*Article* **Compatibility and Washing Performance of Compound Protease Detergent**

**Wei Zhang 1, Jintao Wu 1, Jing Xiao 1,2,\*, Mingyao Zhu <sup>1</sup> and Haichuan Yang <sup>1</sup>**


**Abstract:** Protease is the main enzyme of detergent. Through the combination of different proteases and the combination of protease and detergent additives, it can adapt to different washing conditions to improve the washing effect. In this experiment, whiteness determination, microscope scanning, Fourier transform infrared spectroscopy, and X-ray photoelectron spectroscopy were used to detect the whiteness values of the cloth pieces before and after washing, as well as the stain residue between the fibers on the surface of the cloth pieces. The protease detergent formula with better decontamination and anti-deposition effects was selected. The combination of alkaline protease, keratinase, and trypsin was cost-effective in removing stains. Polyacrylamide gel electrophoresis showed that the molecular weight of the protein significantly changed after adding the enzyme preparation during washing, and the molecular weight of the protein was directly proportional to protein redeposition. The composite protease had a better comprehensive decontamination effect, and when compatible with suitable surfactants, anti-redeposition agents, and water-softening agents, the compound protease detergent exhibited a stronger decontamination ability than commercial detergents.

**Keywords:** protease; detergent; surfactant; cleaning

### **1. Introduction**

Protease is added to detergents to decompose stains during laundry [1,2]. The protease breaks the polypeptide chain, the macromolecular protein is broken down into small molecule polypeptides or amino acids, and it is peeled from the fabric under the action of surfactant and external force [3]. The earliest protease used for washing was trypsin, which only needs a small amount to achieve good washing results. However, trypsin is mainly extracted from animal materials [4,5], and the raw materials and processes have certain limitations. Thus, it has been gradually replaced with alkaline protease, which can be produced by large-scale fermentation. Current enzyme-added detergents only add alkaline protease and exhibit a single washing effect. With the large-scale production of multiple varieties of proteases, the production cost of proteases has been reduced. With the improvement of living conditions, people's requirements for washing have increased, and new types of protein stain washing solutions are needed.

In 1963, Novozymes introduced protease, which led to a revolution in the industrial enzyme market and began a rapid expansion of detergent enzyme products. Currently, enzymes for detergents already account for 40% of all industrial enzymes. In the European and American markets, enzyme detergents already account for 80% of the detergent market, and almost all detergents are enzyme detergents in Japan. The current trend of enzymatic detergent research and development is extended from single protease to multiple enzymes, such as lipase, amylase, cellulase, mannanase, peroxidase, laccase, etc., and from a single type of stain to a comprehensive washing for multiple stains [6]. Improving the storage stability of proteases in detergents and washing stability under high/low temperature and

**Citation:** Zhang, W.; Wu, J.; Xiao, J.; Zhu, M.; Yang, H. Compatibility and Washing Performance of Compound Protease Detergent. *Appl. Sci.* **2022**, *12*, 150. https://doi.org/10.3390/ app12010150

Academic Editor: In Jung Kim

Received: 29 October 2021 Accepted: 18 December 2021 Published: 24 December 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

high alkaline/acidic conditions have also become new research hotspots. In order to meet the diverse functional requirements of the consumer market for detergents, Novozymes has developed a hybrid liquid detergent Medley solution. The enzymes used in the new Medley solution maintain stability and washing performance at high water content. Alkaline proteases have become key ingredients in detergent formulations [7]. With the emergence of new strains, researchers have found that other proteases can be added to detergents, such as keratinase [8]. The current problems of protease detergents are the compatibility of protease with washing auxiliaries and the stability of protease at different temperatures and pH [9,10].

The effect of trypsin [11], keratinase [12,13], and alkaline protease [14] on cleaning is obvious. The compound washing effect is better after the protease is proportioned. Detergents are added with different ingredients [15], such as surfactants [16], water-softening agents [17], anti-redeposition agents [18], softening agents [19,20], and stabilizers, to improve the washing efficiency. We selected 30 common detergent auxiliaries, matched them with protease, and compared the washing effects of different compound protease detergents, and the optimal formula was obtained. The washing power of the composite protease detergent was better than those of several common commercial detergents [21].

#### **2. Materials and Methods**

#### *2.1. Materials*

Selected proteases included alkaline protease (5.0 × 105 U/g), weakly alkaline protease (4.8 × 105 U/g), neutral protease (1.5 × 105 U/g), acid protease (8.0 × 105 U/g), trypsin (4.0 × <sup>10</sup><sup>3</sup> U/g), aminopeptidase (6.0 × 104 U/g), flavourzyme (5.0 × 105 U/g), and keratinase (1.0 × <sup>10</sup><sup>5</sup> U/g) in food-grade solid powder form from Lonct Enzymes Co., Ltd. (Linyi, China). Protease activity is expressed as protease activity units. At 40 ◦C and a certain pH, the amount of enzyme required for protease to hydrolyze casein to produce 1 μg of tyrosine per minute is one unit of enzyme activity. The pH of the reaction conditions for measuring the enzymatic activity of alkaline proteases, weakly basic proteases, aminopeptidases, and keratinases was 10.5. The pH of the reaction conditions for measuring the enzymatic activity of neutral proteases, trypsin, and flavored proteases was 7.5. The pH of the reaction conditions for measuring the enzymatic activity of acidic proteases was 3.0.

Detergent builders included polyethoxylated fatty alcohols, sodium ethoxy alkyl sulfate, dodecylbenzenesulfonic acid, triethanolamine, anhydrous sodium citrate, SNS-80, each in industrial-grade (≥90%) from Research Institute of Daily Chemical Industry (Taiyuan, China). 2-morpholineethanesulfonic acid (MES) in molecular biology grade (>99%) from Coolaber. Fatty acid methyl ester ethoxylate (FMEE), alkyl glycoside (APG), layered sodium disilicate (SKS-6) from Yousuo Chemical Technology Co., Ltd. (Linyi, China), modified oil ethoxylate (SOE) from Junxin Chemical Technology Co., Ltd. (Guangzhou, China), tea saponin from Zhongye Biotechnology Co., Ltd. (Lishui, China), sodium alginate, hyaluronic acid from Boxbio Science & Technology Co., Ltd. (Beijing, China), all of the above are industrial grade (≥90%). Sodium carboxymethyl cellulose (CMC), polyvinyl alcohol type 1799 (PVA), polyethylene glycol 6000 (PEG), polyaspartic acid (PASP), silicon dioxide (SiO2), ethylenediaminetetraacetic acid tetrasodium salt (EDTA), disodium maleate were used in chemically pure forms (≥99.5%) from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). Hydroxypropyl methylcellulose sodium (HPMC), hydroxyethyl cellulose sodium, and sodium polyacrylate were used in food-grade (>99%) from Best Food Additives Co., Ltd. (Zhengzhou, China). 4A zeolite was industrial-grade (≥90%) from Runfeng Synthetic Technology Co., Ltd. (Nantong, China). Sodium tartrate and sodium gluconate in food-grade (>99%) were acquired from Gukang Biological Engineering Co., Ltd. (Jinan, China). Sodium laurate in industrial-grade (≥90%) was from Longhui Chemical Co., Ltd. (Jinan, China). α-cyclodextrin, β-cyclodextrin, and γ-cyclodextrin were used in food-grade (>99%) from Youlezi Food Ingredients Co., Ltd. (Shanghai, China). Sulfobutylβ-cyclodextrin (Captisol), methyl-β-cyclodextrin, and 2-hydroxypropyl-β-cyclodextrin were analytical reagents (≥99.9%) from Aladdin (Shanghai, China). In the experiment, the

materials used to simulate protein stains were eggs and carbon powder. Blood used was porcine anticoagulated whole blood. Double distilled water was used in the experiments.

