**Figure 2.** Comparing the synthesis of α-hydroxy-β-keto ester by the BMO (biogenic manganese oxides) from dry m-MnB1 and CMO (chemical manganese oxides): (**a**) Effects of reaction media consisting of H2O/MeCN with different ratios; (**b**) Effects of dosage of BMO and CMO in H2O/MeCN (9:1); (**c**) Correlation of yield of **2** with time; (**d**) Pseudo-first-order plots of α-hydroxylation of β-keto Ester by BMO and CMO in the first 3 h.

Notably, in all tested solvent conditions, BMO in dry m-MnB1 revealed significantly higher reactivity than CMO, and almost 2-fold higher yields were attainable in any given medium (Figure 2a). These results proved that the dry m-MnB1 possesses superior reactivity and selectivity than conventional CMO.

### 2.2.2. Influence of Dosage of Manganese Oxides on Reaction Rate

We subsequently explored the effects of dosage of manganese oxides on the oxidation of compound **1**. A mixture of H2O/MeCN (9:1) was employed as the reaction medium and the reaction time was set at 1 h. As shown in Figure 2b, the ratio of manganese oxides to substrate **1** was found to significantly impact the oxidation process. As the dosage of BMO from dry m-MnB1 was increased from 0.5 eq. to 1.0 eq., the yield of **2** raised from 42% to 72%. The outcome could be further improved to 91% with 2.0 eq of oxidant, and it became marginally higher with further addition of BMO. These results suggest 2.0 eq. of BMO is adequate. As the substrate **1** dissolved in the reaction medium was originally 5.0 mM, the factor limiting it from approaching full conversion might be the extremely low concentration at this point. Analogously, the same set of experiments were conducted with CMO. Although the conversion of substrate **1** was also evidently promoted along the addition of oxidant, the yield of **2** was constantly lower than BMO under identical conditions, for instance, as seen from 56% yield versus 93% yield with 2.5 eq. of manganese oxides. The results also back the notion that the BMO from dry m-MnB1 is more efficient than the CMO for this oxidation reaction.

### 2.2.3. Reaction Kinetics

The kinetic parameters for both BMO and CMO mediated oxidation were studied (Figure 2c,d). It was found that α-hydroxylation of substrate **1** with both BMO from dry m-MnB1 and CMO in

the first 3 h followed Pseudo-first order kinetics, which were expressed as in equation (1), with *C*: concentration of 1 (mmol/L) at time *t* (h), *C*0: the initial concentration of **1** (mmol/L), and *k*: the rate constant (h−1). By transforming Equation (1) into Equation (2),

$$\mathcal{C} = \mathcal{C}\_0 \cdot \mathfrak{e}^{-k \cdot t} \tag{1}$$

$$
\ln\left(\frac{c}{c\_0}\right) = -k \cdot t \tag{2}
$$

the rate constant can be calculated from the slope of the graph as depicted in Figure 2d. An average value for the rate constant *k* of 0.9843 ± 0.0215 h−1 was calculated with BMO from dry m-MnB1, while on the contrary, the rate constant of CMO was merely 0.296 ± 0.0286 h−1 (Table 1). It is clear that the oxidation activity of the BMO from dry m-MnB1 is higher than CMO.

**Table 1.** Kinetic parameters of manganese oxides.

1 BMO: biogenic manganese oxides; 2 CMO: chemical manganese oxides.

### *2.3. E*ff*ects of* β*-Keto Ester 1 on Bacteria Growth*

In order to achieve the α-hydroxylation of **1** by directly using live MnB1 bacteria, we moved on to investigate the influence substrate **1** on the growth of *P. putida* MnB1 at different concentrations. As illustrated in Figure 3a and Figure S4, the MnB1 growth profile did not deviate notably from the standard strain incubation in the presence of 5.0 mM of substrate, while larger concentration (10.0 mM) exhibited obvious toxicity. Further increase of concentration to 15.0 and 20.0 mM both significantly suppressed the bacterial growth. Nevertheless, the bacteria were still proliferating even with 20.0 mM substrate, highlighting the robustness of MnB1.

**Figure 3.** Effects of β-keto ester on the bacteria growth (**a**) measured by spread plate method and (**b**) the manganese mineralization; various colors refer to different concentrations of **1**; CFU: colony forming units.

### *2.4. E*ff*ects of* β*-Keto Ester 1 on Manganese Mineralization*

The effects of various concentrations of **1** on the manganese mineralization by the model strain were also probed. However, the manganese mineralization of MnB1 was found to be much more sensitive to substrate **1**. A concentration of 5.0 mM has already shown a significant inhibition on mineralization process, which was found to be completely repressed at 10.0 mM or higher (Figure 3b). Since the generation of BMO by *P. putida* MnB1 was fundamentally accomplished via multicopper oxidase-catalyzed oxidation of Mn(II) into Mn(VI) in cell extracts within its active center with three Cu(II) sites [34–36], and the β-keto ester is known to chelate divalent metal ions, we speculate that

compound **1** might potentially act as a multicopper oxidase inhibitor [37] to prohibit manganese oxide mineralization. Therefore, a maximum substrate concentration of 5.0 mM was selected for the following whole-cell biocatalytic oxidation of substrate **1**.

### *2.5. Continuous Live MnB1 Catalyzed Synthesis of* α*-Hydroxy-*β*-Keto Ester*

As the whole-cell catalysts have been previously demonstrated as a continuous and repeated-batch reaction system [38], herein we explored such potential of live MnB1 for α-hydroxylation of **1**. The MnB1 bacteria were cultivated in Lept medium containing 1.0 mM MnCl2 until 0.45 mM BMO were formed according to leucoberbelin blue (LBB) test [39]. The substrate **1** was then directly added to the culture medium to reach the concentration of 5.0 mM. During the first round of synthesis, the reaction progress was monitored as shown in Figure 4a. The reaction rate was fast at the initial stage but slowed down with the BMO being consumed. The highest conversion could reach 92% after 24 h (Figure 4b). Afterwards, the MnB1 bacteria were recycled by centrifugation, and washed by PBS before re-culturing in a fresh medium. The same concentration of fresh substrate was added to the medium when the BMO concentration reached 0.45 mM again. The yield of product **2** was consistently determined after 24 h. Notably, in consecutive four cycles, virtually identical yield was attainable for each run, and all were over 90% (92%, 91%, 93% and 94% respectively, Figure 4c). These results clearly suggest that the MnB1-based whole-cells catalyst can be easily regenerated and recycled for continuous reactions, which is highly valuable for low cost and sustainable catalysis and flow chemistry.

**Figure 4.** Synthesis of α-hydroxy-β-keto ester by the whole cells based on the biogenic manganese oxides: (**a**) Schematic diagram of the recycling experiments; (**b**) Profile of substrate consumption and product formation; (**c**) Yield of product **2** in recycling reaction.

### **3. Discussion**

Manganese oxides are known to be generated via oxidation of Mn2+ through both abiotic and biotic channels in the environment. Related reports indicated that the abiotic manganese oxidation pathway is much slower than biological processes, which are performed by a large variety of bacteria and fungi [40,41]. Here, we used *P. putida* MnB1 as the model strain to generate the BMO. Notably, *P. putida* MnB1 is ubiquitous in the freshwater and soil, and its optimal growth temperature is 26~30 ◦C [20], which indicates that the strains are readily available and can be cultivated even in complicated environments. Thus, the robustness of *P. putida* MnB1 lays the groundwork for the prospective industrial application of BMO.

In recent years, BMO have received broad attention due to its marked redox reactivity, for instance, for the degradation of the organic pollutants, removal of heavy metal ions, etc. [42,43]. Although these processes involve various organic transformations, no rational synthesis of organic compounds with specific bond formation using BMO has been disclosed. In this study, we realized the synthesis of valuable α-hydroxy-β-keto ester **2** by utilizing BMO as an oxidant to promote the highly selective C-O bond formation event. Its superior performance than commercially available CMO highlights a strong capacity for synthetic oxidative reactions, and this is consistent with the previous studies on degradation applications [44]. In fact, oxidation of substrate **1** with BMO was a spot-to-spot process based on thin layer chromatography (TLC) analysis, and it was much faster and cleaner than that with CMO. Mechanistically, BMO produced by the *P. putida* MnB1 is known to contain larger inter-layer space and less structural Mn(III) than the CMO [45], thus providing larger specific surface areas [46]. We presume that these characteristics of BMO are beneficial to render defined interaction with substrates, and likely account for their better performance observed in our experiments.

Despite remarkable performance of isolated BMO for the oxidation of β-keto ester **1**, we were further motivated to develop an environmentally friendly and efficient catalytic system by using whole-cell bacteria as a novel type of catalyst. However, tremendous challenges need to be taken into consideration, such as the compatibility with reaction media, the toxicity of substrate to cells and others [47]. Results of solvent optimization (Figure 2a) guided us to take mixture of H2O and MeCN as a suitable reaction medium. However, toxicity experiments showed that higher concentration (10 mM) of substrate would drastically inhibit the mineralization function of bacteria (Figure 3b). Jung [35] and Francis [34] independently concluded that the generation of BMO by *P. putida* MnB1 was fundamentally accomplished by multicopper oxidase, which catalytically oxidizes Mn(II) into Mn(VI) in cell extracts within its active center with three Cu(II) sites. This activity would get inhibited by heating or by treatment with a protease. The β-keto ester is known to chelate divalent metal ions, an essential component in the active site that interacts with amino acid residues of the enzyme. Therefore, this type of substrate might potentially act as a protein inhibitor [37]. We estimated that the higher concentration of β-keto ester **1** suppressed the manganese oxidizing partially due to the toxicity of substrate to the bacteria growth (Figure 3a and Figure S4), and this was probably a result of the impact of substrate on the activity of the multicopper oxidase. Based on these findings, we chose 5 mM as an appropriate concentration of substrate to conduct the whole-cell biocatalysis. As the content of β-keto ester **1** was 5-fold to Mn(II), the obtained 92% yield of product **2** clearly indicates Mn is operative in a catalytic manner, i.e., the regeneration of BMO by *P. putida* MnB1. Although a similar notion has been offered in the degradation of organic pollutants by Ko et al. [18], however, the recyclability of the bacteria for continuous catalysis has never been demonstrated.

In conclusion, we have realized the synthesis of valuable α-hydroxy-β-keto ester (**2**) by utilizing BMO as an oxidant to promote the highly selective C-O bond formation. Its superior performance than commercially available CMO highlights a strong capacity for synthetic oxidative reactions. Consecutive repeated-batch synthesis with recovered bacteria by the whole-cell catalytic system was achieved with consistently high levels of yield recorded [38]. Collectively, the whole-cell catalytic *P. putida* MnB1 with biogenic manganese oxides is highly robust, even amenable to organic transformation with somewhat toxic substrates. As Mn(IV) is extensively utilized in oxidative organic reactions, we anticipate that this efficient biological system is promising for the benign synthesis of various bioactive substances as well as bulk fine chemicals to meet the increasing demand for sustainable chemistry.

### **4. Materials and Methods**

### *4.1. Preparation of Freeze-Dried BMO*

The bacterial strain *Pseudomonas putida* MnB1 [American Type Culture Collection (ATCC) no. 23483] was cultured in the Lept medium (0.5 g/L yeast extract (Ruji, Shanghai, China), 0.5 g/L Casamino Acids (Coolaber, Beijing, China), 5 mM D(+)-glucose (Macklin, Shanghai, China), 10 mM

HEPES (*N*-2-hydroxyethylpiperazine-*N*'-2-ethanesulfonic acid, pH 7.5, GBCBIO, Guangzhou, China), 0.48 mM CaC12 (Macklin), 0.83 mM MgSO4 (Macklin), 3.7 μM FeCl3 (Macklin), and 1 mL of trace element solution (10 mg/L CuSO4•5H2O, 44 mg/L ZnSO4•7H2O, 20 mg CoCl2•6H2O and 13 mg/L Na2MoO4•2H2O, (Macklin)) containing 1mM MnCl2 (Aladdin, Shanghai, China) at 30 ◦C and shaken at 150 rpm for 5 d [48]. The suspensions were centrifuged at 8000× *g* for 20 m and the supernatant was discarded, sediments of bacteria and BMO were diluted three times with deionized water by means of centrifugation (20 m at 8000× *g*), then the mixture of precipitates were collected for freeze-drying to obtain the dried BMO sample.

### *4.2. Quantification of Freeze-Dried BMO by Inductive Coupled Plasma Optical Emission Spectrometry (ICP-OES)*

The 25 μg dry m-MnB1 was treated with 200 μL 90% HNO3 overnight and heated at 70 ◦C for 30 min, then was added H2O2 (30%, 200 μL). The resulting mixture was treated to evaporate at 200 ◦C for 1 h and then dissolved in deionized water to 4 mL solution to be measured by Inductive Coupled Plasma Optical Emission Spectrometry (ICP-OES 7000 Plus, ThermoFisher, Waltham, MA, USA) to quantify the amount of Mn2+. ICP-OES combined with a water cross-flow nebulizer and Ar was run as the carrier gas; auxiliary energy flow, coolant flow, and nebulizer flow were set as 0.7 L/min, 13.00 L/min, and 0.7 L/min, respectively. The analyses were calibrated by gravimetric standards with different concentrations (2 ppm, 4 ppm, 8 ppm, 16 and 32 ppm, respectively) that were measured before sample quantification.

### *4.3. Synthesis of* α*-Hydroxy-*β*-Keto Ester by the Dry m-MnB1 and CMO*

A reaction mixture (400 μL, MeCN: H2O = 1:1) containing 50 mM β-keto ester **1**, 5.0 mg dry m-MnB1 or 1.75 mg CMO was shaken at 150 rpm, 30 ◦C for 3 h. A control experiment was conducted without manganese oxide under otherwise identical reaction conditions. All reaction mixtures were monitored by TLC (hexanes/ethyl acetate = 3:1). Then, the resulting samples were centrifuged at 8000× *g* for 5 m, supernatant was extracted with dichloromethane and evaporated under reduced pressure. The dried residues were purified by silica gel chromatography with hexanes/ethyl acetate (20:1–10:1) to get desired products for the NMR analysis. Finally, the purified samples were used as standards for the following HPLC detection.

### 4.3.1. Optimization of the Reaction Medium

The reaction mixtures (400 μL, MeCN: H2O = 3:1, 1:1, 1:3, 1:6 or 1:9) containing 50 mM β-keto ester (1) (Preparation of this substrate followed a known protocol [49]), 5.0 mg dry m-MnB1 or 1.75 mg CMO (Aladdin) were shaken at 150 rpm, 30 ◦C for 3 h. The reaction samples were centrifuged at 8000× *g* for 5 m and supernatant was analyzed using HPLC to calculate the yield of oxidation product.

### 4.3.2. Assays of Different Dosage of Manganese Oxides on the Oxidation Reaction

Substrate **1** (0.02 mmol) and dry m-MnB1/CMO (Chemical manganese oxides, Innochem, Beijing, China) (0.01, 0.02, 0.03, 0.04, or 0.05 mmol) were added to test tubes and dissolved in MeCN (40 μL) and H2O (360 μL). The reaction samples were stirred at 150 rpm, 30 ◦C for 1.5 h. Then, the reaction systems were centrifuged at 8000× *g* for 5 m and supernatants were analyzed using HPLC to obtain the yield of the oxidation product.

