were added to solutions with different organic solvents, temperatures, pH values, and different ionic liquids, respectively. The solutions were allowed to stand for a period of time, and then sodium carboxymethyl cellulose solution was added. The absorbance was measured at 540 nm using the DNS method, and the activity was calculated. The study showed that cellulase@NKPOP-1 had a significant protective effect compared to cellulase, and its activity was higher than that of cellulase, indicating that it has higher stability. See Figures 32-33 .

[0129] Example 16: Scale-up study of cellulase@NKPOP-1. The specific implementation steps are as follows: Weigh 13.2g of TAPB and add it to a 10L reactor. Add 1.25L of pre-prepared 0.4M hydrochloric acid. After the TAPB is completely dissolved, add 0.75L of prepared 0.15M sodium nitrite solution and stir at 0-5℃ for 30min. Weigh 6.2g of HQ and add 0.75L of prepared sodium carbonate solution, maintaining the temperature at 0-5℃. Weigh 50g of cellulase and dissolve it in 1L of deionized water to prepare a 50g / L cellulase aqueous solution. Pour this solution into the TAPB solution and quickly add HQ dropwise. Stir for 1h, then filter, wash with deionized water, and dry to obtain brown powdered cellulase@NKPOP-1. Figure 35 As shown, this indicates that the composite material was successfully synthesized.

Claims

1. An amorphous porous organic polymer (POPs), characterized in that, The porous organic polymer is a porous organic polymer linked by nitrogen-nitrogen bonds, which is obtained by further coupling organic monomer 1, which has undergone diazotization, with organic monomer 2. Wherein, organic monomer 1 is a di-, tri-, or tetra-linked monomer containing an amino functional group that can be substituted by any substituent, and the substituent is selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 2 is a monomer containing a phenolic hydroxyl functional group that can be optionally substituted, either di-, tri-, or tetra-linked. The substituents are selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I.

2. The porous organic polymer as described in claim 1, characterized in that, The organic monomer 1 is selected from the following structures: Where X is N or C, Y can be C or O, and M is Fe. 2+ Cu 2+ Mg 2+ Mn 3+ Zr 4+ Pd 2+ Co 2+ Ni 2+ Zn 2+ R represents -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 1 is further preferably 1,4-phenylenediamine, 2,5-pyridinediamine, 1,5-diaminonaphthalene, biphenyldiamine, 2,2'-bipyridine-5,5'-diamine, 3,8-diaminephenanthroline, 4,4'-diaminodimethyl ketone, 4,4'-diaminodimethane, 4,4'-diaminodiphenyl ether, 4,4'-diaminodiphenyl sulfone, 4,4'-diaminodiphenyl disulfide, 4,4'-diaminoterphenyl, 4,4'-(1,4-phenylenebis(acetylene-2,1-diyl))diphenylamine, 1,3,5-triaminobenzene, melamine, 1,3,5-tris(4 Any one of the following: (-aminophenyl)benzene, 2,4,6-tris(4-aniline)-1,3,5-triazine, 4,4',4”-triaminotriphenylmethane, 4,4',4”-triaminotriphenylamine, tetra(4-aminophenyl)methane, N,N,N',N'-tetra(p-aminophenyl)p-phenylenediamine, 4,4',4”,4”'-silanetetraethyltetraphenylamine, tetra-(4-aminophenyl)ethylene, 4,4',4”,4”'-(pyrene-1,3,6,8-tetraphenyl)tetraphenylamine, and 4,4',4”,4”-(porphyrin-5,10,15,20-tetraphenyl)tetraphenylamine.

3. The porous organic polymer as described in claim 1, characterized in that, The organic monomer 2 is selected from the following structures: Where M is Fe 2+ Cu 2+ Mg 2+ Mn 3+ Zr 4+ Pd 2+ Co 2+ Ni 2+ Zn 2+ R represents -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 2 is further preferably any one of hydroquinone, resorcinol, 1,2,4,5-tetrahydroxybenzene, 1,5-dihydroxynaphthalene, 2,3,6,7-tetrahydroxynaphthalene, 4,4'-biphenyl, 2,3,6,7-tetraol-anthracene, phloroglucinol, and 2,3,6,7,10,11-hexahydroxytriphenyl.

4. A method for preparing a biomolecular reactor based on amorphous porous organic polymers (POPs), characterized in that, The reaction steps include the following: (1) Dissolve organic monomer 1 in an acidic reagent, and then react it under the action of a diazotizing reagent. (2) Dissolve organic monomer 2 in an inorganic salt solution. (3) Add a solution containing biomolecules to the mixed solution obtained in step (1), then add the organic monomer 2 solution obtained in step (2) to carry out a coupling reaction. (4) After mixing, a biocatalyst consisting of biomolecules coated with porous organic polymers is obtained; The porous organic polymer is a porous organic polymer linked by nitrogen-nitrogen bonds. Organic monomer 1 is a di-, tri-, or tetra-linked monomer containing an amino functional group that can be substituted by any substituent. The substituent is selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 2 is a monomer containing a phenolic hydroxyl functional group that can be optionally substituted, either di-, tri-, or tetra-linked. The substituents are selected from any one or more of -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I.