#### *2.2. Preparation of Soiled Cloth*

The protein stain was prepared as follows: Weigh 2.4 g of gum arabic powder and dissolve it with a little water, add 1.6 g of carbon black powder and grind for about 2 min. Transfer this carbon black stain to 120 mL of aqueous solution containing 13.8 g of whole milk powder, add another 120 mL of distilled water, homogenize with an emulsifier at 4000–5000 r/min for 30 min, then slowly add 120 mL of aqueous solution containing 25 g of egg liquid (egg white: yolk = 3:2) and continue to homogenize for 1 h.

White cotton cloth was cut into circular pieces of ~6 cm. When preparing the protein fouling cloth, the protein stain solution was heated to 40 ◦C and filtered. Then, 200 μL was added dropwise onto the white cotton cloth, soaked, pressed, and then dried. The same method was followed when preparing the dirty blood cloth.

#### *2.3. Washing Procedure*

The preparation method of the basal detergent is as follows. Add 4% polyethoxylated fatty alcohol, 2% ethoxylated alkyl sulfate, 8% dodecylbenzene sulfonic acid, 0.5% triethanolamine, and 0.5% anhydrous sodium citrate in a volume of water, stir to dissolve, and use sodium hydroxide solution to adjust the pH of the solution to 8.5–9.0.

The water used during washing was hard water (250 mg/kg), and the molar ratio of Ca2+ to Mg2+ was 6:4. The configuration method is as follows: weigh 1.67 g CaCl2 and 2.04 g MgCl2·6H2O, add water to make 10.0 L, which is 250 mg/kg hard water.

Then, 400 U/g of protease was added to the basal detergent and mixed well to prepare a 0.2% solution. A 100 mL aliquot of the solution was added to an Erlenmeyer flask, and a cloth piece was added. Various proteases have their maximum enzyme activity at 50–60 ◦C. Thus, 50 ◦C was selected as the reaction temperature [22,23]. The rotating speed was maintained at 150 r/min at 50 ◦C and washed for 50 min. Each piece of cloth was rinsed, dehydrated, and then dried.

#### *2.4. Characterization of the Cleaning Effect*

A whiteness tester was used to detect the whiteness reflectance value of the cloth surface before and after washing at the wavelength of 457 nm. We took two points on the front and back sides of the cloth piece before washing or after washing and measured the whiteness value. The average value of the four measurements is the whiteness value of the cloth piece. The stain residues on the surface and inside of the fiber before and after washing were observed with a super depth-of-field microscope (Leica DVM6A, Chongqing, China) at magnifications of 200× and 500× [24]. The state of the fibers before and after washing the cloth and the stain residues between the fibers were observed via scanning electron microscopy (SEM, Phenom pure plus, Shanghai, China) at a voltage of 10 kV [13,25]. The magnifications were 410×, 430×, and 440×. Fourier transform infrared spectroscopy (ATR-FTIR, Nicolet10, Waltham, MA, USA) was performed before and after washing of the fabric sheet [26,27]. The wavenumber range of residual functional groups was 500–4000 cm<sup>−</sup>1. Energy dispersion analysis was performed using X-ray photoelectron spectrometry (XPS, ESCALABXi+, Waltham, MA, USA) [27].

#### *2.5. Detection of Stain Protein Molecular Weight*

Eighty units of enzyme activities of alkaline protease, keratinase, and 4 U enzyme activities of trypsin were added to the protein-contaminated liquid to verify the decomposing effect of the proteases on protein stains. The reaction was carried out at 50 ◦C for 50 min, and the reaction solution was analyzed by SDS–PAGE [14,28]. Eighty units of alkaline protease, keratinase, and trypsin were added to the blood to verify the decomposing effect of the proteases on bloodstains. The reaction was carried out at 50 ◦C for 10–50 min, and the reaction solution was analyzed by SDS–PAGE. The electrophoresis gel was prepared using

the Meilun protein gel kit. The sample was added with 80 μL Tricine-SDS-PAGE loading buffer (5×), boiled for 5 min, centrifuged to obtain the supernatant, and then loaded with 10–15 μL for each sample. Electrophoresis was performed after adding the electrophoresis buffer to the electrophoresis system.

#### *2.6. Evaluation of the Effect of Blood Stains Redeposition*

5 mL of 4% blood dilution solution was prepared; added with 80 U of alkaline protease, keratinase, and trypsin; and then shaken in a water bath at 50 ◦C for 10, 20, 30, 40, and 50 min. The reaction solution was boiled for 3 min, and then protein gel electrophoresis was performed. Double-distilled water (5 mL) was added to the reaction solution, in which the white cotton cloth was immersed and allowed to stand for 12 h at 30 ◦C. The cloth was rinsed with water and dried, and then the extent of surface deposition and the relationship between the extent of protein deposition on the surface of the cloth and the molecular weight of the protein were observed.

#### *2.7. Optimization of Protease Washing Performance*

Alkaline protease and keratinase were mixed in the detergent at different ratios, and the total enzyme activity of each experiment was set to 80 U. In the experiment, the enzyme activity ratios of alkaline protease and keratinase were 20 U:60 U, 40 U:40 U, 30 U:50 U, 60 U:20 U, and 10 U:70 U. On the basis of adding 80 U protease activity per 0.2 g detergent, the enzyme activity of trypsin was added as 5 U, 10 U, 15 U, 20 U successively, and the remaining enzyme activity was supplemented to 80 U by alkaline protease and keratinase. The selected detergent auxiliaries were added to the basal detergent, and the addition amount was 1% of the mass of basal detergent. One or two additives, enzymes, and basal detergents with the best cleaning effect were selected from the surfactants, antideposition agents, water softeners, and cyclodextrins for subsequent experiments. The optimized formula was compared with four common commercial detergents in terms of washing effect.

#### **3. Results and Discussion**

#### *3.1. Washing Performance Test of Different Proteases*

The washing effect of different proteases on protein fouling cloths and dirty blood cloths was analyzed. Saleem et al. [29] reported the capability of protease to digest and convert the insoluble form of egg white and blood clot into their soluble forms. Bersic et al. [30] reported that adding protease to detergent has a better washing effect. Through the above washing method, different proteases were used to clean protein-fouling cloths and blooddirty cloths under the conditions of 80 U, 50 ◦C, and 50 min (Table 1). In the washing of protein stains, the highest reflectivity was obtained when trypsin was added, equivalent to 1.3 times upon the addition of alkaline protease, followed by keratinase. In the washing of bloodstains, the highest reflectivity was obtained when trypsin was added, equivalent to 1.6 times upon the addition of alkaline protease, followed by keratinase. Neutral protease and aminopeptidase have the worst washing effects. In washing blood-stained cloths with protease, some proteases could not completely decompose hemoglobin on the cloth piece, which caused hemoglobin to be retained by the fibers on the surface of the cloth piece and redeposited on the surface of the cotton cloth. Paul et al. [13] reported that crude keratinase could effectively remove blood and egg yolk stains and can be added to detergent products as a washing aid. In addition, the sewage used for washing does not pollute water resources. Emran et al. [31] reported that alkaline protease is the most commonly used protease in detergents because of its good thermal stability and compatibility with detergents. However, continuing research and development on detergents is important to find proteases with better effects. Commercial keratinase and trypsin have become promising choices. As a protease added to detergents, commercial keratinase is a more promising option. Trypsin has better detergent effects but is limited by price and production conditions and can be added to special detergents as appropriate.