### 4.3.3. Kinetic Measurements of Dry m-MnB1 and CMO

To further compare the reaction efficiency of dry m-MnB1 and CMO, kinetic parameters were detected. The reaction mixtures (**1** (0.02 mmol), dried-BMO/CMO (0.02 mmol), MeCN (40 μL) and H2O (360 μL) were stirred at 150 rpm, 30 ◦C, and the reaction progress was monitored by yield of the oxidation product.

### *4.4. Bioassays of Substrate* β*-Keto Ester Tolerance of Pseudomonas putida MnB1*

The bacterial strain *Pseudomonas putida* MnB1 was precultured overnight in LB medium (10 g/L tryptone, 10 g/L NaCl, 5 g/L yeast extract) at 150 rpm, 30 ◦C and transferred into fresh Lept medium containing 1 mM MnCl2 and various concentrations of **1** (0 mM, 5 mM, 10 mM, 15 and 20 mM, respectively). The colony forming units (CFU) were quantified by using the standard spread plate method [50] every 5 h until the generation of BMO. Simultaneously, the OD600 nm level of bacteria was performed with microplate reader (3020-675, ThermoFisher).

### *4.5. Investigation of the E*ff*ects of Substrate on Manganese Mineralization*

The oxidized Mn content generated in the suspension that was described in the Section 4.1 was measured at the indicated time points by using leucoberbelin blue (LBB, Sigma-Aldrich, St. Louis, MO, USA) method as illustrated by Krumbein [39]. With negligible modifications, bacteria culture (10 μL) was added to 50 μL LBB. After 15 m in the dark at room temperature, deposits were removed by centrifugation. KMnO4 was used as the standard to confirm the absorbance at 620 nm of supernatant.

### *4.6. Continuous Biocatalytic Experiments by Whole Cells Based on BMO*

The bacterial strain *Pseudomonas putida* MnB1 was cultured in the Lept medium containing 1 mM MnCl2 at 150 rpm, 30 ◦C for 5 d to generate the BMO. The concentration of BMO was quantified by LBB method before adding the 5 mM substrate **1** to the medium. The mixture was then continued to shake at 150 rpm, 30 ◦C to get target product and the reaction progress was monitored by HPLC analysis. After the first-round reaction, the bacteria with BMO were collected by centrifugation and diluted three times with deionized water. The precipitates were added to fresh Lept medium containing 1 mM MnCl2 to allow the continuous mineralization of manganese at 30 ◦C and 150 rpm. The mixture was used for the second-round reactions under comparable conditions when the concentration of BMO was similar with the first-round initial content. Four consecutive rounds of biocatalysis by whole-cell were conducted and yield of product in each round was measured. All procedures were performed under rigorous aseptic conditions.

### *4.7. Methods and Conditions of HPLC Analysis*

For HPLC detection, all of the collected supernatant samples were filtered over a 0.22 μm filter (Millipore, Billerica, MA, USA) and analyzed by HPLC system containing a LC-UV 100 absorbance detector (Wufeng, Shanghai, China). The compounds were separated on a reverse phase HPLC C18 column (C18 250 × 4.6 mm, 5 μm, Shodex, Tokyo, Japan) at a constant flow rate of 1.0 mL/min and analysed by UV/Vis detection at 254 nm. Solvent A was deionized water containing 5% trifluoroacetic acid (TFA, Aladdin) and solvent B was acetonitrile containing 5% TFA. A gradient from 0% to 70% solvent B was applied from 0 to 5 min, the solvent B decreased 70% to 55% from 5 to 20 m, then dropped to 5% during 5 m. All analyses were calibrated by the absorption of substrate and product standards with different concentrations.

### *4.8. Statistical Analysis*

All data were presented as means ± standard deviation of three replicates. Statistical analysis was completed using SPSS 16.0 (Statistical Package for the Social Sciences, IBM Corp., Armonk, NY, USA).

### **Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/21/5/1709/ s1.

**Author Contributions:** Conceptualization, J.L., Y.W. and F.Z.; methodology, J.G., H.G. and J.L.; data curation, J.G. and J.L.; investigation, J.G. and H.G.; writing—original draft preparation, J.G.; writing—review and editing, Y.W. and F.Z.; supervision, Y.W. and F.Z.; funding acquisition, J.L. and Y.W. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the National Key R&D Program of China (2018YFA0903500), the China Postdoctoral Science Foundation funded project (2018M630849), the 1000 Young Talent Program of China, and Hubei Technological Innovation Project (2019ACA125).

**Acknowledgments:** We thank the Analytical and Testing Centre of HUST, Analytical and Testing Centre of School of Chemistry and Chemical Engineering (HUST), and Research Core Facilities for Life Sciences (HUST) for instrument support.

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

### **Abbreviations**

### **References**

© 2020 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Improving the Performance of Horseradish Peroxidase by Site-Directed Mutagenesis**

### **Diana Humer and Oliver Spadiut \***

TU Wien, Institute of Chemical, Environmental and Bioscience Engineering, Research Area Biochemical Engineering, Gumpendorfer Straße 1a, 1060 Vienna, Austria<EMAIL_ADDRESS>
**\*** Correspondence<EMAIL_ADDRESS>
Received: 23 January 2019; Accepted: 16 February 2019; Published: 20 February 2019

**Abstract:** Horseradish peroxidase (HRP) is an intensely studied enzyme with a wide range of commercial applications. Traditionally, HRP is extracted from plant; however, recombinant HRP (rHRP) production is a promising alternative. Here, non-glycosylated rHRP was produced in *Escherichia coli* as a DsbA fusion protein including a Dsb signal sequence for translocation to the periplasm and a His tag for purification. The missing *N*-glycosylation results in reduced catalytic activity and thermal stability, therefore enzyme engineering was used to improve these characteristics. The amino acids at four *N*-glycosylation sites, namely N13, N57, N255 and N268, were mutated by site-directed mutagenesis and combined to double, triple and quadruple enzyme variants. Subsequently, the rHRP fusion proteins were purified by immobilized metal affinity chromatography (IMAC) and biochemically characterized. We found that the quadruple mutant rHRP N13D/N57S/N255D/N268D showed 2-fold higher thermostability and 8-fold increased catalytic activity with 2,2'-azino-*bis*(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) as reducing substrate when compared to the non-mutated rHRP benchmark enzyme.

**Keywords:** *E. coli*; recombinant horseradish peroxidase; site-directed mutagenesis; periplasm; glycosylation sites

### **1. Introduction**

The enzyme horseradish peroxidase (EC <IP_ADDRESS>) is a heme-containing oxidoreductase used in both research and diagnostics for a wide range of applications (e.g., immunoassays, diagnostic kits, probe-based assay techniques as ELISA, EMSA, Western blotting and Southern blotting, waste water treatment and as a reagent in organic synthesis [1–7]. This 308 amino acid metalloenzyme incorporates two calcium atoms and four disulphide bridges [8,9]. In the plant, HRP also contains a hydrophobic 30 amino acid N-terminal leader sequence and a 15 amino acid C-terminal extension [10]. The C-terminal sequence is a sorting signal responsible for secretion to the vacuole [11] and the N-terminal sequence directs the enzyme to the endoplasmatic reticulum (ER) for post translational modifications, namely heme and calcium incorporation, disulphide bond formation and *N*-glycosylation [12]. The asparagine-linked glycans represent about a fifth of the 44 kDa plant holoenzyme. HRP possesses nine potential glycosylation sites and the pattern as well as the occupation of these sites is heterogeneous between HRP variants [13,14]. At least 28 different native HRP isoforms have been described so far [15] out of which HRP C is the most abundant and therefore the most studied one [9].

The commercially available HRP is extracted from *Armoracia rusticana* roots. However, only seasonal availability and long cultivation times paired with low yields make the classical production process undesirable. Moreover, the content of single isoenzymes is often very low and downstream processing is tedious. As a consequence, there is a need for a uniform enzyme preparation with defined characteristics and recombinant protein production would mitigate this issue. Many studies have addressed *Saccharomyces cerevisiae* or *Pichia pastoris* as host organisms for rHRP production because

yeasts are easy to cultivate and commonly used for glycosylated and disulphide bond containing proteins. Alas, in yeast hyper-glycosylation occurs and the downstream process is cumbersome. Prokaryotes on the other hand lack the organelles necessary for glycosylation, namely ER and Golgi apparatus. In addition, the reducing environment in the cytoplasm of bacteria impedes the formation of disulphide bridges. Hence, recombinant glycoproteins with disulphide bridges are usually not produced in bacteria. However, *E. coli* is a very convenient host organism because of its cheap and easy cultivation at high cell densities. Moreover, there are no obstacles due to hyper-glycosylation as it is the case in yeast. This substantially facilitates downstream processing and allows application of the product for therapeutic use. It has also been shown that glycosylation is not essential for HRP activity or folding [16] although enzyme activity and thermal stability are considerably reduced when compared to the native enzyme [15,17]. Many studies have already been performed with the goal to enhance the general stability and activity properties of rHRP (Table 1).

**Table 1.** List of rHRP mutations that improve enzyme performance, listed by authors.

**Table 1.** *Cont.*

\* 13A7 = A85(GCC→GCT)/N212D/Q223L; \* H2-10G5 = A85(GCC→GCT)/N175S/N212D; \* 13A7-N175S = A85(GCC→GCT)/N212D/Q223L/N175S; \* 13A10 = R93L/T102A/L131P/N135(AAC→AAT)/L223Q/T257 (ACT→ACA)/V303E; \* 17E12 = N47S/T102A/G121(GGT→GGC)/L131P/N135(AAC→AAT)/L223Q/P226Q/T257 (ACT→ACA)/P289(CCT→CCA).

Lin et al. [21] identified a N255D mutant by random mutagenesis with 14-fold higher activity than the non-mutated benchmark enzyme but they concluded that this increase was due to better folding of the enzyme rather than improved catalytic performance. Directed evolution was used to identify mutants 13A10 and 17E12 (for mutant descriptions see Table 1, Footnotes) in *P. pastoris,* which were associated with increased specific activity with ABTS (5.4-fold and 2.8-fold) and guaiacol (2.4-fold and 1.2-fold) as substrates. The thermostability of 13A10 was comparable to the non-mutated benchmark enzyme but it was remarkably decreased in 17E12 mutants [27]. Variant 13A10 was used as starting point for successive rounds of directed evolution and gave rise to 13A7, H2-10G5 and 13A7-N175S (for mutant descriptions see Table 1, Footnotes). These variants were found to be more stable towards pH, temperature, SDS, urea and sodium chloride but enzyme activity was not further improved [26]. Ryan et al. [23] intensely studied the influence of site-directed mutations on hydrogen peroxide tolerance. They identified T110V, K232N and K241F, which were 25-, 18- and 12-fold more resistant towards hydrogen peroxide than the non-mutated benchmark enzyme. These variants also showed increased tolerance to heat and solvents. In addition, K232N as well as K241F displayed higher turnover numbers (kcat) with ABTS as reducing substrate [22]. Asad et al. [18] changed the amino acids present at two *N*-glycosylation sites of rHRP by site-directed mutagenesis. They described variants N13D and

N268D, which showed increased catalytic efficiency with phenol/4-aminoantipyrine and were both more stable in terms of hydrogen peroxide and heat tolerance. A follow-up study identified N268G, which showed 18-fold higher resistance towards hydrogen peroxide and 2.5-fold higher thermal stability [20]. Capone et al. [19] performed a profound investigation of *N*-glycosylation mutants in *P. pastoris*, where asparagines at eight sites were replaced by aspartic acid, serine or glutamine. They showed that the positive influence of N13D and N268D on thermal stability is also valid for expression in yeast and that the variant identified by Lin et al. [21] was apparently slightly beneficial in terms of catalytic activity. Interestingly, a variant with mutations at all eight *N*-glycosylation sites showed substantially decreased activity and thermal stability.

At the moment, HRP is not used for in vivo medical applications, because the plant glycosylation pattern differs significantly from human glycoforms and therefore has immunogenic potential [28]. This can be circumvented by reducing the glycosylation pattern to mannose-type glycans, which can be achieved by adding an ER retention sequence. Unfortunately, these glycans lead to rapid clearance from circulation in humans [28]. Nevertheless, a combination of HRP and paracetamol or the plant hormone indole-3-acetic acid (IAA) was found to be medically active in targeted cancer treatment [29,30]. Although this cytotoxic effect has been known since the nineties [31,32], up to now, HRP is not considered suitable for therapeutic use. For this application, a recombinantly produced single isoform free of glycosylation with sufficient stability and activity would be desirable. In this study, we investigated the *N*-glycosylation mutants N13D, N57S, N255D, N268D and combinations thereof for thermal stability and catalytic efficiency with the substrate ABTS for rHRP expressed in *E. coli.* Soluble rHRP was preferred for mutant screenings because refolding of rHRP from *E. coli* inclusion bodies is a complex and cumbersome procedure which still has to be optimized. Therefore, we chose an expression system that leads to translocation of rHRP into the periplasm. The aim of this work was to improve the traits of non-glycosylated rHRP towards higher stability and catalytic efficiency to increase suitability for medical applications. Indeed, a promising rHRP *N*-glycosylation mutant was identified and biochemically characterized.

### **2. Results and Discussion**

### *2.1. Protein Production*

Recombinant HRP variants were produced and then translocated to the periplasm by the DsbA signal sequence. Soluble proteins were isolated from the periplasm and rHRP was purified to gain active, correctly folded enzyme. The imidazole concentration in the IMAC binding buffer was at the upper limit given by the column manufacturer (GE Healthcare, Chicago, IL, USA) to avoid unspecific interactions between *E. coli* host cell proteins and the stationary IMAC phase. Nevertheless, several impurities were visible on the SDS PAGE of the IMAC eluate (data not shown). Therefore, rHRP concentrations were calculated using an SDS-PAGE HRP standard curve with known concentrations (Figure S1). The peak area of rHRP was determined using Fiji Image Analysis Software (https: //fiji.sc) [33] and the protein content was calculated using the slope of the linear regression line of known rHRP concentrations. This led to final enzyme titres of 0.05–0.09 mg rHRP g−1 DCW. The final rHRP product yield was between 0.04–0.08 g L−1 and is similar to reported values from Gundinger et al. [17] for soluble rHRP in pET39b+ (0.048 g L−1).

### *2.2. Biochemical Characterization*

### 2.2.1. Biochemical Characterization of Benchmark rHRP and Seven rHRP Mutants

### <IP_ADDRESS>. Enzyme Kinetics

Plant HRP VI-A (Sigma-Aldrich, St. Louis, MO, USA), non-mutated benchmark rHRP and the seven rHRP variants N13D, N57S, N255D, N268D, N57S/N268D, N57S/N255D/N268D and N13D/N57S/N255D/N268D were analysed for steady-state kinetics with ABTS as reducing substrate. The kinetic constants are presented in Table 2. N13D and N255D showed less catalytic efficiency (Kcat/Km) than the benchmark enzyme and for N13D this is in accordance with Capone et al. [19]. For N255D on the other hand, Capone et al. [19] observed almost the same catalytic activity compared to the benchmark enzyme (1.1-fold increase). N268D had a 2-fold increased turnover number (kcat) when compared to the non-mutated rHRP and the same trend was shown by Asad et al. [20], where a 2.6-fold enhanced kcat with phenol/4-aminoantipyrine was reported. The slightly enhanced catalytic efficiency of N57S, when compared to the benchmark rHRP (1.2-fold), is in accordance with Capone et al. [19] (1.4-fold). The triple mutant N57S/N255D/N268D reached a 3.2-fold higher turnover number than the benchmark rHRP and with a 10-fold increase, N13D/N57S/N255D/N268D showed the highest fold change of kcat when compared to the benchmark rHRP. However, only N57S and N13D/N57S/N255D/N268D had an increased catalytic efficiency when compared to the non-mutated rHRP (1.2-fold and 2-fold, respectively). In general, the results were greatly affected by the unusually high Michaelis Menten constants (Km) and the considerable standard deviations (see Section 2.2.2).