5. The preparation method according to claim 4, characterized in that, The organic monomer 1 is selected from the following structures: Where X is N or C, Y can be C or O, and M is Fe. 2+ Cu 2+ Mg 2+ Mn 3+ Zr 4+ Pd 2+ Co 2+ Ni 2+ Zn 2+ R represents -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 1 is further preferably 1,4-phenylenediamine, 2,5-pyridinediamine, 1,5-diaminonaphthalene, biphenyldiamine, 2,2'-bipyridine-5,5'-diamine, 3,8-diaminephenanthroline, 4,4'-diaminodimethyl ketone, 4,4'-diaminodimethane, 4,4'-diaminodiphenyl ether, 4,4'-diaminodiphenyl sulfone, 4,4'-diaminodiphenyl disulfide, 4,4'-diaminoterphenyl, 4,4'-(1,4-phenylenebis(acetylene-2,1-diyl))diphenylamine, 1,3,5-triaminobenzene, melamine, 1,3,5-tris(4 Any one of the following: (-aminophenyl)benzene, 2,4,6-tris(4-aniline)-1,3,5-triazine, 4,4',4”-triaminotriphenylmethane, 4,4',4”-triaminotriphenylamine, tetra(4-aminophenyl)methane, N,N,N',N'-tetra(p-aminophenyl)p-phenylenediamine, 4,4',4”,4”'-silanetetraethyltetraphenylamine, tetra-(4-aminophenyl)ethylene, 4,4',4”,4”'-(pyrene-1,3,6,8-tetraphenyl)tetraphenylamine, and 4,4',4”,4”-(porphyrin-5,10,15,20-tetraphenyl)tetraphenylamine.

6. The preparation method according to claim 4, characterized in that, The organic monomer 2 is selected from the following structures: Where M is Fe 2+ Cu 2+ Mg 2+ Mn 3+ Zr 4+ Pd 2+ Co 2+ Ni 2+ Zn 2+ R represents -H, -NH2, -CN, -NO2, -OCH3, -SO3H, -CH3, CF3, -COOH, -COOCH3, -F, -Cl, -Br, and -I. Organic monomer 2 is further preferably any one of hydroquinone, resorcinol, 1,2,4,5-tetrahydroxybenzene, 1,5-dihydroxynaphthalene, 2,3,6,7-tetrahydroxynaphthalene, 4,4'-biphenyl, 2,3,6,7-tetraol-anthracene, phloroglucinol, and 2,3,6,7,10,11-hexahydroxytriphenyl.

7. The preparation method according to claim 4, characterized in that, The biomolecule is selected from antibodies, nucleic acids, proteolytic enzymes, oxidoreductases, transferases, isomerases, lyases, or synthases. More preferably, the biomolecule is selected from one or more of cellulase, lipase, glucose oxidase, pepsin, laccase, amylase, lysozyme, cytochrome C, and bovine serum albumin.

8. The preparation method according to claim 4, characterized in that, The acidic reagent is a solution containing one or more inorganic acids such as hydrochloric acid, sulfuric acid, perchloric acid, and fluoroboric acid. The diazotizing reagent is a solution containing sodium nitrite; the inorganic salt solution is a solution of inorganic salts such as sodium carbonate, sodium sulfate, sodium phosphate, potassium carbonate, potassium sulfate, and potassium phosphate.

9. A biomolecular reactor based on amorphous porous organic polymers (POPs), which is prepared by the method of claims 4-8.

10. The biomolecular reactor as described in claim 8, characterized in that, The biomolecular reactor is composed of enzyme@POPs composite catalytic materials, including but not limited to cellulase@NKPOP-1, cellulase@NKPOP-2, cellulase@NKPOP-3, cellulase@NKPOP-4, cellulase@NKPOP-5, lipase@NKPOP-1, glucose oxidase@NKPOP-1, glucose oxidase@NKPOP-1, cytochrome c@NKPOP-1, bovine serum albumin@NKPOP-1, and lysozyme@NKPOP-1.

11. Use of the biomolecular reactor as described in claims 9-10, characterized in that, This biomolecular reactor is used as a heterogeneous catalyst, preferably for catalyzing cellulase-based hydrolysis, lipase-based hydrolysis and transesterification, glucose oxidase-based oxidation, laccase-based oxidation and amylase-based hydrolysis.

12. A method for large-scale preparation of the biomolecular reactor as described in claims 9-10, wherein the reactor is prepared by the method of claims 4-8.

13. The method for large-scale preparation of the biomolecular reactor as described in claim 12, characterized in that, This biomolecular reactor rapidly prepares a kilogram-scale powdered biological agent in 1 hour under mild conditions (room temperature aqueous phase) through simple filtration, drying and other procedures. The scale of the agent increases with the scale of the reactor.

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