**Table 1.** Washing performance of protease on protein-fouling and blood-dirty cloths.

The stain residue on the surface and interior of the protein fouling cloth and blood-dirty cloth before cleaning and after protease washing was observed under an ultra-depth-offield microscope and a desktop scanning electron microscope (Figure 1). A large number of stains were attached to the fiber surface and between the fibers before washing, and the amount of residual stains on the surface and between the fibers of the experimental group after washing with enzymes was significantly reduced, and the fibers were arranged loosely and smoothly. The cleaning effect of the cloth after washing with trypsin and keratinase was better than that after washing with alkaline protease.

The total reflectance of the protein-fouling cloth and blood-dirty cloth before and after washing with different proteases was analyzed using FTIR in the wavenumber range of 500–4000 cm−1. McCutcheon et al. [26] reported that the special spectral characteristics of protein detected by FTIR can reflect the residual protein stains on the surface of the fabric and the interior of the fabric fiber. ATR-FTIR analysis before washing the dirty cloth showed significant infrared absorption peaks at 1640 and 1526 cm−1, which represent the two infrared absorption peaks in the protein-peptide bond C=O stretching vibration absorption peaks and β-sheet conformation amide III band characteristic absorption band (Figure 2). The area of absorption peaks at 1640 and 1526 cm−<sup>1</sup> of the stained cloth washed with enzymes was significantly reduced, indicating that protein stains were separated from the cloth after being decomposed by enzymes. Alkaline protease, keratinase, and trypsin can all decompose proteins to different degrees, but trypsin and keratinase were better than alkaline protease in decomposing proteins.

Elemental analysis on the cloth was performed using XPS before and after washing with alkaline protease, keratinase, and trypsin (Figure 3). The protein-fouling cloth and blood-dirty cloth before washing showed obvious peaks at 398–400 eV, indicating that they contain N elements. After washing with protease detergent, the peaks of N elements were significantly reduced. This result indicates that the protein on the surface of the dirty cloth was decomposed and removed by the proteases.

#### *3.2. Evaluation of the Ability of Protease to Resist Stain Redeposition*

The reaction solution was analyzed by SDS–PAGE (Figure 4), the molecular weight of alkaline protease was approximately 30 kDa, and the molecular weight of keratinase was approximately less than 15 kDa. After the reaction of protein stain and alkaline protease, the molecular weight of the protein was concentrated in 50–80 kDa, and a small part of the molecular weight was concentrated in 25–30 kDa. After the protein stain liquid was hydrolyzed by keratinase, only a small amount of protein had a molecular weight of 30 kDa. This result indicates that the protein-decomposing effect of keratinase is significantly better than that of alkaline protease. The 4 U enzyme activity of trypsin was reacted with the protein stain solution. The molecular weight of the protein after the reaction was less than 25–35 kDa. The amount of trypsin added was much smaller than those of alkaline protease and keratinase, but the protein stains were decomposed effectively. A comparison

of the SDS-PAGE results with the application results showed that the protease with a better washing effect could break down protein stains from large molecules into small molecules, and the stains were easier to remove under the action of detergents.

**Figure 1.** Surface image of protein-fouling cloth. (**a**) Unwashed protein-fouling cloth (500×). (**b**) Proteinfouling cloth after washing with alkaline protease (500×). (**c**) Protein-fouling cloth after washing with keratinase (500×). (**d**) Protein-fouling cloth after washing with trypsin (500×). Surface image of blood-dirty cloth (**e**) Unwashed blood-dirty cloth (500×). (**f**) Blood-dirty cloth after washing with alkaline protease (500×). (**g**) Blood-dirty cloth after washing with keratinase (500×). (**h**) Blood-dirty cloth after washing with trypsin (500×). SEM images of protein-fouling cloth before and after washing. (**i**) Unwashed protein-fouling cloth (420×). (**j**) Protein-fouling cloth after washing with alkaline protease (420×). (**k**) Protein-fouling cloth after washing with keratinase (420×). (**l**) Protein-fouling cloth after washing with trypsin (420×). (**m**) Blood-dirty cloth after washing with keratinase (440×). (**n**) Blood-dirty cloth after washing with trypsin (410×).

**Figure 2.** Infrared spectra of dirty cloth before and after washing with different proteases. (**A**) Proteinfouling cloth. (**B**) Blooddirty cloth.

**Figure 3.** X-ray photoelectron spectroscopy of dirty cloth before and after washing with different proteases. (**A**) Protein-fouling cloth. (**B**) Blood-dirty cloth.

**Figure 4.** SDS-PAGE result graph. (**A**) M is Marker. (i) Protein stain and alkaline protease. (ii) Alkaline protease and double-distilled water. (iii) Protein stain and keratinase. (iv) Keratinase and doubledistilled water. (v) Protein stain and alkaline protease and keratinase. (vi) Alkaline protease and keratinase and double-distilled water. (vii) Protein stain and trypsin. (viii) Trypsin and doubledistilled water. (**B**) M is Marker. (i) Protein stain.

The bloodstains on the surface of the cloth treated with alkaline protease, keratinase, and trypsin were compared (Figure 5). Protein molecular weights after treatment with alkaline protease were 70 and 25 kDa. Bloodstains were deposited on the surface of the

cloth, but the deposit was relatively reduced as the enzymatic hydrolysis time was prolonged. Bloodstains were analyzed after treatment with keratinase. The protein molecular weights were mainly 50, 40, and 25 kDa. The bloodstains were deposited in the cloth piece. As the enzymatic hydrolysis time was prolonged, the deposition gradually decreased. Compared with the alkaline protease, there were less bloodstains deposited after the keratinase hydrolysis. The solution was treated with trypsin precipitation, seen from the electrophoresis pattern analysis, after 10 min, 20 min, and 30 min reactions. The protein molecular weights were mainly 50 kDa, 22 kDa, 15 kDa, 10 kDa, and macromolecular protein gradually reduced; after 40 min and 50 min reactions, protein molecular weight was mainly concentrated below 10 kDa. Blood from the cloth sheet deposition of view was observed, and the trypsin-treated blood was not deposited onto the fabric sheet again.

**Figure 5.** Deposition effect after the blood is decomposed by proteases. (**A**) Alkaline protease. (**B**) Keratinase. (**C**) Trypsin. (i) Reaction 10 min. (ii) Reaction decomposition 20 min. (iii) Reaction 30 min. (iv) Reaction 40 min. (v) Reaction 50 min. (vi) No added protease.