**Table 2.** Kinetic characteristics of plant HRP, rHRP and seven rHRP variants with ABTS as reducing substrate measured in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl.

### <IP_ADDRESS>. Thermal Stability

The thermal stability of plant HRP VI-A, non-mutated benchmark rHRP and the seven rHRP variants was determined at 60 ◦C (Table 3). N13D and N268D were found to enhance stability towards heat, which is in accordance with Asad et al. [18] and Capone et al. [19]. However, Capone et al. [19] also reported a positive effect of N57S, which could not be confirmed. Variant N255D seemed to have no effect on thermal stability. The double mutant N57S/N268D was similar to the rHRP benchmark enzyme concerning temperature susceptibility, so apparently the effect of the mutations is not additive. N268D had the highest benefit and was 3.6-fold more stable than the non-mutated rHRP, whereas the quadruple mutant N13D/N57S/N255D/N268D was 2.4-fold enhanced. Although N268D was the most thermostable variant, the quadruple mutant was chosen for further investigations, as this variant showed promising results for both catalytic activity and thermal stability.

**Table 3.** Half-life of plant HRP, rHRP and seven rHRP variants at 60 ◦C in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl.

*t***1/2** = half life.

### 2.2.2. Catalytic Activity of Plant HRP under Different Conditions

Interestingly, all obtained Michaelis Menten constants (Table 2) were much higher than previously reported for soluble rHRP variants [17] and this was also the case for commercially available plant HRP [17,34,35]. The substantial variability of the obtained Michaelis Menten constants may be a direct result of the high Km values we observed due to fast reactions and high slopes, as variability increases with reaction velocity (see Table 2). We assumed that the measurement buffer was influencing the results of the kinetic measurements, as this was the only apparent difference to the previous procedure from Gundinger et al. [17]. Therefore, the catalytic activity of plant HRP was determined in both buffers: 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl and 50 mM KH2PO4 pH 6 (Table 4). The results indicated that the high Michaelis Menten constants, as well as the high standard deviations, are indeed buffer dependent. The Km value was 6-fold higher in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl buffer when compared to 50 mM KH2PO4 pH 6. As a consequence, the catalytic efficiency was 9-fold enhanced when the measurements were performed in potassium phosphate buffer. The kinetic parameters observed with 50 mM KH2PO4 pH 6 are in accordance with Gundinger et al. [17], as they reported a Km-value of 1.75 mM and a Vmax-value of 567 U mg−1.

**Table 4.** Comparison of kinetic characteristics measured with plant HRP Type VI-A in two different buffers.

However, at this point it was still unclear whether these results were obtained because of the buffer substance, the pH change or the additives in the BisTris buffer. Therefore, the specific activity of plant HRP with ABTS as reducing substrate was examined under different conditions (Figure 1). Apparently, the additives glycerol and sodium chloride have a negative influence, as the specific activity was 2-fold enhanced when they were omitted. The two buffers 50 mM KH2PO4 pH 7 and 50 mM BisTris/HCl pH 7 led to comparable results. The catalytic activity at pH 5 was 3-fold enhanced when compared to pH 7 when the measurements were conducted in potassium phosphate buffer. Based on these results, we decided to use 50 mM KH2PO4 pH 5 for all consecutive measurements (Section 2.2.3).

**Figure 1.** Specific activity of plant HRP Type VI-A with ABTS under various conditions. Five different buffers were used to measure the catalytic activity of plant HRP with 5 mM ABTS and 1 mM hydrogen peroxide. Horizontal stripes: 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl; white: 50 mM BisTris/HCl pH 7; light grey: 50 mM KH2PO4 pH 5; vertical stripes: 50 mM KH2PO4 pH 6.5; dark grey: 50 mM KH2PO4 pH 7.

### 2.2.3. Optimized Biochemical Characterization of Benchmark rHRP and Mutant N13D/N57S/N255D/N268D

The first investigation (Section 2.2.1) showed that the quadruple mutant N13D/N57S/N255D/N268D was the most promising variant when compared to the non-mutated benchmark rHRP (Tables 2 and 3). Therefore, a second protein purification and biochemical characterization with optimized assay conditions was performed to confirm these results.

### <IP_ADDRESS>. Enzyme Activity

Enzyme kinetic measurements with plant HRP, non-mutated benchmark rHRP and rHRP variant N13D/N57S/N255D/N268D were performed in 50 mM KH2PO4 pH 5 using 96-well plates. Here, N13D/N57S/N255D/N268D showed an 8-fold enhanced catalytic efficiency (Kcat/Km) and an 8-fold increased turnover number (kcat) when compared to the benchmark enzyme (Table 5). The increase in kcat for N13D/N57S/N255D/N268D was similar to the results obtained with the previous assay (Table 2, 10-fold). Besides, the Km (0.27 mM) for plant HRP was similar to values reported in literature: 0.27 mM [35] and 0.11 mM [34]. However, the Km-values observed for plant HRP in potassium phosphate buffer differed significantly between the two assays (Tables 4 and 5). We assume that this might be due to differences in pH, as the Michaelis Menten constant in 50 mM KH2PO4 pH 6 was 1.5 mM (Table 4) and Gundinger et al. [17] reported 1.75 mM in 50 mM KH2PO4 pH 6.5. Gilfoyle et al. [35] and Grigorenko et al. [34] used sodium phosphate/citrate buffer at pH 5 and sodium acetate buffer at pH 5, respectively, which resulted in Km-values of 0.27 mM [35] and 0.11 mM [34] which are in accordance with 0.27 mM in potassium phosphate (Table 5).

**Table 5.** Kinetic characteristics of plant HRP, rHRP and N13D/N57S/N255D/N268D with ABTS as reducing substrate in 50 mM KH2PO4 pH 5.

### <IP_ADDRESS>. Thermal Stability

The thermal stability of plant HRP VI-A, non-mutated benchmark rHRP and rHRP variant N13D/N57S/N255D/N268D was determined at 60 ◦C (Table 6). The measurement was performed at pH 7 to guarantee comparability with the previous assay (Section <IP_ADDRESS>). Interestingly, the half-life of all HRP species measured in 50 mM KH2PO4 buffer pH 7 was slightly higher than the half-life in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl (Table 4). Asad et al. [36] and Haifeng et al. [37] found that phosphate buffers can influence the thermostability of horseradish peroxidase. Although the samples were kept in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl during heat exposition in both experiments, it might still be possible that potassium phosphate had a stabilizing effect during the activity measurement. In both experiments the plant HRP was far more stable than the rHRP variants (Tables 3 and 6) because glycans improve enzyme stability [18]. The thermostability of the quadruple mutant N13D/N57S/N255D/N268D was 2-fold higher than the stability of the benchmark rHRP enzyme (Table 6) which is in accordance with the previous assay (Table 3; 2.4-fold).

**Table 6.** Half-life of plant HRP, rHRP and N13D/N57S/N255D/N268D at 60 ◦C in 50 mM KH2PO4 pH 7.

Summarizing, the quadruple mutant showed a significantly augmented performance concerning thermal stability as well as catalytic activity with ABTS as substrate. Therefore, this variant is considered a good starting point for further enzyme engineering approaches.

### **3. Materials and Methods**

### *3.1. Chemicals*

Chemicals were purchased from Carl Roth (Karlsruhe, Germany) or AppliChem (Darmstadt, Germany). Plant HRP Type VI-A (Cat. No.: P6782) was purchased from Sigma-Aldrich (St. Louis, MO, USA), enzymes were purchased from New England Biolabs (Ipswich, MA, USA) or Thermo Fisher Scientific Inc. (Waltham, MA, USA), 2,2'-azino-*bis*(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) was purchased from Sigma-Aldrich or AMRESCO® biochemical (Solon, OH, USA).

### *3.2. Strains and Plasmids*

The *hrp* gene coding for HRP variant C1A was codon-optimized for *E. coli* and obtained from GenSript USA Inc. (Piscataway, NJ, USA). HRP was produced as a His6-tagged recombinant protein from pET39b+ in the *E. coli* strain BL21 (DE3) (Lucigen, Middleton, WI, USA). The plasmid pET39b+ (Novagen, San Diego, CA, USA) encodes a Dsb tag for export and periplasmic folding, so that a DsbA fusion protein with a HIS tag between the *dsbA* sequence and the *hrp* sequence is generated.

### *3.3. Strain Generation by Site-Directed Mutagenesis*

The following plasmids were constructed with standard molecular cloning techniques [38]. Whole plasmid PCR was used to introduce mutations in the *hrp* gene by site-directed mutagenesis. The 7 kb fragment was amplified with the respective oligonucleotides to generate single, double, triple and quadruple mutations (Table 7). All oligonucleotides were purchased from Microsynth (Balgach, Switzerland). Each PCR reaction contained 1× Q5 Reaction Buffer, 200 μM dNTP Mix, 200 nM of both forward and reverse primer, 100 ng template vector DNA and 1 U Q5 High-Fidelity DNA Polymerase. The PCR products were purified with the Monarch PCR & DNA Cleanup Kit from New England Biolabs (NEB, Ipswich, MA, USA) and the template plasmid DNA was removed by FastDigest *Dpn*I (Thermo Scientific™, Waltham, MA, USA) digestion. 1 FDU (FastDigest unit, see Abbreviations) of *Dpn*I was added to the cleaned PCR products and incubated overnight at 37 ◦C. After heat inactivation at 80 ◦C for 20 min, the plasmids were transformed into BL21 (DE3). All DNA inserts of the recombinant plasmids were verified by DNA sequencing (Microsynth, Balgach, Switzerland).

The mutated sites are underlined. fwd: forward; rev: reverse.

### *3.4. Growth Conditions and Protein Production*

LB medium (10 g L−1 tryptone, 10 g L−1 NaCl and5gL−1 yeast extract) or SB medium (32 g L−1 tryptone; 20 g L−1 yeast extract; 5 g L−1 NaCl; 5 mM NaOH) was used for cultivation of BL21 (DE3) strains. Kanamycin was added to a final concentration of 50 mg L−1 to ensure plasmid maintenance. Pre-cultures were grown overnight at 37 ◦C with shaking (250 rpm) in 50 mL LBKan or SBKan medium and 2.5 L Ultra Yield Flasks (UYF) were inoculated to reach an optical density (OD600) of 0.3 in a final volume of 500 mL LBKan or SBKan medium. The cells were grown at 37 ◦C with shaking (250 rpm) until an OD600 of 0.5, subsequently *hrp* expression was induced by adding 0.1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG). After growth for 20 h at 25 ◦C and 250 rpm, the cells were harvested by centrifugation (4500 rpm, 30 min, 4 ◦C).

### *3.5. Protein Purification*

The cell pellets were resuspended in buffer A (50 mM BisTris/HCl pH 7, 500 mM NaCl, 40 mM imidazole) with cOmplete™ Protease Inhibitor Cocktail (Roche, Basel, Switzerland). The cell suspension was homogenized with an Avestin Emulsiflex C3 high pressure homogenizer (Avestin, Ottawa, ON, Canada) for 10 passages at 1000 bar and centrifuged afterwards at 10,000 rpm for 1 h. Particles were removed from the supernatant by filtration (0.2 μm) prior to protein purification with IMAC using the Äkta pure system (GE Healthcare, Chicago, IL, USA). The column (HisTrap™ Fast Flow Crude 1 mL, GE Healthcare) was equilibrated with 10 column volumes (CV) buffer A and the crude extract was loaded with a linear flow rate of 156 cm h−1. Subsequently, the column was washed with 10–20 CV buffer A before step elution with 100% buffer B (50 mM BisTris/HCl pH 7, 500 mM NaCl, 500 mM imidazole) at a linear flow rate of 156 cm h−1. The eluted protein fractions were desalted with Sephadex G-25 PD-10 desalting columns (GE Healthcare, Chicago, IL, USA) and eluted in buffer C (50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl). Glycerol and sodium chloride were added to the buffer as these substances were found to positively impact on rHRP stability (data not shown). Total protein content was measured using the Bradford assay [39] and the eluate was analysed by SDS PAGE. The concentration of rHRP was calculated as μM mL−1 using 61.5 kDa (with DsbA Protein and His tag) as molecular mass of the fusion protein. Hemin and calcium chloride were added in a 2-fold and 4-fold molar amount, respectively and the enzyme preparations were incubated over night at 4 ◦C with slight agitation.

### *3.6. SDS PAGE*

SDS PAGE was performed according to the Laemmli protocol [40]. Mini-PROTEAN® TGX Stain-Free™ Precast Gels (Bio-Rad, Hercules, CA, USA) were used and the gel was run with SDS running buffer (25 mM Tris, 200 mM glycine, 0.1% SDS) in a Bio-Rad Mini-PROTEAN® Tetra Cell. Proteins were separated at 125 V for 1 h and the bands were visualized with Coomassie Brilliant Blue solution. Bio-Rad Precision Plus Protein™ Dual Xtra Prestained Protein Standard or Thermo Scientific PageRuler™ Plus Prestained Protein Ladder, 10 to 250 kDa (Waltham, MA, USA) were used as mass standards. The SDS PAGE was analysed using a ChemiDoc™ MP System with Image Lab™ Software (Bio-Rad, Hercules, CA, USA).

### *3.7. Biochemical Enzyme Characterization*

### 3.7.1. Assay Development

### <IP_ADDRESS>. Pathlength Correction for Microplates

The pathlength for 200 μL reaction volume was determined experimentally by measuring the absorbance of MilliQ water in the Tecan Infinite M200 PRO (Tecan, Männedorf, Switzerland) and the Hitachi U-2900 spectrophotometer (Hitachi, Tokyo, Japan). Water absorption can be measured at near infrared wavelength with a maximum absorbance at 975 nm, the measurement at 900 nm subtracts background absorbance (e.g., plastic of 96-well plate). The correct pathlength for the 96-well plates was then calculated according to Equation (1):

$$pathlength = \frac{A975 \text{ nm (well)} - A900 \text{ nm (well)}}{A975 \text{ nm (cuvette)} - A900 \text{ nm (cuvette)}} \times 10 \text{ mm} \tag{1}$$

Thus, a reaction volume of 200 μL results in a pathlength of 0.58 cm in 96-well plates.

### <IP_ADDRESS>. Determination of Extinction Coefficient for ABTS

The extinction coefficient of the ABTS radical was determined experimentally with a Tecan Infinite M200 PRO system (Tecan, Männedorf, Switzerland) for ABTS purchased from Sigma-Aldrich (St. Louis, MO, USA) and AMRESCO® biochemical (Solon, OH, USA). ABTS concentrations ranging from 0 mM to 0.15 mM were measured in 50 mM KH2PO4 pH 5 buffer with 1 mM hydrogen peroxide in a final volume of 200 μL and 22.5 ng plant HRP Type VI-A (Sigma-Aldrich, Cat. No.: P6782). The reaction was followed to the plateau phase in a 96-well plate at 420 nm for 40 min at 30 ◦C. Finally, the extinction coefficient was calculated from the slope of the linear regression line of ABTS concentration plotted against maximal absorbance. The extinction coefficient of ABTS at 420 nm is 27 mM−1 cm−1 for ABTS purchased from both companies (Sigma-Aldrich: y = 15.455x + 0.0548; R2 = 0.9977/ AMRESCO® biochemical: y = 15.455x + 0.0314; R2 = 0.9986).