#### *3.3. Alkaline Protease and Detergent Auxiliary Washing Effect Test*

The effect of detergents mixed with different detergent auxiliaries and alkaline protease ratios to clean protein-soiled cloth and blood-dirty cloth was tested (Tables 2 and 3). The evaluation criterion is whether the decontamination effect of the mixed detergent is better than that of only the alkaline protease detergent. Better washing surfactants include sodium alginate, FMEE, SNS-80, hyaluronic acid, APG [32], SOE, MES, and tea saponin. Watersoftening agents with a better washing effect are 4A zeolite [33,34], sodium gluconate, sodium tartrate, and sodium laurate. The ones that inhibit the effect of alkaline protease are SKS-6, SiO2, and EDTA. Experimental results show that in the washing process of using alkaline protease detergent, whether it is for protein or bloodstains, it can be mixed with water-softening agent sodium gluconate, surfactant sodium alginate, FMEE, hyaluronic acid, APG, MES, and tea saponin [35,36]. In addition, bloodstain washing can be mixed with anti-deposition agent HPMC, sodium hydroxyethyl cellulose [37], sodium polyacrylate, and PVA. SOE, SiO2, CMC, disodium maleate, PASP, PEG, EDTA, and EDTA exert inhibitory effects on the protein decomposing effect of alkaline protease.

**Table 2.** Washing effect of different surfactants and water-softening agents mixed with alkaline protease on dirty cloth.


**Table 3.** Washing effect of anti-redeposition agent mixed with alkaline protease on blood dirty cloth.


Adding cyclodextrin to the detergent can help remove blood stains. A variety of cyclodextrins [38] mixed with alkaline protease can be used to wash blood-dirty cloth. Results showed that the addition of α-cyclodextrin and β-cyclodextrin to alkaline protease detergent was better than that of γ-cyclodextrin. The solubility of α-cyclodextrin and β-cyclodextrin was not good (Table 3), and 2-hydroxypropyl-β-cyclodextrin showed better water solubility than β-cyclodextrin. Adding sulfobutyl-β-cyclodextrin to the alkaline protease detergent demonstrated a better washing effect, but it is currently mainly used in the medical field and is expensive.

#### *3.4. Trypsin, Keratinase, and Detergent Auxiliaries Washing Effect Test*

The cleaning effect of a single alkaline protease is limited. The compatibility of detergent auxiliaries suitable for alkaline protease with keratinase and trypsin must be tested to improve protease compounding and compatibility with detergent auxiliaries. The protein-fouling and blood-dirty cloths were washed after mixing the detergent auxiliaries and trypsin (Table 4 (a)). The evaluation criterion is whether the washing effect of the mixed detergent is better than that of the detergent only added with trypsin. Results showed that the washing effect of the mixed detergent was better than that of only trypsin detergent. The effect of washing a blood-dirty cloth with mixed detergent was not much different from that of only adding trypsin detergent. Adding a detergent to trypsin detergent exerted no obvious effect on washing blood-dirty cloths.


**Table 4.** Washing effect of detergent auxiliaries mixed with trypsin or keratinase on dirty cloths.

The protein-fouling and blood-dirty cloths were washed after mixing the detergent auxiliaries and keratinase (Table 4 (b)). The evaluation standard is whether the cleaning effect of the detergent after mixing is better than that of using only the keratinase detergent. For washing protein stains, it can be mixed with sodium alginate, sodium gluconate, hyaluronic acid, and 4A zeolite. For washing bloodstains, it can be mixed with sodium hydroxyethyl cellulose and CMC, followed by sodium polyacrylate, sodium alginate, hyaluronic acid, and HPMC.

#### *3.5. Compound Protease Washing Test*

The washing effect of alkaline protease alone is limited. Niyonzima et al. [39] reported that using multiple enzymes to work together in the washing process can decompose and remove stains efficiently. In this experiment, a compound detergent formula composed of alkaline protease, keratinase, and trypsin was prepared, and the total enzyme activity of each group of experiments was set to 80 U. In the experiment, the mixing ratios of alkaline protease and keratinase were 2:6, 4:4, 3:5, 6:2, and 1:7. Stain removal showed a slight difference when alkaline protease and keratinase were added in different proportions to the detergent during washing protein-soiled cloths. When the detergent alkaline protease was mixed with keratinase at a ratio of 1:7, the effect was greatest in washing soiled cloths with blood (Table 5).


**Table 5.** Cleaning effect of protease compound on protein-fouling cloth and blood-dirty cloth.

The detergent added with trypsin exerted obvious washing effects on protein stains. The addition amount of trypsin ranged from 5 U to 20 U. Each group to add 80 U enzyme activities of protease, wherein the proportion of alkaline protease and keratinase added in the detergent was set to 1:7. Four experimental groups had 5, 10, 15, and 20 U of trypsin added. When the amount of trypsin added in the compound detergent was 10 U, the effect of washing the protein-soiled cloth improved. When the addition amount of the composite detergent trypsin was 5 U, the washing effect with bloodstains became more pronounced (Table 6).



#### *3.6. Comparison of the Effect of Detergent Products*

Liquid detergents usually have surfactants, enzymes, water-softening agents, and anti-redeposition agents added to improve their effect [40]. The current source of trypsin is mainly extracted from animal tissues, which is relatively expensive. When the ratio of alkaline protease and keratinase was set to 1:7, the washing effect can be close to the effect of using only trypsin. Therefore, alkaline protease and keratinase were selected for the compound experiment. One or two detergent auxiliaries, enzymes, and basal detergents were combined to improve washing effects in the above experiments. Four liquid detergents with the highest sales volume on the e-commerce platform and the mixed protease detergent samples were selected to compare the effects and verify the decontamination performance of the composite detergent. The whiteness value of the protein-fouling cloth washed with formulas A, B, C, and D were significantly higher than the whiteness value of the protein-fouling cloth after washing with the selected four commercial detergents. The whiteness value of the blood-dirty cloths washed with formula B, C, and D were slightly higher than that of samples 1 and 4 and was basically the same as that of samples 2 and 3. The whiteness value of the blood-dirty cloths washed with formula A, B, C, and D were slightly higher than the whiteness value of the -cloths after washing with the selected four commercial detergents. Formula A exhibited stronger washing effects and detergency than commercial detergents (Table 7, Figure 6).

**Table 7.** Cleaning effect of protease liquid detergent samples and commercial liquid detergents on dirty cloth.


**Figure 6.** Surface image of protein-fouling cloth. (**a**) Unwashed protein-fouling cloth. (**b**) Protein-fouling cloth after alkaline protease washing. (**c**) Protein-fouling cloth after keratinase washing. (**d**) Proteinfouling cloth after trypsin washing. (**e**) Protein-fouling cloth after complex enzyme detergent. (**f**) Protein-fouling cloth after commercial detergent. Surface image of blood dirty cloth. (**g**) Unwashed blood-dirty cloth. (**h**) Blood-dirty cloth after alkaline protease washing. (**i**) Blood-dirty cloth after keratinase washing. (**j**) Blood-dirty cloth after trypsin washing. (**k**) Blood-dirty cloth after complex enzyme detergent. (**l**) Blood-dirty cloth after commercial detergent. Microscopic images of protein stains after washing protein stains with protease liquid detergent. (**m**) Protein-fouling cloth after washing (500×). (**n**) Blood-dirty cloth after washing (500×).