### <IP_ADDRESS>. Enzyme Activity Calculation

HRP activity in Units mL−1 (1 Unit is defined as the amount of enzyme which oxidizes 1 μmol of ABTS per minute) was calculated according to Equation (2):

$$\frac{\text{dL}}{\text{mL}} = \frac{(\Delta A 420 \text{ nm} - \Delta A 420 \text{ nm} \text{ blank}) \times \text{total volume of reaction} \times \text{d}}{\varepsilon \times \text{p} \times \text{sample volume}} \tag{2}$$

*d* = dilution factor

p = pathlength in cm, which is 1 for cuvettes and 0.58 in a 96-well plate when the reaction volume is 200 μL (see Section <IP_ADDRESS>).

ε = 27 mM−1 cm−1

### 3.7.2. Catalytic Activity of Plant HRP under Different Conditions

Enzyme activity of plant HRP Type VI-A (Sigma-Aldrich, Cat. No.: P6782, St. Louis, MO, USA) was determined either in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl or in 50 mM KH2PO4 pH 6. The reaction mixture in the cuvette contained 27.6 ng HRP, a saturating hydrogen peroxide concentration of 1 mM and varying ABTS concentrations (0.1–10 mM) in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl or in 50 mM KH2PO4 pH 6 with a final volume of l mL. The increase in absorption was followed at 420 nm for 180 s at 30 ◦C in a Hitachi U-2900 spectrophotometer (Hitachi, Tokyo, Japan). All measurements were performed in triplicates. Kinetic parameters were calculated using OriginPro software (OriginLab Corporation 2016, Northampton, MA, USA). As these measurements revealed substantial differences concerning the kinetic parameters between the two buffers, it was decided to further investigate the influence of buffer substance, pH and buffer additives in a 96-well plate format. Here, the reaction mixture in each well contained a saturating hydrogen peroxide concentration of 1 mM and 5 mM ABTS in a final volume of 200 μL. Commercially available plant HRP (9 ng; Sigma-Aldrich, Cat. No.: P6782, St. Louis, MO, USA) was added to the reaction mixture and the increase in absorption was followed at 420 nm for 180 s at 30 ◦C in a Tecan Infinite M200 PRO instrument (Tecan, Männedorf, Switzerland). The following buffers were used: 50 mM KH2PO4 pH 5; 50 mM KH2PO4 pH 6.5; 50 mM KH2PO4 pH 7; 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl or 50 mM BisTris/HCl pH 7. All measurements were performed in duplicates.

### 3.7.3. Enzyme Kinetics

Enzyme kinetic parameters for the substrate ABTS were determined with non-mutated rHRP benchmark enzyme, seven rHRP mutants and commercially available plant HRP Type VI-A. The samples were either measured in cuvettes with a U-2900 spectrophotometer (Hitachi, Tokyo, Japan) or in 96-well plates using a Tecan Infinite M200 PRO instrument (Tecan, Männedorf, Switzerland).

### <IP_ADDRESS>. Hitachi U-2900 Spectrophotometer

The reaction mixture in the cuvette contained 20 μL protein sample, a saturating hydrogen peroxide concentration of 1 mM and varying ABTS concentrations (0.1–10 mM) in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl buffer with a final volume of l mL. The increase in absorption was followed at 420 nm for 180 s at 30 ◦C in a Hitachi U-2900 spectrophotometer. All measurements were performed in triplicates. The kinetic parameters were calculated using OriginPro software (OriginLab Corporation 2016, Northampton, MA, USA).

### <IP_ADDRESS>. Tecan Infinite M200 PRO

The reaction mixture in each well of the 96-well plate contained a saturating hydrogen peroxide concentration of 1 mM and varying ABTS concentrations (0.1–10 mM) in 50 mM KH2PO4 pH 5 buffer in a final volume of 200 μL. Protein sample (5 μL) was added to each well and was filled up with 195 μL reaction mixture. The increase in absorption was followed in a 96-well plate at 420 nm for 180 s at 30 ◦C in a Tecan Infinite M200 PRO instrument. Eight replicates were used for all measurements. The kinetic parameters were calculated using OriginPro software (OriginLab Corporation 2016, Northampton, MA, USA).

### 3.7.4. Thermal Stability

The thermal stability of the enzyme was assessed at 60 ◦C in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl and the residual activity with ABTS was measured after 0, 1, 2, 3 and 5 min for the non-mutated rHRP benchmark enzyme and the seven rHRP mutants. The residual activity of plant HRP Type VI-A (Sigma-Aldrich, Cat. No.: P6782, St. Louis, MO, USA) was measured after 0, 30, 60, 90 and 120 min, respectively. The protein concentration was normalized to 0.05 g L−1 for all samples to minimize potential influences of protein concentrations on thermal stability. The reaction mixture contained 20 μL of protein, a saturating hydrogen peroxide concentration of 1 mM and 10 mM ABTS in 50 mM BisTris/HCl pH 7, 7% glycerol, 100 mM NaCl or in 50 mM KH2PO4 pH 7 with a final volume of l mL. The increase in absorption was followed at 420 nm for 180 s at 30 ◦C in a Hitachi U-2900 spectrophotometer (Hitachi, Tokyo, Japan). The residual enzyme activity was plotted against incubation time and the half-life at 60 ◦C was calculated using the rate of inactivation in Equation (3):

$$t\_{1/2} = \frac{lm2}{k\_{in}}$$

*t*1/2 = half life *kin* = slope of the logarithmic residual activity

### **4. Conclusions**

HRP has many features that make it suitable for therapeutic use: it is stable at 37 ◦C, shows high activity at physiological pH and can be conjugated to antibodies or lectins [9]. Thus, it is highly interesting to engineer rHRP for increased activity and stability for use in medical applications. In our study we discovered a new non-glycosylated rHRP variant with improved characteristics by site-directed mutagenesis of amino acids at the *N*-glycosylation sites. N13D/N57S/N255D/N268D was found to substantially increase activity with ABTS as substrate, the catalytic efficiency of this variant was 8-fold higher when compared to the rHRP benchmark enzyme. Moreover, N13D/N57S/N255D/N268D is

2-fold more stable towards high temperature exposure than the non-mutated rHRP. Currently, we work on further improvement of N13D/N57S/N255D/N268D by directed evolution, as well as the additional introduction of selected mutants (see Table 1). Finally, our future goal will be to produce the resulting non-glycosylated rHRP variant in a large-scale inclusion body process.

**Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/20/4/ 916/s1.

**Author Contributions:** Conceptualization, O.S.; Methodology, O.S. and D.H.; Validation, O.S. and D.H.; Formal analysis, D.H.; Investigation, D.H.; Resources, O.S.; Data curation, D.H.; Writing—original draft preparation, D.H.; Writing—review and editing, O.S. and D.H.; Visualization, D.H.; Supervision, O.S.; Project administration, O.S.; Funding acquisition, O.S.

**Funding:** This research was funded by Open Access Funding by the Austrian Science Fund (FWF), grant number P30872-B26.

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

### **Abbreviations**

### **References**

© 2019 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **2-Ketoglutarate-Generated In Vitro Enzymatic Biosystem Facilitates Fe(II)**/**2-Ketoglutarate-Dependent Dioxygenase-Mediated C–H Bond Oxidation for (2**s**,3**r**,4**s**)-4-Hydroxyisoleucine Synthesis**

**Xiao-Ran Jing 1, Huan Liu 1, Yao Nie 1,\* and Yan Xu 1,2,\***

Received: 24 June 2020; Accepted: 20 July 2020; Published: 28 July 2020

**Abstract:** Fe(II)/2-ketoglutarate-dependent dioxygenase (Fe(II)/2-KG DO)-mediated hydroxylation is a critical type of C–H bond functionalization for synthesizing hydroxy amino acids used as pharmaceutical raw materials and precursors. However, DO activity requires 2-ketoglutarate (2-KG), lack of which reduces the efficiency of Fe(II)/2-KG DO-mediated hydroxylation. Here, we conducted multi-enzymatic syntheses of hydroxy amino acids. Using (2s,3r,4s)-4-hydroxyisoleucine (4-HIL) as a model product, we coupled regio- and stereo-selective hydroxylation of l-Ile by the dioxygenase IDO with 2-KG generation from readily available l-Glu by l-glutamate oxidase (LGOX) and catalase (CAT). In the one-pot system, H2O2 significantly inhibited IDO activity and elevated Fe2+ concentrations of severely repressed LGOX. A sequential cascade reaction was preferable to a single-step process as CAT in the former system hydrolyzed H2O2. We obtained 465 mM 4-HIL at 93% yield in the two-step system. Moreover, this process facilitated C–H hydroxylation of several hydrophobic aliphatic amino acids to produce hydroxy amino acids, and C–H sulfoxidation of sulfur-containing l-amino acids to yield l-amino acid sulfoxides. Thus, we constructed an efficient cascade reaction to produce 4-HIL by providing prerequisite 2-KG from cheap and plentiful l-Glu and developed a strategy for creating enzymatic systems catalyzing 2-KG-dependent reactions in sustainable bioprocesses that synthesize other functional compounds.

**Keywords:** Fe(II)/2-ketoglutarate-dependent dioxygenase; 2-ketoglutarate generation; regio- and stereo-selective synthesis; hydroxy amino acids; sequential cascade reaction

### **1. Introduction**

The C–H functionalization of small molecules is an important reaction in organic synthesis [1,2]. The oxidized products were used as building blocks in pharmaceutical syntheses [3]. The C−H bond activation by biocatalysis has been reported to date, with less environmental impact compared with conventional counterparts [4]. Fe (II)- and 2-ketoglutarate-dependent dioxygenases (Fe(II)/2-KG DOs) catalyze various C–H-activation-mediated reactions including hydroxylation, deoxygenation, desaturation, and ring extension [5]. Recently, research attention has been directed to the asymmetric hydroxylation of inactivated carbon atoms via direct C–H bond functionalization [1]. Amino acids are substrates of Fe(II)/2-KG DOs and are usually converted into hydroxy amino acids [6]. Various amino acid hydroxylases producing hydroxy amino acids have been characterized among the Fe(II)/2-KG DOs. These include lysine hydroxylase [7], asparagine hydroxylase [8], l-Ile dioxygenase (IDO) [9], leucine hydroxylase [10], and proline hydroxylase [11,12].

Hydroxy amino acids have numerous applications in the pharmaceutical and food industries. They serve as chiral building blocks and food additives [13]. For example, *trans*-4-hydroxy-l-proline (*trans*-4-Hyp) is a major component of collagen [14]. It is a crucial precursor in the pharmaceutical synthesis of n-aryl pyrrole [15] and (−)-kainic acid [16]. Moreover, (2s,3r,4s)-4-hydroxyisoleucine (4-HIL) naturally occurs in *Trigonella foenum-graecum* (fenugreek) seeds, accelerates insulin secretion, and could be administered to treat type II diabetes [17]. In addition, 4-hydroxynorvaline isolated from *Lathyrus odoratus* seeds [18] stimulates insulin secretion and has antidiabetic properties [19]. Further, 5-hydroxyleucine participates in griselimycin synthesis. Griselimycin may have efficacy against drug-resistant *Mycobacterium tuberculosis* and strong antituberculosis activity [10].

Fe(II)/2-KG-DOs catalyze C–H oxidation using 2-KG as a co-substrate. Thus, the amount of 2-KG directly influences the catalytic efficiency of Fe(II)/2-KG DOs. Several studies applied metabolic engineering to generate 2-KG and facilitate Fe(II)/2-KG DO-mediated C–H bond oxidation. Smirnov et al. redirected the tricarboxylic acid (TCA) cycle intermediate 2-KG to biotransform l-Ile into 4-HIL using IDO. They achieved 82% yield using 100 mM l-Ile as the substrate [20]. Zhang et al. optimized the TCA cycle by dynamically modulating the activity of the 2-ketoglutarate dehydrogenase complex in *Corynebacterium glutamicum.* In this manner, they generated ample 2-KG and yielded 232.52 mM of 4-HIL after 64 h [21]. A similar strategy was applied to other Fe(II)/2-KG-DOs-mediated reactions in order to overcome insufficient 2-KG supply. Lin et al. reported a deacetoxycephalosporin-C synthase (DAOCS) that converted penicillin substrates into cephalosporins. They reconstituted the TCA cycle to force 2-KG into an enzymatic reaction [22]. However, this modification had a negative effect on cell growth [21,22], low conversion efficiency, and long reaction time. In contrast, 2-KG has also been produced by chemical synthesis and enzymatic oxidation [23,24].  Liu et al. reported the use of l-amino acid oxidase (LAAO) for 2-KG production with l-glutamic acid (l-Glu) as the substrate. Nevertheless, LAAO had low activity towards l-Glu and was inhibited by high substrate and product concentrations. l-glutamate oxidase (LGOX) has been identified with high l-Glu-to-2-KG conversion activity without requiring exogenous (flavin adenine dinucleotide) FAD [25]. Since the oxidative deamination from l-Glu to 2-KG is along with H2O2 generation, Wu et al. developed a cascade strategy of co-expression LGOX and catalase (CAT) to eliminate H2O2 [26].

In vitro and in vivo (whole-cell) enzymatic syntheses of complex molecules have been successfully implemented in recent years [27]. Here, we implemented an in vitro three-enzyme system to synthesize hydroxy amino acids. We used IDO to hydroxylate l-amino acids and performed oxidative decarboxylation of 2-KG to succinate (SA) [28]. As the consumption of superstoichiometric amount of 2-KG is one of the limitations of the system, an adequate supply of 2-KG derived from inexpensive substrates is necessary for an in vitro system. Hence, 2-KG generation from l-Glu catalyzed by LGOX was adopted in combination with CAT to eliminate the H2O2 by-product. An in vitro multi-enzymatic system was successfully established to synthesize the novel insulin secretion accelerant 4-HIL for the treatment of type II diabetes. Moreover, this system is highly efficient at catalyzing reactions involving other hydrophobic aliphatic l-amino acids and could synthesize hydroxy amino acids [28]. Here, Fe(II)/2-KG DOs-mediated C–H oxidation of small molecules was accomplished in vitro using an efficient and sustainable 2-KG supply system that could generate this substrate for other enzymes catalyzing the synthesis of various functional compounds.

### **2. Results and Discussion**

### *2.1. Design of a Fe(II)*/*2-KG DO-Based In Vitro Enzymatic Biosystem for Hydroxy Amino Acid Synthesis*

Fe(II)/2-KG DOs catalyze various reactions including hydroxylation by C–H activation. Binding of the 2-KG co-substrate facilitates substrate binding and forms a ferryl intermediate that is critical for substrate hydroxylation [29]. The quantity of 2-KG directly affects the catalytic efficiency of Fe(II)/2-KG DOs. Thus, we implemented an in vitro bio-cascade reaction system to synthesize hydroxy amino acids by coupling Fe(II)/2-KG DOs-mediated C–H hydroxylation with 2-KG derived from cheap, readily available l-Glu (Figure 1). In this multi-enzymatic cascade system, IDO, the Fe(II)/2-KG DO, catalyzes l-amino acid hydroxylation and synthesizes useful hydroxy amino acids and certain sulfur-containing l-amino acids. This process is accompanied by oxidative 2-KG decarboxylation using Fe2+ as a cofactor [28]. LGOX exhibits high specificity for the conversion of l-Glu to 2-KG and simultaneously generates H2O2. Catalase (CAT) eliminates the H2O2 [25], thereby preventing its potentially negative effects on the cascade reaction such as the oxidization of 2-KG to succinic acid. Overall, hydroxy amino acid synthesis is accomplished with a simultaneous 2-KG supply by constructing a cell-free reaction system with no additional substrate consumption or membrane-induced mass transfer [30]. Compared with in vivo 2-KG accumulation from the TCA cycle, in vitro cell-free enzymatic system for 2-KG generation has the advantages of high space-time yield and easy operation in product separation. Moreover, this strategy is applicable to other enzymes that catalyze 2-KG-coupled reactions and synthesize other functional compounds.

**Figure 1.** Multi-enzymatic cascade system for hydroxy amino acid synthesis by combining Fe(II)/2-ketoglutarate-dependent dioxygenase (Fe(II)/2-KG DO), l-glutamate oxidase (LGOX), and catalase (CAT).

### *2.2. Enzyme Kinetics*

IDO and LGOX in the cascade reaction were purified by His-Trap HP affinity chromatography (Figure S1), and we optimized the reaction conditions to achieve maximum system efficiency (Figure S2). According to the optimized results, we observed the kinetics of IDO and LGOX under 50 mM Tris-HCl, pH 7.0, and 30 ◦C. In these conditions, the IDO and LGOX activity levels were relatively high. The kinetic parameters were fitted to a Michaelis–Menten model (Table 1). IDO exhibited high oxidative decarboxylation activity on 2-KG (*V*max = 19.79 ± 0.08 U·mg−1). It also had relatively high activity in the presence of l-Ile (*V*max = 8.99 ± 0.09 U·mg−1). In contrast, l-Ile hydroxylase from *Bacillus cereus* 13658 had an activity of only 0.68 ± 0.06 U·mg−1 [31]. LGOX had high specificity and affinity for l-Glu (*K*m = 2.65 ± 0.11 mM and *V*max = 2.93 U·mg−1). IDO and LGOX did not markedly differ in terms of *k*cat. Therefore, the rate of 2-KG regeneration aligned with that of l-Ile hydroxylation and facilitated 4-HIL synthesis.

**Table 1.** Kinetic parameters of IDO and LGOX.

Kinetic parameters were measured in 50 mM Tris-HCl (pH 7.0) at 30 ◦C.

### *2.3. E*ff*ects of Reaction Components on IDO and LGOX Activity*

There are generally some factors to be considered for constructing a cascade reaction involving different reaction components [32]. Various intermediates and components may influence the reaction process [33]. We investigated the inhibitory effects of l-Glu, l-Ile, SA, l-ascorbic acid (Vc), Fe2+ (FeSO4·7H2O), and H2O2 on single enzyme activity by varying their concentrations. Figure 2 shows that H2O2 strongly inhibited IDO and significantly decreased its activity. Exposure of the enzyme to the oxidizing environment created by H2O2 oxidized the Fe2+ in the IDO active site and decreased enzyme activity. Vc is a reductant that promotes C–H oxidation catalyzed by Fe(II)/2-KG DOs [34]. Vc was required by this system for 4-HIL synthesis. Further, 50 mM SA lowered IDO activity to 74%. However, all other components tested only slightly affected IDO activity. High Fe2+ concentrations inactivate LGOX [25]. The relative activity of LGOX was <20% at 20 mM Fe2+. IDO is a member of the Fe (II)/2-KG-dependent dioxygenase family. It requires 2-KG and Fe2+ as cofactors to catalyze hydroxylation. Here, IDO exhibited little activity towards the substrate even in the absence of Fe2+, possibly because of the Fe2+ pool in the *E. coli* cells [35]. Therefore, the influence of Fe2+ addition on the cascade reaction needs to be further investigated.

**Figure 2.** Effects of various reaction components on IDO (blue) and LGOX (orange) activity. IDO and LGOX activity levels were measured using various concentrations of (**A**) l-Glu, (**B**) l-Ile, (**C**) Fe2+, (**D**) Vc, (**E**) H2O2, and (**F**) SA. The specific activities of the enzymes under standard conditions (5.84 ± 0.07 U·mg−1 for IDO and 2.67 ± 0.09 U·mg−1 for LGOX) were designated as 100%, respectively.

### *2.4. Enzymatic Cascade for 4-HIL Synthesis*

Here, we attempted to develop an in vitro system producing hydroxy amino acids and using IDO, LGOX, and CAT as biocatalysts. We tested one-pot 4-HIL production using a combination of the aforementioned enzymes (Figure 3A).

**Figure 3.** Enzymatic cascade production of 4-HIL. (**A**) Scheme of one-pot production of 4-HIL. (**B**) Scheme of multistep reaction to produce 4-HIL.

We assessed the influence of exogenous Fe2+ on 4-HIL production. We mixed 100 mM l-Glu and 100 mM l-Ile in a 25 mL shaker flask and added 1 g·L−1 IDO, 0.5 g·L−1 LGOX, and 2 mg·L−1 CAT to it. Figure 4 shows that 166 mM of 4-HIL was produced in the absence of Fe2+ whereas 28 mM of 4-HIL was produced in the presence of Fe2+. IDO and LGOX activity obviously decreased in the cascade reaction relative to the single-step reaction (control). The final 4-HIL product did not continue to accumulate after the first hour of the reaction. The reaction mixture presented with low 4-HIL yield (16%) in the absence of Fe2+ and low IDO activity (Figure 4A). As shown in Figure 4B, Fe2+ obviously inhibited LGOX activity. Minimal l-Glu was consumed and little 4-HIL accumulated after 0.5 h. However, residual 2-KG was detected in the reaction mixture after sufficient CAT was added to eliminate the H2O2. IDO and LGOX were simultaneously inhibited by H2O2 and Fe2+, respectively. Therefore, the generation of 2-KG from l-Glu by LGOX was incompatible with the hydroxylation of l-Ile by IDO in one pot. For this reason, it was preferable to conduct a stepwise cascade reaction to produce hydroxy amino acids efficiently.

**Figure 4.** One-pot cascade reaction to produce 4-HIL. (**A**) One-pot cascade reaction in the absence of Fe2+; (**B**) one-pot cascade reaction in the presence of Fe2+.

We then attempted two-step 4-HIL production (Figure 3B) and further optimized the biocatalyst and substrate concentrations. l-Ile in the concentration range of 100–500 mM was hydroxylated with 1 g·L−1 purified IDO. Figure 5A shows that IDO had robust l-Ile hydroxylation activity. We obtained 444 mM of 4-HIL with 500 mM l-Ile as substrate. Thus, the product yield was 89%. Moreover, no substrate inhibition was detected at any substrate concentration. We examined the effects of IDO on l-Ile transformation at various substrate concentrations. The corresponding time courses are shown in Figure 5B. The yield and conversion rate increased with IDO concentration. After 9 h, 2 g·L−1 IDO generated 461 mM of 4-HIL from 500 mM l-Ile at a product yield of 92%. After 9 h, 1 g·L−1 IDO generated 459 mM of 4-HIL at a product yield of 92%. However, concerning the reaction productivity, 4.9 g·(L·g·h)−1 of 4-HIL was obtained with 2 g·L−1 IDO, while 7.5 g·(L·g·h)−1 of 4-HIL was obtained with 1 g·L−1 IDO. Therefore, 1 g·L−1 IDO was selected as the catalyst for l-Ile hydroxylation. Hydrogen peroxide is co-produced with 2-KG and may degrade it to SA. The H2O2 is decomposed to H2O and O2 by CAT [36]. We explored the effect of varying CAT concentration on 2-KG production using 200 mM l-Glu as the substrate. Figure 5C shows that the SA accumulation was <15 mM in the presence of 2–3 mg·L−1 CAT. The 2-KG production slightly increased with increasing CAT concentration. At 2 mg·L−1 CAT, 186 mM of 2-KG was generated within 120 min and the yield was 93%. In contrast, only 2.5% SA was formed. No further increase in 2-KG was observed with increasing CAT concentration. We also tested various l-Glu concentrations to establish the optimal reaction conditions (Figure 5D). There was slight substrate inhibition with increasing l-Glu concentration. LGOX generated 491 mM of 2-KG in the presence of 550 mM L-Glu and the yield was 90%.

**Figure 5.** Optimization of biocatalyst and substrate concentrations. (**A**) 4-HIL production at different L-Ile concentrations with 1 g·L−1 IDO; (**B**) 4-HIL production at various IDO concentrations; (**C**) 2-KG and SA yield at different CAT concentrations in presence of 0.5 g·L−1 LGOX; (**D**) 2-KG yield at various L-Glu concentrations in presence of 0.5 g·L−1 LGOX.

### *2.5. Fed-Batch 4-HIL Synthesis Reaction*

L-Ile is the most hydrophobic amino acid. Its solubility is 32.5 g·L−1 (247.75 mM) in aqueous solution at 25 ◦C [37]. To further improve transformation efficiency of cell-free enzymatic system for 4-HIL synthesis, we explored methods of feeding l-Ile in a 500 mL bioreactor (Figure 6). l-Glu at 550 mM was fully converted to 2-KG in ~5 h and only <35 mM SA accumulated. In the first oxidation reaction, 491 mM of 2-KG was obtained at a reaction rate of 14.5 g·L−1·h−1. Optimal concentrations of the l-Ile hydroxylation components were added to the system and 500 mM l-Ile was added either in one dose or in batches at an initial concentration of 200 mM. IDO maintained robust l-Ile hydroxylation activity when the substrate was added in one dose. We obtained 434 mM of 4-HIL after 6 h of l-Ile hydroxylation. The l-Ile hydroxylation reaction rate significantly decreased when the initial substrate concentration was 200 mM as the l-Ile was rapidly consumed. Nevertheless, the reaction rate increased after substrate supplementation. In the second hydroxylation reaction, when l-Ile was incrementally supplemented, IDO yielded 440 mM of 4-HIL after 9 h. In contrast, IDO produced 465 mM of 4-HIL after the l-Ile was added in one dose. The total reaction time for the entire cascade reaction was 14.5 h. The first step produced 491 mM of 2-KG from l-Glu within 5 h and the yield was 89%. The second step generated 464.96 mM of 4-HIL after 9 h when l-Ile was added in one dose. The yield was 93%. Hence, it is feasible to scale up the foregoing 4-HIL production system. In contrast, an in vivo 2-KG supply for 4-HIL synthesis has been successfully implemented by redirecting the TCA cycle [21,38]. However, metabolic engineering strategies of shifting the carbon flux from l-Ile to 2-KG by boosting the TCA cycle and further increasing the pool of oxaloacetate were adopted to obtain 34.21 g/L (232.45 mM) of 4-HIL at 64 h, inevitably hindering normal cell growth [21]. The cell-free enzymatic system generating 2-KG from l-Glu was favorable for 4-HIL production with high substrate concentration and space–time yield. Furthermore, the methodology optimized here could lay the foundation for the development of other industrial bioprocess technologies.

**Figure 6.** Two-step reaction for 4-HIL production.

### *2.6. In Vitro Enzymatic Biosystem for Hydroxy Amino Acids Synthesis*

Hydroxy amino acids are active ingredients or building blocks in pharmaceutical synthesis. Thus, effective, sustainable hydroxy amino acid production is urgently required. IDO has enzymatic activity towards certain hydrophobic aliphatic l-amino acids and sulfur-containing amino acids [28]. It hydroxylates or sulfonates C–H bonds and generates hydroxy amino acids and l-amino acids sulfoxides, respectively. Here, we used 100 mM of l-amino acids substrates in the presence of IDO. All enzymatic reactions achieved high substrate conversion rates. Figure 7 shows that LGOX generated 102.19 mM of 2-KG from l-Glu within 1 h. IDO produced 16.47 g·L−1 of l-methionine sulfoxide (>99.9% conversion) within the first 0.5 h in the presence of l-methionine (l-Met). IDO generated 14.61 g·L−1 of 4-hydroxy-l-leucine (>99.9% conversion) within 2 h in the presence of l-leucine (l-Leu). IDO converted 89.2% of l-norvaline (l-Nva) after 3 h. When l-norleucine (l-Nle) was the substrate, the putative diastereomers 4-hydroxy-l-norleucine and 5-hydroxy-l-norleucine were detected. The reaction products were characterized by mass spectrometry (Figure S3). The aforementioned

reactions demonstrated that the application of a two-step cascade system was highly efficient for the production of hydroxy amino acids and l-amino acid sulfoxides via direct C–H functionalization.

**Figure 7.** In vitro enzymatic synthesis of hydroxy amino acids. (**A**) Hydroxy amino acids were generated from a two-step system using various l-amino acids as substrates. Products were analyzed by LC-MS. (**B**) Biotransformation data for the productions of different hydroxy amino acids.

### **3. Materials and Methods**

### *3.1. Reagents*

The (2s,3r,4s)-4-hydroxyisoleucine standard was purchased from Cambridge Sigma-Aldrich (Munich, Germany). All other analytical-grade chemicals were obtained from Sinopharm Chemical Reagent Co. (Shanghai, China). All enzymes for genetic manipulations were obtained from TaKaRa Biotechnology Co. (Dalian, China). Protein purification columns were acquired from GE Healthcare (Munich, Germany).

### *3.2. Preparation of Recombinant Enzymes*

Commercial catalase (CAT) was obtained from Aladdin Bio-Chem Technology Co. Ltd. (Shanghai, China). The 6×His-tagged IDO-expressing strain (*E. coli* BL21(DE3)-IDO) [28] and 6×His-tagged LGOX-expressing strain (*E. coli* BL21(DE3)-LGOX) [25,26] were constructed as previously described. IDO and LGOX precultures were incubated overnight in 5 mL of Luria–Bertani (LB) medium (Sangon Biotech, Shanghai, China) with 50 μg·mL−1 kanamycin (Sinopharm Chemical Reagent, Shanghai, China) at 37 ◦C on a rotary shaker (200 rpm). IDO and LGOX were induced with 0.5 mM isopropyl-β-d-1-thiogalactopyranoside (IPTG) in 200 mL of LB medium containing 50 μg·mL−1 kanamycin. The cultures were incubated for 3 h at 37 ◦C and then for 16 h at 17 ◦C.

The cells were harvested by centrifugation (10,800× *g*, 4 ◦C, 5 min) and resuspended in 20 mM Tris-HCl buffer (pH 7.5). The bacterial pellet was disrupted and centrifuged (10,800× *g*, 4 ◦C, 40 min). The recombinant IDO and LGOX supernatants were purified by Ni-affinity chromatography as previously reported [39]. The supernatant was purified in a His-Trap HP affinity column (GE Healthcare, Little Chalfont, UK) [40]. The elution buffer in the affinity columns created a

high salt content in the purified fractions. The excess salt was removed with disposable PD-10 desalting columns (GE Healthcare, Little Chalfont, UK) [41]. The proteins were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) [42] and the yields were determined by NanoDrop 8000 Microvolume UV-Vis spectrophotometry (Thermo Fisher Scientific, Waltham, MA, USA).