#### **4. Conclusions**

Adding protease to detergents can remove protein stains. The washing performance of different proteases was tested through the protein-fouling cloth model, and the dirty cloth before and after washing was characterized and analyzed. The compound enzyme of the protease selected in the experiment was more efficient in removing protein stains than a single enzyme. Protein stains were disintegrated and peeled off, and stain redeposition resistance was effective. By step-by-step matching with the protease, washing aids such as sodium alginate, sodium gluconate, and sodium carboxymethyl cellulose were selected to further improve the washing effect of the composite protease. Formula A exhibited superior washing performance over common commercial detergents and, therefore, can provide new product solutions for the expansion and development of washing products.

**Author Contributions:** Conceptualization and methodology: W.Z. and J.X.; validation: W.Z. and J.W.; data analysis: M.Z.; investigation: H.Y.; writing—original draft preparation: W.Z. and J.W.; writing—review & editing: W.Z. and J.X.; funding acquisition: J.X. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Major Science and Technology Innovation Project in Shandong Province (2019JZZY011001).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** All generated and analyzed data used to support the findings of this study are included within the article.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


### *Review* **Glucose Isomerase: Functions, Structures, and Applications**

**Ki Hyun Nam 1,2**


**Abstract:** Glucose isomerase (GI, also known as xylose isomerase) reversibly isomerizes D-glucose and D-xylose to D-fructose and D-xylulose, respectively. GI plays an important role in sugar metabolism, fulfilling nutritional requirements in bacteria. In addition, GI is an important industrial enzyme for the production of high-fructose corn syrup and bioethanol. This review introduces the functions, structure, and applications of GI, in addition to presenting updated information on the characteristics of newly discovered GIs and structural information regarding the metal-binding active site of GI and its interaction with the inhibitor xylitol. This review provides an overview of recent advancements in the characterization and engineering of GI, as well as its industrial applications, and will help to guide future research in this field.

**Keywords:** glucose isomerase; xylose isomerase; high-fructose corn syrup; HFCS; bioethanol; structure

#### **1. Introduction**

Glucose isomerase (GI, EC 5.3.1.5; also known as D-xylose ketol isomerase, xylose isomerase (XI), xylose ketoisomerase, and xylose ketol-isomerase) is widely distributed in bacteria, actinomycetes, fungi, and plants [1,2]. This enzyme is an intramolecular oxidoreductase that can interconvert aldoses and ketoses. GI occupies a pivotal position with respect to its physiological role and commercial applications [3]. It is one of the three most commonly produced industrial enzymes, along with amylase and protease [2,3]. In particular, this enzyme is extensively used in the industrial production of high-fructose corn syrup (HFCS) [2]. Moreover, the bioconversion of xylose to ethanol by GI is important for the production of bioethanol from hemicellulose [4]. In this review, I introduce the molecular function, structural characteristics, and industrial applications of GI and describe the recent crystallographic results related to GI structure.

#### **2. Function**

#### *2.1. Substrate Specificity*

Glucose/xylose isomerase (GI/XI) is known to catalyze the reversible isomerization of D-glucose and D-xylose to D-fructose and D-xylulose, respectively (Figure 1).

**Figure 1.** Catalytic reaction of glucose isomerase. Reversible isomerization between (**a**) D-glucose and D-fructose, and (**b**) D-xylose and D-xylulose.

In addition to its natural substrates, GI can isomerize various pentoses, hexoses, sugar alcohols, and sugar phosphates. It exhibits activity against a broad spectrum of sugar

**Citation:** Nam, K.H. Glucose Isomerase: Functions, Structures, and Applications. *Appl. Sci.* **2022**, *12*, 428. https://doi.org/10.3390/ app12010428

Academic Editor: Hidehiko Hirakawa

Received: 30 October 2021 Accepted: 3 December 2021 Published: 3 January 2022

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**Copyright:** © 2022 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

substrates, including D-ribose [5], L-rhamnulose [6], L-arabinose [7], and D-allose [8], although the substrate specificity of GIs can vary depending on the source strain or organism [1]. For example, GI from *Streptomyces griseofuscus* S-41 shows isomerase activity on D-glucose (activity 100%) and D-xylose (287%) as well as on D-ribose (20%) [5]. The GI from *Streptomyces rubiginosus* exhibits activity on L-arabinose, in addition to D-glucose and D-xylose [7]. L-arabinose is widely used in food products with a low glycemic index, and in pharmaceutical and chemical industries [9]. The commercially available immobilized GI from *Streptomyces murinus* (SmGI) has previously been used to produce L-rhamnulose (6-deoxy-L-sorbose) from 300 g L−<sup>1</sup> L-rhamnose in a packed-bed reactor, with a 45% conversion yield [6]. L-rhamnulose plays an important role in sugar metabolism and has wide applications in the flavor industry [6,10]. Furthermore, SmGI has been employed to produce D-allose from D-allulose in a packed bed reactor [11]. At the optimal conditions of pH 8.0 and 60 ◦C, SmGI produced an average of 150 g/L D-allose over 20 days from 500 g/L D-allulose as the substrate (productivity of 36 g/L/h). D-allose is a rare sugar used as a non-caloric and non-toxic sweetener [8]; it has attracted increasing research interest owing to its beneficial medical properties, including anti-cancer, anti-oxidant, anti-inflammatory, and anti-hypertensive effects [11]. Moreover, SmGI can isomerize D-glucose, D-xylose, D-ribose, and L-rhamnose to D-fructose, D-xylulose, D-ribulose, and L-rhamnulose, respectively [11]. Thus, besides their normal ability to interconvert D-glucose/D-fructose and D-xylose/D-xylulose, GIs can also produce rare sugars that are used in food or medicinal products. GIs from various species could be developed for industrial applications by exploiting their intrinsic substrate specificity for the isomerization for rare sugars.

#### *2.2. Metal Ions*

GI requires divalent cations as cofactors for its isomerization activity [12]. Typically, GI is known to exhibit the highest activity in the presence of divalent metal ions such as Mg2+, Mn2+ and Co2+; however, the specific metal ion requirement can vary depending on the source of the enzyme. For example, the GI from *Bacillus coagulans* shows the maximum enzyme activity in the presence of Mg2+ or Mn2+ [13], and Mn2+ is the most favorable ion for catalytic activity of the GI from *Piromyces* sp. E2 [14]; in contrast, the GI from the *Streptomyces* strain YT-5 prefers Co2+ [15]. Moreover, using a combination of these divalent metal ions has been shown to further enhance the isomerization activity. For example, the activity of EDTA-treated GI from *Streptomyces* sp. CH7 was reportedly increased by 3.6-, 2.8-, and 2.1-fold in the presence of 1 mM Mg2+, Mn2+, and Co2+, respectively, compared to its activity in the absence of any metal ions [16]. However, the enzyme activity increased 6.2-fold and 44.2-fold in the presence of the combinations 1 mM Mg2+ and 0.1 mM Co2+, and 10 mM Mg2+ and 0.1 mM Co2+, respectively [16]. Thus, during optimization, it is important to measure GI activity not only with one metal ion, but also with a combination of several metal ions. In general, Mg2+ and Mn2+ act as GI activators, whereas Co2+ acts as a protein stabilizer, maintaining the ordered conformation in the quaternary structure of GI [1].