### *3.3. Enzymatic Activity Assays*

For IDO, a mixture of 10 mM l-Ile, 1.5 mM FeSO4, 10 mM l-ascorbic acid, 10 mM 2-KG, and 50 mM Tris-HCl buffer (pH 7.0) was incubated in a Thermomixer Comfort incubator (Eppendorf; Hamburg, Germany) at 30 ◦C with 800 rpm agitation. The reaction was initiated with 0.1 g·L−1 purified IDO in a final volume of 300 μL [43]. Samples were collected at 0, 5, and 10 min after enzyme addition and analyzed by HPLC [44]. One unit of enzymatic activity was defined as the amount of enzyme required to catalyze the synthesis of 1 μmol of 4-HIL min−1 under standard conditions.

For LGOX, a mixture of 10 mM l-Glu, 1 mg·L−1 CAT, and 50 mM Tris-HCl buffer (pH 7.0) was incubated in a Thermomixer Comfort incubator (Eppendorf; Hamburg, Germany) at 30 ◦C, with 800 rpm agitation. The reaction was initiated with 0.1 g·L−1 purified LGOX in a final volume of 300 μL. Samples were collected at 1, 3, and 5 min after enzyme addition and analyzed by HPLC. One unit of enzymatic activity was defined as the amount of enzyme required to catalyze the synthesis of 1 μmol of 2-KG· min−1 under standard conditions.

### *3.4. Determination of Km and Vmax*

For IDO, the optimal concentration of the cascade reaction was used and the l-Ile concentration range was 0.5–20 mM. The concentration of 2-KG was in the range of 1–40 mM. The kinetic parameter assays were conducted in triplicate. For LGOX, the optimal concentration of the cascade reaction was used and the l-Glu concentration range was 0.5–20 mM. IDO and LGOX activity were measured as previously described. The kinetic parameters were fitted to a Michaelis–Menten model [45].

### *3.5. E*ff*ects of Reaction Components on Enzyme Activities*

The inhibitory effects of l-Ile, l-Glu, Fe2+, Vc, H2O2, and SA were tested by comparing the initial reaction rates of IDO and LGOX in the presence of various concentration of the aforementioned compounds. IDO and LGOX activity were measured as previously described.

### *3.6. One-Pot Synthesis of 4-HIL*

The reaction mixture consisted of 100 mM l-Ile, 50 mM Vc, 100 mM l-Glu, 1 g·L−1 IDO, 0.5 g·L−1 LGOX, 2 mg·L−1 CAT, and 50 mM Tris-HCl buffer (pH 7.0). Then 5 mM FeSO4·7H2O was added to the mixture to compare the effects of Fe2+ on the cascade reaction. The system comprised 5 mL of the reaction mixture heated to 30 ◦C in a 25-mL shaker flask rotating at 200 rpm.

### *3.7. Two-Step Process for Hydroxy Amino Acid Synthesis*

For the 2-KG regeneration step, the reaction mixture consisted of 0.5 g·L−1 LGOX, various concentrations of l-Glu (300–550 mM) and CAT (1-3 mg·L−1), and 50 mM Tris-HCl buffer (pH 8.0) in a final volume of 5 mL. The reaction was conducted at 30 ◦C and 200 rpm. For the l-amino acid hydroxylation step, l-amino acids hydroxylation by IDO was examined after 2-KG production by LGOX. The reaction mixture comprised 5 mM FeSO4, 50 mM Vc, 1 g·L−1 IDO, and 50 mM Tris-HCl buffer (pH 7.0) in a final volume of 5 mL. Using various substrate concentrations (100–500 mM), the 2-KG obtained from the first step was added in equimolar concentrations with the l-amino acids. The mixture was incubated in a shaker flask at 30 ◦C and 200 rpm.

### *3.8. Fed-Batch Reaction for 4-HIL Synthesis*

For 2-KG generation, the reaction mixture was added to a 500-mL reactor consisting of 0.5 g·L−1 LGOX, 550 mM l-Glu, 2 mg·L−1 CAT, and 50 mM Tris-HCl buffer (pH 8.0) in a final volume of 100 mL. The reaction was conducted at 30 ◦C and 400 rpm. After 5 h, the components containing 10 mM FeSO4, 50 mM Vc, and 1 g·L−1 IDO were mixed into the reactor. The pH was fixed at 7.0. The one-dose and incremental L-Ile addition methods were further compared as l-Ile has low solubility in aqueous solvents. l-Ile 500 mM was added to the reactor in one dose to synthesize 4-HIL. The fed-batch reaction was conducted using 200 mM l-Ile as the initial concentration. Then 150 mM l-Ile was supplemented every 2 h until a final concentration of 500 mM l-Ile was reached. The reactions were conducted at 30 ◦C and 400 rpm. The supernatants were sampled every hour and analyzed.

### *3.9. Analysis of Organic Acids*

The organic acids in the samples were analyzed in a Waters 2695 HPLC system (Waters Co., Milford, MA, USA) fitted with an Atlantis T3 column (4.6 mm × 250 mm). Compounds were detected at 210 nm using an injection volume of 10 μL and a column temperature of 40 ◦C. The mobile phase was 20 mM phosphate buffer (pH 2.8) and the flow rate was 0.8 mL·min−1.

### *3.10. Analysis of Amino Acids*

The samples were analyzed in a Waters 2695 HPLC system (Waters Co., Milford, MA, USA) fitted with a Diomansil C18 column (4.6 mm × 250 mm). The chromatographic conditions were as follows: mobile phase A (NaAc-HAc buffer (50 mM, pH 4.2):acetonitrile = 50:50); mobile phase B (acetonitrile); gradient elution program; flow rate, 1 mL min−1; column temperature, 25 ◦C; injection volume, 10 μL. Post-column derivatization was conducted with Fmoc-Cl [44]. The Fmoc-Cl derivatives of the amino acids were detected at 263 nm.

LC-MS analysis was conducted in a Waters ACQUITY UPLC-MS system fitted with a Waters ACQUITY UPLC HSS C18 reverse-phase column (inner diameter, 1.8 μm) (Waters Co., Milford, MA, USA). The inlet, MS transfer line, and ion source temperatures were set to 280, 280, and 230 ◦C, respectively.

### *3.11. Nuclear Magnetic Resonance (NMR) Analysis*

All products were isolated by cation exchange chromatography using a strong cation resin (C100E (H+ form), Purolite, King of Prussia, PA, USA) [46]. l-Ile hydroxylation generated 40 mM of 4-HIL and 42% yield. The 4-HIL spectrum was recorded by 1H-NMR spectroscopy (Figure S4) as previously described [9].

(2*s*,3r,4*s*)-4-hydroxyisoleucine: δ 3.84 (d, J = 4.4 Hz, 1H), 3.79 (dd, J = 13.6, 6.8 Hz, 1H), 1.92–1.82 (m, 1H), 1.19 (d, J = 6.3 Hz, 3H), 0.91 (d, J = 7.1 Hz, 3H).

#### **4. Conclusions**

Here, we developed an in vitro enzymatic system to synthesize hydroxy amino acids via Fe(II)/2-KG DOs-mediated C–H bond oxidation. Amino acid hydroxylation by Fe(II)/2-KG DO was coupled with generation of the prerequisite 2-KG from inexpensive and abundant l-Glu. After systematically assessing the catalytic parameters of the catalysts, a sequential cascade reaction was preferable for the efficient synthesis of hydroxy amino acids. Optimizing the catalyst and substrate concentrations in the two-step process resulted in 465 mM of 4-HIL and 93% yield. Otherwise, IDO was highly active towards other hydrophobic aliphatic l-amino acids and could, therefore, produce other hydroxy amino acids from them. Hence, this system is promising for in vitro biosynthesis of hydroxy amino acids, and more efforts should be necessary to enhance the resistance of IDO and LGOX towards the inhibition of reaction generated components, so as to facilitate the one-pot reaction system. It is also a viable

alternative to other Fe(II)/2-KG DO-mediated C–H bond functionalization strategies for the synthesis of functional compounds.

### **Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/21/15/ 5347/s1.

**Author Contributions:** Conceptualization, X.-R.J. and Y.N.; methodology, X.-R.J.; software, X.-R.J.; validation, X.-R.J. and H.L.; formal analysis, X.-R.J.; investigation, X.-R.J.; resources, Y.N. and Y.X.; data curation, X.-R.J. and H.L.; writing—original draft preparation, X.-R.J., Y.N. and Y.X.; writing—review and editing, X.-R.J., Y.N. and Y.X.; visualization, X.-R.J., Y.N. and Y.X.; supervision, Y.N. and Y.X.; project administration, Y.N. and Y.X.; funding acquisition, Y.N. and Y.X. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was financially supported by the National Natural Science Foundation of China (NSFC) (No. 21676120), the 111 Project (No. 111-2-06), the High-End Foreign Experts Recruitment Program (No. G20190010083), the Program for Advanced Talents within Six Industries of Jiangsu Province (No. 2015-NY-007), the National Program for Support of Top-Notch Young Professionals, the Fundamental Research Funds for the Central Universities (No. JUSRP51504), the Project Funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions, Top-Notch Academic Programs Project of Jiangsu Higher Education Institutions, the Jiangsu Province "Collaborative Innovation Center for Advanced Industrial Fermentation" Industry Development Program, the Postgraduate Research & Practice Innovation Program of Jiangsu Province (No. KYCX19\_1833), the Program for the Key Laboratory of Enzymes of Suqian (No. M201803), and the National First-Class Discipline Program of Light Industry Technology and Engineering (No. LITE2018-09).

**Conflicts of Interest:** There are no conflicts of interest to declare.

### **References**

© 2020 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Genetic and Functional Diversity of Nitrilases in Agaricomycotina**

### **Lenka Rucká 1, Martin Chmátal 2, Natalia Kulik 3, Lucie Petrásková 2, Helena Pelantová 4, Petr Novotný 2, Romana Pˇríhodová 2, Miroslav Pátek 1,\* and Ludmila Martínková 2,\***

Received: 24 October 2019; Accepted: 22 November 2019; Published: 28 November 2019

**Abstract:** Nitrilases participate in the nitrile metabolism in microbes and plants. They are widely used to produce carboxylic acids from nitriles. Nitrilases were described in bacteria, Ascomycota and plants. However, they remain unexplored in Basidiomycota. Yet more than 200 putative nitrilases are found in this division via GenBank. The majority of them occur in the subdivision Agaricomycotina. In this work, we analyzed their sequences and classified them into phylogenetic clades. Members of clade 1 (61 proteins) and 2 (25 proteins) are similar to plant nitrilases and nitrilases from Ascomycota, respectively, with sequence identities of around 50%. The searches also identified five putative cyanide hydratases (CynHs). Representatives of clade 1 and 2 (NitTv1 from *Trametes versicolor* and NitAg from *Armillaria gallica*, respectively) and a putative CynH (NitSh from *Stereum hirsutum*) were overproduced in *Escherichia coli*. The substrates of NitTv1 were fumaronitrile, 3-phenylpropionitrile, β-cyano-l-alanine and 4-cyanopyridine, and those of NitSh were hydrogen cyanide (HCN), 2-cyanopyridine, fumaronitrile and benzonitrile. NitAg only exhibited activities for HCN and fumaronitrile. The substrate specificities of these nitrilases were largely in accordance with substrate docking in their homology models. The phylogenetic distribution of each type of nitrilase was determined for the first time.

**Keywords:** Basidiomycota; Agaricomycotina; nitrilase; cyanide hydratase; nitrile; substrate specificity; overproduction; homology modeling; substrate docking; phylogenetic distribution

### **1. Introduction**

Nitrilases (NLases; EC 3.5.5.-) constitute branch 1 of the nitrilase (NLase) superfamily, which comprises 13 branches of hydrolases acting on non-peptide C-N bonds [1]. NLases have attracted the attention of academia and industry due to their ability to hydrolyze nitriles under mild conditions and with stereo- or regioselectivities [2–4]. They are also promising for bioremediation, especially as far as their cyanide-degrading subtypes cyanide hydratases (CynHs; EC <IP_ADDRESS>) and cyanide dihydratases (CynDs; EC <IP_ADDRESS>) are concerned [5]. Our understanding of the natural roles of NLases and the cyanide-degrading enzymes still falls behind our knowledge of their biotechnological applications. Nevertheless, some probable functions of these enzymes were proposed such as nitrogen recycling,

protection against herbivores, participation in the metabolism of cyano glycosides and glucosinolates, or the detoxification of hydrogen cyanide (HCN) [6].

The first NLases characterized were almost exclusively of bacterial origin [7], with some exceptions of fungal NLases, which, however, were not sequenced [8–10]. The bacterial NLases were usually classified into a number of substrate-specificity subtypes, i.e., aromatic NLases, aliphatic NLases, arylacetoNLases, bromoxynil-specific NLases, CynHs and CynDs [7]. However, a recent study suggested that aliphatic NLases may not actually exist, since the activities for aliphatic nitriles are present in other NLase types such as arylacetoNLases [11]. The plant NLases could be classified into two main subtypes, namely NIT1 through NIT3 with broad substrate specificities on the one hand [12] and NIT4 with a strict specificity for β-cyanoalanine (β-CA) on the other [13].

Recently, more than 10 new NLases in fungi of the division Ascomycota were characterized [14–16]. These enzymes could be roughly classified into aromatic NLases with a preference for (hetero) aromatic nitriles and arylacetoNLases with a preference for phenylacetonitrile (PAN) or mandelonitrile (MN). In contrast, NLases in the second largest division of fungi, Basidiomycota, have been underexplored. A single NLase was characterized in this taxon (in *Auricularia subglabra*, formerly *Auricularia delicata*) [16]. Yet understanding the function of the nitrile-converting enzymes in this taxon seems to be important with regard to the interactions of these fungi with plants. For instance, pathogenic fungi are among the factors causing damage to agricultural crops. Understanding the defense mechanisms of these fungi can lead to the identification of new targets for their suppression. In contrast, biocontrol fungi as components of the plant microbiome are known to exhibit beneficial effects on plants [17].

Basidiomycota comprise saprophytic and parasitic fungi equipped with a wealth of enzymes enabling them to degrade and utilize the plant biomass. Thus the occurrence of peroxidases, laccases, cellulases and hemicellulases in these fungi has been well documented (e.g., [18,19]). Nitrile-degrading enzymes, if active in these fungi, may support their growth on plant biomass by detoxifying natural nitriles, which are widespread in the plant kingdom [20]. For instance, toxic β-CA is formed by β-CA synthase from cysteine and HCN, whereas NIT4 transforms β-CA into a mixture of asparagine (Asn), aspartic acid (Asp) and ammonia. Thus this pathway not only serves for the detoxification of β-CA but also for nitrogen recycling. HCN, which is produced in plants during the synthesis of ethylene (phytohormone) from methionine or by cyano glycoside breakdown, can be also removed directly through its hydration catalyzed by CynHs in fungi [5].