#### *2.3. Reaction Mechanism*

The catalytic reaction of GI occurs in three major steps: (i) ring opening, (ii) isomerization, and (iii) ring closure [17] (Figure 2). The action of GI has been attributed to a hydride shift mechanism, based on results from several studies employing chemical modification, X-ray crystallography, and isotope exchange methods [18–20]. Accordingly, it is proposed that the substrate at the active site of GI exists in an open ring state, and a closed ring form product is created after isomerization via a hydride shift from C2 to C1 [18–20]. During this process, a conserved histidine residue catalyzes the proton transfer from O1 to O5. Crystallographic studies have shown that, during its isomerization, xylose initially binds to the enzyme in an open chain conformation. The GI structure contains two sites (M1 and M2) for metal binding. The metal ion at the M1 site binds to O2 and O4 of the xylose molecule, and once bound, the metal ion at the M2 site binds to O1 and O2 in the transition

state. These interactions, along with a conserved lysine residue, help catalyze the hydride shift necessary for isomerization [18–20].

**Figure 2.** Isomerization mechanism of GI. (**a**) Three major steps involved in the configuration change from D-glucose to D-fructose catalyzed by GI. (**b**) Hydride shift mechanism of GI.

#### **3. Structure**

#### *3.1. Overall Structure*

The first crystal structure of GI was reported from *Streptomyces rubiginosus* at a 4 Å resolution in 2001 [21]. To date, more than 120 crystal structures of GI/XIs from various species, such as *S. rubiginosus*, *Arthrobacter* sp. NRRL B3728, *Piromyces* sp. E2, *Actinoplanes missouriensis*, *Streptomyces olivochromogenes*, *Streptomyces diastaticus*, *Streptomyces* sp. F-1, *Streptomyces* sp. SK, *Bacteroides thetaiotaomicron* VPI-5482, *Geobacillus stearothermophilus*, *Paenibacillus* sp. R4, *Streptomyces albus*, *S. murinus*, *Thermoanaerobacterium thermosulfurigenes*, *Thermotoga neapolitana*, *Thermus caldophilus*, and *Thermus thermophilus* HB8, have been elucidated and deposited in the Protein Data Bank (PDB, www.rcsb.org). These include more than 100 crystal structures of the GI from *S. rubiginosus*; however, many of these structures have been solved and used only as model samples for the development of various X-ray technologies or studying radiation damage, because the GI from *S. rubiginosus* exhibits high-quality diffraction intensity [22–29].

Structurally, GI consists of a TIM barrel-like domain and an extended α-helix domain (Figure 3A,B) which are assembled into a functional tetramer (Figure 3C) [21]. The substratebinding pocket is formed by two protomers and the GI active site is located on the TIMbarrel fold (Figure 3C). The GI exists as a tetramer with 222 crystallographic symmetry [30] and includes four active sites for substrate isomerization.

**Figure 3.** Crystal structure of the GI from *Streptomyces rubiginosus* (SruGI). (**a**) GI protomer consists of a TIM-barrel domain and an extended α-helical domain. (**b**) The extended α-helical domain interacts with the TIM barrel domain from neighboring GI molecule. (**c**) Tetrameric GI shows 222 symmetry.

The typical GI active site contains two sites, M1 and M2, for metal binding. M1 and M2 are referred to as the structural metal and catalytic metal sites, respectively [31], as the metals bound at the M1 and M2 sites are involved in the substrate binding and isomerization mechanism, respectively. In *S. rubiginosus*, the metal at the M1 site is coordinated by Glu181, Glu217, Asp245, and Asp287, and the metal at the M2 site binds the protein via Glu217, His220, Asp255, and Asp257. These metal-bound amino acids are conserved across the GI family [32,33] (Figure 4).

Most of the crystal structures of GI are bound to the metal ions Mg2+, Mn2+, or Co2+, which are involved in the isomerase activity [1]. However, some GI crystal structures are known to contain biologically less related metal ions in their active sites [34,35]. For example, in the crystal structures of GIs from *S. rubiginosus* (PDB code: 4W4Q) [36], *Paenibacillus* sp. *R4* (PDB code: 6INT) [34], and *Piromyces* (6T8E) [35], Ca2+ is located in the M1 and M2 sites of the active site, although enzyme activity in the presence of Ca2+ has not been verified. In addition, in some crystal structures, the metal ion was placed in a position other than at the M1 and M2 sites [37]. Thus, metal ions in the crystal structures of GI require verification through biochemical experiments or X-ray analysis; additionally, precise analysis of electron density is required to confirm their position and involvement in GI substrate binding and activity.

**Figure 4.** Amino acid sequence alignment of GIs from *Streptomyces rubiginosus* (UniProt Accession: P24300), *Arthrobacter* sp. NRRL B3728 (P12070), *Piromyces* sp. E2 (Q9P8C9), *Actinoplanes missouriensis* (P12851), and *Streptomyces olivochromogenes* (P15587). Residues involved in metal binding at the M1 and M2 sites are indicated by red and blue triangles, respectively. Alignment was performed using Clustal Omega [38] and displayed using Espript [39].

#### *3.2. Metal-Binding State at the GI Active Site*

With regard to metal binding, the active site of GI can exist in three different states: (i) twometal-binding state, (ii) one-metal-binding state, and (iii) metal free-state (Figure 5) [23,33,40]. Although GI crystal structures exhibit different metal-binding states at their active site, the overall GI fold is almost similar, indicating that metal binding does not influence the overall protein folding [40].

**Figure 5.** Three different metal-binding states in the active site of the GI from *Streptomyces rubiginosus* (SruGI). (**a**) Two-metal-binding mode (PDB code: 6IRK), (**b**) one-metal-binding mode (PDB code: 5Y4I), and (**c**) metal-free state of SruGI (PDB code: 7CJP).

In the two-metal- binding state of GI, the M1 and M2 sites at active site are occupied by divalent metal ions such as Mg2+, Mn2+, and Co2+, and the differences in enzyme activity occur depending on the metal occupying the active site [41,42]. In the crystal structure of the GI from *S. rubiginosus*, determined via cryo-crystallography, the metal ion at the M1 site is coordinated by Glu181, Glu217, Asp245, and Asp287, as well as the glycerol or ethylene glycol molecule, which is used as a cryoprotectant [33,40]. Meanwhile, in the room-temperature structure of *S. rubiginosus* GI, the metal ion at the M1 site is coordinated by the conserved metal-binding residues and water molecules [23,37], and the metal ion at the M2 site is coordinated by the conserved Glu217, His220, Asp255 (binate interaction), Asp257, and water molecules. The distance between the metal-binding residues and the metal is shorter at the M1 site than at the M2 site, indicating that the M1 site has tighter metal–protein interactions than the M2 site.

In the one-metal-binding mode of GI, the metal ion exists only at the M1 site involved in substrate recognition [33], indicating the higher affinity of the M1 site for metal binding than the M2 site. This result is consistent with the tighter interaction previously observed between the metal-binding residues and the metal at the M1 site, than the interaction at the M2 site [37]. In the one-metal-binding mode, the sugar substrate or inhibitor can interact with the metal ion at the M1 site [33]. However, isomerization activity through a hydride shift mechanism cannot occur due to the absence of a metal ion at the M2 site, leading to an inactive state of GI.