Studies describing the transformation of nitriles or HCN in Basidiomycota are rare. Thus a psychrophilic "basidiomycete" synthesized 2-aminopropionitrile or 4-amino-4-cyanobutyric acid from HCN, ammonia and the corresponding aldehyde. The hydrolysis of these nitriles proceeded in the same organism, producing alanine and glutamic acid, respectively [21,22]. However, the enzymes catalyzing the synthesis or hydrolysis of these nitriles were not identified and the organism was not classified taxonomically. Another study reported on the elimination of CN− in some species of Basidiomycota (*Trametes versicolor*, *Phanerochaete chrysosporium*, *Pleurotus sajor-caju*) but the mechanism of this process was not explained and no degradation products were detected except for a small amount of ammonia [23].

This work focused on the NLases in Agaricomycotina (mushroom-forming fungi). The aim of this work is to analyze their sequences, which are retrieved from the GenBank, and to predict their enzyme activities from sequence similarities to characterized NLases and from homology models. The sequences are classified into clades and representatives of three clades overproduced in *Escherichia coli* for the first time. The phylogenetic distribution of the homologues of the experimentally confirmed NLases is determined.

### **2. Results**

### *2.1. Occurrence of Putative Nitrilases (NLases) in Agaricomycotina*

The GenBank database was searched for putative NLases in Basidiomycota (subdivisions Agaricomycotina, Ustilaginomycotina and Pucciniomycotina). The templates were the bacterial NLase from *Rhodococcus rhodochrous* (GenBank: ABO46008.1) [24], plant NLases NIT1 (GenBank: NP\_851011.1) [12] and NIT4 (GenBank: AAM65906) [13] from *Arabidopsis thaliana*, fungal NLases from *Fusarium verticillioides* (formerly *Gibberella moniliformis*; GenBank: ABF83489.1) [15] and *Auricularia subglabra* (formerly *Auricularia delicata*; EJD42068.1) [16] and a CynH from *Aspergillus niger* (GenBank: ABX75546) [25]. The sequences which exhibited at least 25% identity (>60% cover) to at least one of the templates were retrieved. These sequences were approximately 200 in total, and the majority of them were in Agaricomycotina (around 150 sequences). If sequences exhibiting identities of ≥ 99% to each other occurred in the same species, one of them was discarded, as well as the sequences with a length atypical of NLase (≤300 or ≥400 amino acids).

Deleting the highly similar sequences and the sequences with atypical lengths resulted in obtaining a set of 135 sequences in Agaricomycotina (Supplementary Materials, Table S1). This set was used to construct a phylogenetic tree (Figure 1A) which enabled the identification of a few clades. The largest of them (clade 1; Figure 1B) consisted of 61 sequences similar to plant NLases (NIT1, NIT4) with identities of around 50%; nitrilases outside clade 1 showed lower identities to plant NLases. A smaller clade (clade 2; Figure 1C) was identified, consisting of 25 sequences with maximum ca. 50% identity to characterized NLases in Ascomycota as their closest homologues. The third clade (Figure 1D) only contained five sequences with identities of over 60% to the characterized CynH from *A. niger*. Therefore these proteins could be predicted as CynHs with a high probability and the clade was designated accordingly. Proteins outside the three clades could form an outgroup.

**Figure 1.** *Cont.*

(**B**)

**Figure 1.** *Cont.*

(**D**)

**Figure 1.** Phylogeny of nitrilases. (**A**) Full tree, (**B**) clade 1, (**C**) clade 2, (**D**) cyanide hydratases. The most parsimonious trees are shown. The consistency index is (0.340025), the retention index was (0.706964), and the composite index was 0.242788 (0.240385) for all sites and parsimony-informative sites (in parentheses). The percentage of replicate trees, in which the associated taxa clustered together in the bootstrap test (500 replicates) is shown above the branches [26]. Zero values above branches correspond to bifurcating branches and appeared as a result of tree rooting. The sequence from bacteria *Mycobacteroides abscessus* (SII44893.1) was used as the outgroup. The full tree was obtained using the subtree-pruning-regrafting (SPR) algorithm [27] with search level 1 in which the initial trees were obtained by the random addition of sequences (10 replicates). This analysis involved 135 amino acid sequences. The nitrilases (NLases) selected for overproduction are in frame.

### *2.2. Selection of Nitrilases for Overproduction, Multiple Alignment and Substrate Specificity Predictions*

Three of the hypothetical proteins were selected for heterologous production in *E. coli*. Two of them represented the major clades of hypothetical NLases found in Basidiomycota. In addition, one of the putative CynHs was selected. These proteins originated from *Trametes versicolor* (protein NitTv1; GenBank: XP\_008032838.1; clade 1), *Armillaria gallica* (protein NitAg; PBL01211.1; clade 2) and *Stereum hirsutum* (NitSh; GenBank: XP\_007307917.1; CynH). The identity levels between the three proteins were below 30%.

BLAST searches in the pdb database were carried out in order to identify suitable templates that enabled us to construct three-dimensional models of the selected NLases. The search revealed that all these NLases had the highest identities (30–32%) to the NLase from *Synechocystis* sp. (3wuy) [28]. In addition, NitTv1 had a 26% identity to an uncharacterized NLase-related protein 1j31 from branch 13 of the NLase superfamily (a putative apolipoprotein N-acyltransferase from *Pyrococcus horikoshii*) and to the bacterial thermophilic NLase 3ivz [29]. The 3ivz protein exhibits a high-sequence identity (82%)

to 1j31. Furthermore, the structural identity of the proteins is also high, as 75.15% of the heavy atoms of both structures can be superimposed with total distance below 1Å as calculated by Yasara GDT analysis. Therefore, 3ivz does not add any new information to fungal nitrilase sequence description and it was not used in the final sequence alignment. Lower identities were found between yeast ω-amidase designated Nit2 (4hg3) [30] and NitAg (23%) or between the pyrimidine-degrading enzyme 2vhh (EC: <IP_ADDRESS>, branch 5) and NitSh (22%). Protein 2vhh is missing many parts of the sequence (not shown); therefore, it was discarded from further analysis. A multiple sequence alignment of the fungal NLase sequences with pdb structures was constructed (Figure 2).

**Figure 2.** Multiple sequence alignment used for homology modeling. Parts of C-termini in NitAg and NitSh (compare Figure S1) were cut due to a missing template. Secondary structure elements are assigned based on 3wuy. The catalytic triad (E, K, C) is labeled with black dots, an additional E residue important for the catalytic mechanism [7] is marked with a blue dot. Other aa residues which are close to the catalytic center and could influence substrate recognition are marked with cyan dots. The numbers of aa residues above the alignment correspond to 3wuy, and those below the alignment to NitTv1/NitSh/NitAg NLases. Loops which are different in 3wuy and fungal NLases are labeled (HL1-HL4).

Previously, aligning NLases with proteins from other branches of the NLase superfamily demonstrated typical insertions in NLases [31]. Generally, the fungal NLases studied here have longer insertions (loops) than the NLase-related protein 1j31 but also NLase 3wuy. NitTv1 has longer loops, namely HL1, HL2 and HL4, while NitSh and NitAg have a longer HL3 (Figure 2). Loop HL4 in 3wuy was designated "substrate binding" and it was proposed that the larger size of this loop was the reason for the broader substrate range of the NLase from *Synechocystis* sp. in comparison with the thermophilic NLase from *Pyrococcus abyssi* [28]. This loop is similar in 3wuy and NitAg but longer in NitTv1 and NitSh.

The model of NitTv1 was built using three templates: 3wuy, 1j31 (HL3 loop) and 4hg3 (HL4 loop). The other two models were based on 3wuy alone. A few typical substrates of NLases and CynHs (**1a**–**8a** in Figure 3 and HCN) were docked in the models.

**D**5 &1**E**5 &22+**F**5 &21+

**Figure 3.** Nitriles examined as potential substrates of nitrilases NitTv1, NitAg and NitSh: fumaronitrile (**1a**), 3-phenylpropionitrile (**2a**), β-cyano-L-alanine (**3a**), benzonitrile (**4a**), 2-cyanopyridine (**5a**), 4-cyanopyridine (**6a**), (*R*)-mandelonitrile (**7a**), (*S*)-mandelonitrile (**8a**) and potential products of their biotransformations (carboxylic acids **1b**–**8b**, amides **1c**–**8c**).

The substrate binding in the catalytic center of the enzyme was characterized by the binding (Glide) score. Lower values of this score correspond to better binding. However, this parameter may not be sufficient to characterize the binding of small substrates such as nitriles, in which the binding score is higher due to a lack of hydrogen-bonding (HB) or stacking interactions. Thus the binding scores may be higher with nitriles than with other (larger) substrates but must not be positive. Additional geometrical constraints were also applied. Their selection was based on the analysis of the yeast ω-amidase Nit2 (4hg3)-oxaloacetate (reaction product) complex (4hg5) [30], and on the NLase catalytic mechanism [32] in which the catalytic E forms a HB with the cyano nitrogen. This N-atom must be positioned within HB distance of the catalytic K, and also close to the catalytic E [32,33]. Furthermore, the correct orientation of the substrate was characterized by the distance of the C-atom in the CN group from the S-atom in the catalytic C (optimally < 3.1 Å). The broadest substrate specificity was predicted for NitSh. In its model, the Glide scores suggested its possible activities for fumaronitrile (FN) (**1a**), β-CA (**3a**), benzonitrile (BN) (**4a**), 2-cyanopyridine (2CP) (**5a**), (*R*)-MN (**7a**), (*S*)-MN (**8a**) and HCN. The distance between the C-atom of the substrate and the S-atom in the catalytic C indicated a suitable orientation of these substrates, being below or only slightly above 3.1 Å. The model of NitTv1 exhibited suitable Glide scores and substrate orientation with FN, 3-phenylpropionitrile (PPN) (**2a**) and β-CA. In NitAg, these docking parameters were only suitable with FN and HCN (Table 1).

**Table 1.** Docking of selected substrates in fungal NLase models.

1 In this case, the substrate was docked in the active site close to C, but the nitrogen in the cyano group of the substrate is rotated outwards from the catalytic E and K, different from the proposed orientation of the intermediate in the 4hg5 crystal [30]. n.i. = not identified. In this case, the program Glide was unable to dock the substrate with the defined geometrical constraints (see text).

### *2.3. Modeling Substrate Interaction with the Active Site Residues*

The catalytic residues E, K, C interact by HB and the second E residue participates in Lys stabilization by a salt bridge (Figure 4 and Supplementary materials, Table S2). Another residue found in a close contact with the ligand corresponds to W146 in 3wuy (W144, F148 and Y139 in NitTv1, NitAg and NitSh, respectively; Figure 4, Figure 5 and Figure S2).

**Figure 4.** Snapshots of the complexes of nitrilase (**A**) NitTv1, (**B**) NitAg and (**C**) NitSh with fumaronitrile after 5 ns of molecular dynamics simulations. Hydrogen bonds (HBs) are shown with yellow dashed lines, fumaronitrile is colored magenta, protein residues have element color, and hydrogens are hidden. Amino acids within 3 Å from ligand and those marked with dots in Figure 2 are shown; some residues are hidden for clarity.

**Figure 5.** Orientation of ligands docked in the active site of nitrilases NitTv1 (**A**–**C**), NitAg (**D**,**E**) and NitSh (**F**–**J**). Enzyme residues within 3.9 Å from ligands are shown as raindrops and colored according to their properties (see legend in the right bottom angle). Ligands are represented as stick model; carbons and hydrogens are hidden: (**A**,**D**,**G**) fumaronitrile; (**B**) 3-phenylpropionitrile; (**C**,**H**) β-cyano-l-alanine; (**E**,**F**) HCN; (**I**) benzonitrile; (**J**) 2-cyanopyridine.

The corresponding Y142 in the NLase from *Rhodococcus rhodochrous* was proposed to be important for the substrate specificity of the enzyme: the replacement of an aromatic residue with a non-aromatic one (A, V or L) abolished the enzyme activity for aliphatic nitriles [33]. All fungal NLases examined here have an aromatic residue at this position except for the yeast Nit 2 protein (pdb code 4hg3). This residue is not directly in the active site, but close to the catalytic E (E140, E144 and E135 in NitTv1, NitAg and NitSh, respectively; Figure 4) and it may stabilize its correct orientation through a HB formed by its backbone atoms. The exact mechanism of discrimination of substrate affinity by

aromaticity of this residue is not clear. However, mutation to a smaller A residue may allow more space for catalytic E side chain rotation.

Another residue which could influence the substrate recognition corresponds to P194 in 3wuy (V203, W197 and P188 in NitTv1, NitAg and NitSh, respectively; Figures 4 and 5). P194 is not conserved in NLases. In 3wuy it was identified as one of the hydrophobic residues in the "substrate-binding loop" [28]. W197 together with F200 in NitAg narrows the binding site, suggesting that the enzyme will not accept big substrates such as PPN, in contrast to NitTv1 (Supplementary materials, Figure S2). Most of the sequences in clade 2 (similar to NitAg) have a W residue in the corresponding position. From other clades only KII93875.1 from *Plicaturopsis crispa* (assigned to clade 1) has W in this position, while other clade 1 sequences have mostly a nonpolar (hydrophobic) V, A or P substitution. Clade 2 sequences share conserved WWP (residues 196–198 in NitAg) motif and loop HL3. NitSh has WPV residues in the corresponding position. Hence we could conclude that a mutation at this point could modulate the substrate affinity: smaller residues should allow bigger substrates.

The residue one position downstream of the catalytic C (W179, W173 and W164 in NitTv1, NitAg and NitSh, respectively) is also close to the substrate (Figures 4 and 5). The majority of nitrilases in Ascomycota also have a W residue at this position [34] like 3wuy. The aromatic residue at this position seems to be important for the charge stabilization upon catalysis and participates in T-shaped stacking with aromatic substrates (Figure 5I). However there are some sequences in the dataset (see Supplementary materials, Table S1), which have a CGE instead of the CWE motif, e.g., in KIY71061.1 (*Cylindrobasidium torrendii*), KZP 15294.1 (*Fibularhizoctonia* sp.), OCF30510.1 (*Kwoniella heveanensis*), ORY35932.1 (*Naematelia encephala*), or other residues after catalytic C, e.g., in ELU37431.1 (*Rhizoctonia solani*), XP\_012046763.1, OXB39643.1 and OWZ73108.1 (*Cryptococcus neoformans*), XP\_007401608.1 (*Phanerochaete carnosa*) and THH19653.1 (*Bondarzewia mesenterica*). The sequence from *P. carnosa* belonged to clade 1 and those from *R. solani* and *B. mesenterica* to clade 2 but the other ones were not classified into any of these clades.

### *2.4. Determination of Nitrilase Activities in* E. coli *Cells*

Isopropyl β-D-1-thiogalactopyranoside (IPTG) induced whole cells were examined for their activities towards typical NLase and CynH substrates which were also used for docking (see above). Preliminary screening of the substrates was carried out using high-performance liquid chromatography (HPLC) or spectrophotometric methods suitable for determining the expected products (see Materials and Methods). Thus the reaction mixtures from the transformations of substrates **1a**–**6a** (Figure 3) and HCN were checked for the presence of the corresponding acids or amides by HPLC. The retention times and ultraviolet/visible (UV/vis) spectra of the products were compared with those of authentic compounds **1b**–**6b**, **1c**, **3c**, **5c** and **6c** (commercial standards except for **1b** and **1c**, which were prepared in our previous work) [16]. The reaction mixtures from the transformations of HCN were examined for the production of formamide, which was determined spectrophotometrically and by HPLC. The production of ammonia from β-CA was determined spectrophotometrically.