In the metal-free state of GI, no metal is found at both the M1 and M2 sites [40]. In this state, the substrate cannot bind to the active site because neither the metal binding to the M2 site involved in the isomerization activity nor the metal of the M1 site to which the substrate binds exist. In the metal-free state, conformation changes in conserved metalbinding residues occur within 1 Å despite the absence of metal ions at the two metal sites [40]. This is attributed to stabilization of the position of the metal-binding residues by the neighboring residues around the active sites (Glu181 \*-Asn215, Glu217 \*-Asn247, His220 \*-Pro182, Asp245 \*-Asn215, Asp245 \*-His285, Asp255 \*-Lys183, Asp257 \*-Asn247, Asp257 \*-Lys289, and Asp287 \*-Trp16; metal-binding residues are indicated by an asterisk), which helps them maintain their positions without large conformational changes [40]. Accordingly, the metal-free state exhibits a minimum open configuration for the metal to perceive and bind to the sites [40]. Based on the existing biochemical knowledge, this study proposed that, when combining with GI in a metal-free state, the metal first binds to the M1 site and then is filled in the M2 site.

#### *3.3. Xylitol Binding to the Active Site of GI*

Structure-based inhibitor studies of GI are helpful for industrial applications such as HGFS and bioethanol production [37], by identifying the parameters of enzyme engineering to prevent product inhibition. The isomerase activity of GI can be inhibited by some divalent cations such as Ca2+, Cu2+, Zn2+, Ni2+, Ag2+, and Hg2+ [1]. In addition, molecules such as xylitol, arabitol, sorbitol, mannitol, lyxose, and Tris also inhibit GI activity [43]. However, the mechanism of GI inhibition by these compounds remains largely unclear.

Among these compounds, the inhibition of the activity of GI by xylitol, which is a structural analogue of xylose, has been well studied [44,45]. To date, six crystal structures of xylitol-bound GI have been deposited in PDB (PDB code 1XIG, 2XIS, 3GNX, 4DUO, 5Y4J, and 7DFK) [31,33,37,46,47]. All these structures reveal that the xylitol molecule binds at the M1 site in the active site of GI. Three oxygen (O2, O3, and O4) atoms from xylitol interact with the metal ion at the M1 site, and the metal ion is stabilized by octahedral coordination, by the metal-interacting residues of the enzyme, and the xylitol molecules [31,33,37,46,47]. Therefore, xylitol binding in the M1 site prevents substrate access to the metal ion. The crystal structure of one-metal-binding mode of GI shows that the metal ion at the M2 site is not essential to facilitate xylitol binding to the M1 site [33]. Furthermore, xylitol binding to the M1 site stabilizes the geometry of the metal-binding sites.

Meanwhile, the xylitol-bound GI structure shows a high B-factor value of the metal ion at the M2 site, although the reason for this observation remains unknown. A recently obtained room-temperature structure of xylitol-bound GI via serial crystallography revealed a correlation between xylitol binding and low metal occupancy at the M2 site [37]. In native GI, the M1 site shows a distorted octahedral coordination, but it is geometrically stabilized upon xylitol binding to the metal at the M1 site [37] (Figure 6A). This causes a rearrangement of the conformation of metal-binding residues in the M2 site (Figure 6C). This induces the distortion of metal coordination at the M2 site, leading to the release of the unstable coordinated metal ion from the M2 site [37]. This metal release induces further conformational changes in the neighboring residues (Figure 6C). Since the metal ion at the M2 site involved in isomerization activity is released as a result of xylitol binding, an additional catalytic metal ion has to be added for an isomerization reaction in industrial applications after the release of xylitol.

**Figure 6.** Proposed mechanism of xylitol-induced release of metal at the M2 site of GI. (**a**) Xylitolbound state of GI. (**b**) Rearrangement of xylitol-binding residues. (**c**) Release of metal ion from M2 site of GI.

#### **4. Application of GI**

#### *4.1. High-Fructose Corn Syrup (HFCS)*

Owing to its beneficial properties, including a high solubility at low temperature, a lower tendency to crystallize than sucrose, and a high freezing point depression, fructose is widely used in the manufacture of ice cream and frozen desserts to influence taste and texture [48]. Fructose is also used in the production of cake, biscuits, bread, and other confectionery products [49,50]. Moreover, fructose increases the shelf-life of food products, with its high osmotic pressure in solution, making it a better preservative against microbial growth than sucrose syrup [51]. Additionally, fructose is used in the pharmaceutical industry to manufacture diabetes medicine, as it does not influence the blood levels of glucose and insulin [52].

HFCS is an equilibrium mixture of glucose and fructose, which has the advantages of sweetness, low cost, and high solubility [53]. Depending on the fructose content, HFCS can be classified as HFCS-42 (42% fructose, 53% glucose, and 5% polysaccharide), HFCS-55 (55% fructose, 42% glucose, and 3% polysaccharide), and HFCS-90 (90% fructose, 9% glucose and 1% polysaccharide) [54]. Among them, HFCS-55 is the most commonly used, with its commercial production involving several processes such as chromatography, purification, and concentration [55].

HFCS is widely applied in the food, detergent, and pharmaceutical industries [56]. Although glucose can be converted to fructose using a chemical process for HFCS production, this chemical reaction is non-specific and leads to the formation of non-metabolizable sugars [4]. In contrast, GI catalyzes the isomerization of glucose to fructose with excellent specificity, which is critical for the industrial production and application of HFCS [4].

As GI exhibits a reversible isomerization activity, a thermodynamic equilibrium exists in the isomerization reaction between glucose and fructose [57]. The rate of the enzymatic conversion of glucose to fructose can be increased by increasing the reaction temperature; therefore, using a highly thermostable GI capable of sustained operation at high temperatures is critical for one-step HFCS production [57].

Brown et al. reported a GI from the thermophilic eubacterium *Thermotoga maritima* (TmaGI) [58], which is produced when the bacterium is grown in the presence of xylose. TmaGI requires the metal cations Co2+ and Mg2+ for enzyme activity [58]. This enzyme prefers xylose as a substrate and is most active at pH 6.5–7.5. TmaGI displays the maximum activity at an optimum temperature of 105 to 110 ◦C, and its half-life is approximately 10 min at 120 ◦C and pH 7.0. TmaGI is a promising candidate for improving the efficiency of the industrial glucose isomerization process, as it exhibits optimal pH from neutral to slightly acidic, as well as a high thermostability [58].

Deng et al. reported the characterization of GI from *Thermobifida fusca* WSH03-11 (TfuGI) [59]. TfuGI displayed the maximum activity at an optimum temperature of 80 ◦C, with a half-life of approximately 2 h at 80 ◦C or 15 h at 70 ◦C. TfuGI was the most active at pH 10 and retained 95% of its initial activity after incubation at pH 5–10 and 4 ◦C for 24 h. Analysis of TfuGI enzyme kinetics revealed the Km and kcat values to be 197 mM and 1688 min<sup>−</sup>1, respectively [59]. TfuGI was able to convert glucose (45% *w*/*v*) to fructose, with a maximum conversion yield of 53% at pH 7.5 and 70 ◦C.

Jia et al. have characterized GIs from *Thermoanaerobacterium xylanolyticum* (TxyGI), *Thermus oshimai* (TosGI), *Geobacillus thermocatenulatus* (GthGI), and *Thermoanaerobacter siderophilus* (TsiGI) [60]. These enzymes were identified using a genome mining approach for their potential application in manufacturing HFCS at elevated temperatures with a low cost of enriching syrups. Among these enzymes, TosGI showed the highest catalytic efficiency toward D-glucose, along with superior thermostability. The optimum temperature of TosGI was 95 ◦C, and it retained more than 80% of its activity after 48 h at 85 ◦C in the presence of 20 mM Mn2+ [60]. The kinetic parameters Km and kcat/Km of TosGI were calculated to be 81.46 mM and 21.77 min−<sup>1</sup> mM<sup>−</sup>1, respectively [60]. TosGI achieved a maximum yield of 52.16% for the conversion of D-glucose (400 g/L) to D-fructose at 85 ◦C [60].

The GI from *Caldicellulosiruptor bescii* (CbeGI), characterized by Dai et al. [57], exhibited the maximum activity at pH 7.0 and 80 ◦C and retained good thermostability at 85 ◦C. CbeGI showed affinity for D-glucose, with a Km of 42.61 mM, and a conversion efficiency up to 57.3% with 3 M D-glucose [57]. The high catalytic efficiency and affinity of CbeGI make it a valuable enzyme for the direct production of 55% HFCS.

Vieille et al. reported the characterization of GI from the hyperthermophile *T. neapolitana* 5068 (TnGI) [61]. The optimal temperature of TnGI was above 95 ◦C and the optimum pH was 7.1, but the enzyme also showed high activity over a wide pH range. Kinetics studies showed that the catalytic efficiency (kcat/Km) of TnGI remained constant between 60 and 90 ◦C; however, the catalytic efficiency decreased between 90 and 98 ◦C, primarily because of a large increase in Km [61]. TnGI had a higher turnover number and a lower Km for glucose compared to other thermophilic GIs [61]. Taken together, thermophilic GIs exhibit various activities according to the species of origin and are expected to be extensively applied in industrial HFCS production.

#### *4.2. Ethanol Production*

The bioconversion of a renewable biomass into ethanol is attractive in view of the rapid depletion of fossil fuels [62]. GI can catalyze the isomerization of xylose derived from hemicellulosic biomass to xylulose, which can be fermented to ethanol by conventional yeast such as *Saccharomyces cerevisiae*, *Schizosaccharomyces pombe*, and *Candida tropicalis* [63–65]. This property of GI is attractive for the application in bioethanol production because no coenzyme is required and no intermediates are produced during the reaction [66]. However, the typical ethanol production process is plagued by low efficiency, owing to the low conversion yield of xylose to ethanol [67]. The GI enzyme engineering and strain improvement approaches have been explored to improve the conversion yield of xylose to ethanol [1,65,68]. The engineering of yeast strains that exhibit faster xylose metabolism is important in pursuing strain improvement for bioethanol production [69,70].

Ko et al. evaluated the performance of SXA-R2P-E, an engineered isomerase-based xylose-utilizing *S. cerevisiae* strain, for co-fermentation of xylose and glucose to ethanol [65]. SXA-R2P-E produced 50g/L ethanol (yield of 0.43 g ethanol/g sugar) in 72 h via a highsugar fermentation process (70 g/L glucose and 40 g/L xylose). This strain also produced 18-21 g/L ethanol (yield of 0.43–0.46 g ethanol/g sugar) from acid-pretreated lignocellulosic hydrolysates such as rice straw and hardwood [65].

Seike et al. reported improved xylose fermentation through XI expression in the xylose-utilizing *S. cerevisiae* strain IR-2, which has a deletion of the *GR3* gene (encoding an endogenous xylose reductase) and overexpresses *XK21* (encoding an additional xylulose kinase). Evolutionary engineering was performed in the IR-2 strain to select high-efficiency XIs from eight previously reported XIs derived from various species [68]: *Burkholderia cenocepacia* J2315 [71], *Lachnoclostridium phytofermentans ISDg* (LpXI) [67], *Orpinomyces* sp. ukk1 [72], *Piromyces* sp. E2 [73], *Prevotella ruminicola* TC2-24 [74], *Ruminiclostridium cellulolyticum* H10 [75], *Ruminococcus flavefaciens* 17 [76], and *S. rubiginosus* [77]. Among them, the strain expressing LpXI exhibited the highest D-xylose consumption rate after 72 h of micro-aerobic fermentation on D-glucose and D-xylose mixed medium [68]. Furthermore, to enhance the LpXI catalytic activity, the authors performed random mutagenesis using an error-prone polymerase chain reaction and obtained two enzyme constructs with improved fermentation performances [68]. The SS120 strain, expressing double mutant LpXI (T63I/V162A), produced 53.3 g/L ethanol in 72 h (ethanol yield of approximately 0.44 (g/g-input sugars) from 85 g/L D-glucose and 35 g/L D-xylose [68].

XI enzyme engineering is another approach that has been employed to increase the efficiency of bioethanol production by accelerating xylose metabolism [68,78]. For example, the XI from the fungal strain *Piromyces* sp. E2 (PirXI) is employed for xylose isomerization in engineered *S. cerevisiae* strains but has low activity in vivo. To improve the performance of PirXI, Lee et al. constructed a mutant library by substituting residues around the substrate and metal-binding sites of the enzyme [35]. PirXI variants were obtained by transferring

the library to *S. cerevisiae*, followed by in vivo selection for enhanced xylose-supported growth under aerobic and anaerobic conditions. In particular, in the presence of Mg2+ or Mn2+, the Km of PirXI-V270A/A273G variant for xylose was higher, and the kcat was lower than the corresponding values for wild-type PirXI [35].

Thus, further efforts to engineer GI/XI for a high xylose conversion rate through rational engineering or directed evolution are still needed, and applied research should be performed through in vivo studies on improved strains to enable efficient bioethanol production.

#### **5. Perspective**

In addition to its biological importance, GI is an industrially important enzyme that is applied in HFCS and bioethanol production. Although biochemical and structural studies have been conducted on GIs from various organisms, many GIs still need to be identified from the protein database. The development of more efficient GI-based industrial processes requires extensive biochemical analysis and engineering of novel GIs. In addition, it will be necessary to analyze not only in vitro enzymatic activity, but also the industrial performance of GI using whole cell systems. On the other hand, structural studies on the inhibition of GI activity by various metals, sugars, and other inhibitors should be conducted to help better modulate GI activity and enhance the enzyme performance during industrial application. Furthermore, it is necessary to understand the dynamical structural changes occurring in GI to aid enzyme engineering efforts, as most GI structures currently collected in cryogenic environments have only provided limited knowledge with regard to flexibility. The recently developed serial crystallography technique provides information regarding the molecular fluidity of proteins at room temperature, and future studies of molecular fluidity based on the room-temperature crystal structures will provide useful information for GI enzyme engineering.

**Funding:** This work was funded by the National Research Foundation of Korea (NRF) (NRF-2017R1D1A1B03033087, NRF-2017M3A9F6029736 and NRF-2021R1I1A1A01050838) and Korea Initiative for Fostering University of Research and Innovation (KIURI) Program of the NRF (NRF-2020M3H1A1075314).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author. The data are not publicly available due to data protection legislation.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**