The reaction products were detected for FN, PPN, 4CP and β-CA in NitTv1, for FN, BN, 2CP and HCN in NitSh, and for FN and HCN in NitAg. No products or only traces of products were determined in other cases. The identity of the products was then confirmed by NMR (acids **1b**, **3b**; amides **1c**, **2c**) and/or LC-MS (acids **1b**, **2b**, **5b**, **6b**; amides **3c**, **5c**, **6c**) (Supplementary materials, Figures S3–S13). Benzoic acid (**4b**) was identified by comparing its retention time and UV spectrum to those of the authentic standard (λmax 228.7 nm).

The major products were largely carboxylic acids but amides were produced in significant amounts in some cases. Thus amide (formamide) was produced as the major product from HCN by NitSh and NitAg. NitTv1 transformed β-CA (**3a**) into a mixture of Asp (**3b**) and Asn (**3c**) at a ratio of 72:28. Significant amounts of amides **5c** and **6c** were also found in the products from 2CP (**5a**) and 4CP (**6a**) obtained with NitSh and NitTv1, respectively. The transformation of FN (**1a**) largely proceeded on only one of the cyano groups. The ratio of acid **1b** and amide **1c** depended on the origin of the

enzyme. NitTv1 produced more cyano amide **1c** than cyano acid **1b** in contrast to the other two enzymes. The products of NitAg were only detected with high cell densities (OD600 of ca. 10). NitTv1 also formed a minor product **1d** (Figure S3) and traces of fumaric acid (not shown) as a result of the transformation of both cyano groups in FN.

The specific activities for the aforementioned substrates were quantified from the rates of product formation determined by HPLC or spectrophotometry (Table 2). FN was the preferred substrate of NitTv1, while β-CA, PPN and 4CP were transformed at much lower rates (< 8%). The most favored substrate of NitSh was HCN, followed by 2CP, FN and BN transformed at rates one to two orders of magnitude lower. NitAg also exhibited its highest activity for HCN but this activity was almost 50 times lower than at NitSh.

**Table 2.** Relative activities of the selected nitrilases.

1 Activity of 2.30 ± 0.06 U/mg dry cell weight was taken as 100%. 2 Activity of 3.22 ± 0.50 U/mg dry cell weight was taken as 100%. 3 Activity of 153 ± 20 U/mg dry cell weight was taken as 100%.

The experimentally determined activities were in accordance with the predictions, however with some exceptions. NitSh exhibited the broadest substrate specificity (for HCN, 2CP, FN and BN) in accordance with the docking experiments. Nevertheless, the activity predicted for β-CA was not confirmed in this enzyme. The preference of NitTv1 for β-CA, FN, PPN and 4CP and NitAg for HCN corroborated the predictions. However, a significant discrepancy between the docking experiments and the activity assays was observed with MN (the former indicated its suitable orientation in the active sites of NitSh and NitTv1 but no activity was determined in the latter). With NitSh, this may be explained by the racemization of MN resulting in the formation of HCN and benzaldehyde, followed by the removal of HCN by this enzyme. Therefore, MN cannot be restored and benzaldehyde remains in the reaction mixture.

### *2.5. Phylogenetic Distribution of NLases in Basidiomycota*

In the subdivision Agaricomycotina, NLase sequences were found in 18 orders and 64 genera (Table 3). The order with the highest number of NLases (38) was Agaricales, followed by Tremellales and Polyporales with 21 and 16 NLase sequences, respectively. The highest numbers of NLase sequences were found in the genera *Kwoniella* (13), *Rhizoctonia* (11), *Armillaria* (8) and *Moniliophthora* (6).

Sequences of clade 1 were found in 13 orders, while those of clade 2 only occurred in four orders. Four orders contained sequences of both clades. Sequences of clade 1 were most frequent in the orders Agaricales and Polyporales with 14 and 16 sequences, respectively. These sequences occurred in the classes Agaricomycetes and Dacrymycetes but not in the class Tremelomycetes (orders Tremellales and Cystofilobasidiales). Clade 2 NLases were only present in the classes Agaricomycetes and they most often occurred in the orders Agaricales (13) followed by Cantharelasses (seven; all of them in genus *Rhizoctonia*) and Russulases (five). CynHs were also only found in *Agaricomycetes*, and they were quite rare, only occurring in three genera, i.e., *Auricularia*, *Exidia* (Auriculariales) and *Stereum* (Russulales). Sequences not classified into the aforementioned clades were mainly found in Tremellales (21) and Agaricales (10).

**Table 3.** Phylogenetic distribution of nitrilases in Basidiomycota.

1 Assigned according to https://www.ncbi.nlm.nih.gov/Taxonomy. 2 three cyanide hydratases and an arylacetonitrilase. 3 cyanide hydratase. Note: Orders belonging to the class Tremellomycetes and Dacrymycetes are highlighted in light grey and dark grey, respectively. Other orders belong to class Agaricomycetes.

### **3. Discussion**

Basidiomycota are comprised of almost 30,000 known species with 70% of them in their largest subdivision, Agaricomycotina, which, in turn, has Agaricomycetes as its largest class (the smaller ones are Dacrymycetes and Tremellomycetes, together comprising only ca. 2% of the species [35]). In accordance with the sizes of these classes, the searches for putative NLases in Basidiomycota revealed that the majority of these enzymes occurred in the class Agaricomycetes. The study focused on the analysis of a set of 135 non-redundant putative NLases from Agaricomycotina, aiming to elucidate the function and taxonomical distribution of this type of protein.

In this NLase set, a few phylogenetic clades could be distinguished, first of all the two largest clades with 61 and 25 sequences. The distribution of the clades seemed to be taxon-specific. According to the current status of the database, clade 1 occurred in Agaricomycetes and Dacrymycetes, while clade 2 was only present in Agaricomycetes. Putative CynHs were also only found in this class. Tremellomycetes contained a few putative NLases, but these proteins did not belong to any of the aforementioned clades.

To shed light on the activities and functions of the putative fungal NLases from Agaricomycotina, two approaches were used–modeling and substrate docking on one hand and heterologous gene expression followed by activity assays on the other. The results of both approaches were in good accordance with only a few exceptions.

These strategies enabled us, with some degree of certainty, to assign the substrate specificities to the largest NLase clades in Agaricomycotina. The closest characterized homologue of NitTv1 (representative of clade 1) was the NLase NIT4 from *Arabidopsis thaliana* (GenBank: AAM65906) with 52% identity (98% cover), but NIT1, NIT2 and NIT3 NLase isoforms from the same organism (GenBank: NP\_001078234.1, CAA68934.3 and NP\_190018.1, respectively) also shared relatively high identities (47–49%, 98–99% cover) to NitTv1. NIT1-NIT3 on one hand, and NIT4 on the other have distinct substrate specificities: NIT4 was found to have β-CA (an intermediate of cyanide assimilation [6]) as its preferential substrate and hydrolyzed PPN (product of glucosinolate metabolism in plants [6]) with activities more than 100 times lower than those for β-CA [13]. In contrast, NIT1-3 operated on different substrates, primarily arylaliphatic nitriles such as PPN, whereas the conversion of β-CA was less than 1% compared to PPN [12]. Due to these differences between NIT1-NIT3 and NIT4, it was not possible to predict the substrate specificity of NitTv1 on the basis of sequence similarities. Whole-cell assays suggested that NitTv1 was active for both β-CA and PPN (in accordance with the docking experiments) unlike both types of plant NLases. In addition, NitTv1 transformed FN, a substrate not examined with NIT4 [13] or NIT1-NIT3 [12], with activities ca. 12.5 times higher compared to β-CA.

The similarity of NLases from clade 1 to plant NLases suggests that Basidiomycota could obtain the corresponding genes from plants by horizontal gene transfer (HGT). NIT4 is ubiquitous in plants, in contrast to NIT1-3, which is typical for Brassicaceae [12]. It may be hypothesized that the clade 1 NLases in Basidiomycota evolved from the widespread NIT4 enzymes and acquired new substrate specificities and functions. The NLases of clade 1 are the most widespread within Basidiomycota. They have been most frequently found in Agaricales and Polyporales. The transfer of the genes between plants and Basidiomycota is plausible due to frequent interactions between these organisms. For instance, many members of the order Agaricales are symbionts, saprophytes or pathogens of plants, and Polyporales are abundant in wood-rotting species (saprophytes but also pathogens). *Rhizoctonia solani* (Cantharellales), a species with the highest number of NitTv1 homologues, is a plant pathogen able to infect a number of various hosts such as rice, corn, sorghum, bean, soybean, cabbage, lettuce and many others [36].

The closest characterized homologues of NitAg, which was selected as a representative of clade 2, are various NLases from Ascomycota [15,16] with maximum ca. 50% identity to NitAg. Of the tested substrates, only HCN and FN were transformed by this enzyme. The major reaction product of HCN was formamide.

The NitAg homologues are much less abundant in Basidiomycota than NitTv1 homologues and mainly occur in a few genera of the orders Agaricales and Russulales, and in *Rhizoctonia solani* (Cantharellales). In particular *R. solani* is rich in NitAg homologues, which may be due to the exposure of the pathogen to cyanides produced by the plant host. The hypothesis of HGT may be also applied to the NLases in clade 2. It is possible that Basidiomycota acquired these genes from Ascomycota, which co-infect the same host. In fact the infection of a plant host by multiple pathogenic species is common [37].

Similarly, HGT is possible with CynHs, which rarely occur in Basidiomycota but are frequent in Ascomycota, in which a number of CynHs originating from, e.g., *Gloeocercospora sorghi*, *Neurospora crassa*, *Gibberella zeae* or *Aspergillus nidulans* were expressed in heterologous hosts such as *Aspergillus nidulans* or *E. coli* [38,39]. Furthermore, sequenced genomes of Ascomycota contain numerous putative CynHs. In contrast, only five CynHs were identified in Basidiomycota so far (in Auriculariales and Russulales, i.e., saprophytes but also plant pathogens such as *S. hirsutum*). In this case, the prediction of these enzymes as CynHs was possible due to highly conserved primary sequences (with identities of over 60% to characterized CynHs). This assumption was confirmed by activity assays with NitSh from *S. hirsutum*. Its substrate specificity for FN and 2CP, in addition to HCN, was similar to those of some previously characterized CynHs from e.g., *A. niger*, *Botryotinia fuckeliana* or *Pyrenophora teres* [16,25]. NitSh was most similar to the CynH from *N. crassa* (84.5% identity, 94% cover), which was partly characterized previously and was found to be the most stable CynH of those tested [39]. Therefore, the utility of NitSh in the bioremediation of cyanide-containing wastes will be examined.

FN was a substrate of all three of the examined enzymes. This is in accordance with our previous finding that FN was hydrolyzed by all tested NLases of different types and origin [16]. However, the products were dependent on the enzyme. NitTv1 mainly produced the corresponding cyano amide like the previously examined NLase from *Aspergillus kawachii*, which was classified as an "aromatic NLase" [16]. The major product of FN transformation by NitSh was cyano acid instead. This reaction pattern was similar to that of the NLase NitMp from *Macrophomina phaseolina* [16].

In summary, the genomes of Basidiomycota contain genes encoding functional NLases, with the majority of them occurring in the subdivision Agaricomycotina, class Agaricomycetes. The NLases in Agaricomycotina were classified into clades with different substrate specificities and different taxonomical distributions. Three of the enzymes were for the first time characterized in terms of their substrate specificities. Two of them (from *T. versicolor* and *S. hirsutum*) were similar to NLases from other organisms, namely NLases from plants and CynHs from Ascomycota. The third enzyme (from *A. gallica*) had no close characterized homologues among NLases from other organisms. NLases in Basidiomycota need to be studied further with focus on the regulation of their production in fungi. This should help to better understand and exploit the bioremediation potential of these fungi.

### **4. Materials and Methods**

### *4.1. Sequence Analysis*

The GenBank database was searched for NLase sequences using the program BLAST (Available online: http://blast.ncbi.nlm.nih.gov/Blast.cgi) with published NLase sequences (see section Results) as templates. For phylogenetic analysis, multiple sequence alignment was built with the MUSCLE algorithm [40]. Non-informative sites were deleted, i.e., the N- and C-terminal parts were truncated and the regions with less than 90% site coverage were eliminated during alignment preparation in MEGA X [41]. There were a total of 282 positions in the final dataset. Evolutionary analyses were conducted in MEGA X and used for phylogenetic tree building with the maximum parsimony method (500 bootstraps). The cut-off parameter used to isolate the clades from the global tree was a bootstrap value of ≥ 90%. The taxonomical classification of the strains in which NLases were sequenced was according to available online: https://www.ncbi.nlm.nih.gov/Taxonomy.

### *4.2. Nitrilase Modeling and Substrate Docking*

BLAST was used to search for homologous structures in the Protein Data Bank (pdb) database [42]. Multiple sequence alignments were carried out using the program T-Coffee [43] and manually corrected in Jalview [44] based on structure alignment with Yasara [45]. Sequences with the highest identities to the examined NLases were selected as templates for homology modeling with MODELLER 9.16 [46]. The observed models were validated with the structure validation tool MOLPROBITY [47] and minimized using a water model of TIP3P type in Yasara. Structures of substrates were downloaded from the Pubchem database (Available online: https://pubchem.ncbi.nlm.nih.gov/) and minimized with the LigPrep tool in the software Schrödinder [48]. The docking of potential substrates was done using the program Glide (Schrödinger, LLC).

### *4.3. Nitrilase Overproduction*

The synthesis of the NLase genes, which were optimized for *E. coli*, was performed with GeneArt (ThermoFisher Scientific, Regensburg, Germany). The optimized gene sequences are shown in Supplementary Material—Gene and protein sequences (Figure S1). Gene cloning and strain cultivation were as described previously [16]. Briefly, the expression vector was constructed by cloning the gene into the *NdeI* and *XhoI* sites of pET22b(+) and used to transform *E. coli* Origami B (DE3) co-expressing the GroEL/ES chaperone. The cultures were grown in 2 × YT medium and the induction of NLase and chaperone gene expression was performed with 0.02 mM IPTG and 0.011 mM L-arabinose, respectively, at 20 ◦C for 24 h. The cells were centrifuged and used for enzyme assays.

### *4.4. Nitrilase Activity Assays*

The assays were carried out as described previously [16] with minor modifications. Eppendorf tubes (1.5-mL) contained 0.5 mL of the reaction mixture consisting of 50 mM Tris/HCl buffer, pH 8.0, 150 mM NaCl, 25 mM substrate, 5% (*v*/*v*) methanol (cosolvent) and an appropriate amount of whole cells. The substrate was BN, PPN, FN or MN. The reactions with 2CP, 4CP, β-CA or free cyanide (KCN) as substrates were carried out analogously but without methanol. The reactions proceeded at 30 ◦C with shaking, and were terminated after 5–10 min by the addition of 0.050 mL of 2 M HCl for nitriles except β-CA. The reaction of β-CA was terminated by centrifugation of the cells. The reaction of HCN was terminated by adding 1.0 mL of methanol and centrifugation. The activities for β-CA were calculated from the production of ammonia, which was determined spectrophotometrically [11,49]. The activities for HCN were determined from the production of formamide, which was determined spectrophotometrically [50] and by HPLC (see below). The activities for other substrates were determined from the production of carboxylic acids and/or amides, which were determined by HPLC (see below).

### *4.5. Analytical High-Performance Liquid Chromatography (HPLC)*