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Molecular Characterization of a Novel Family VIII Esterase with β-Lactamase Activity (PsEstA) from Paenibacillus sp.

Molecular information about family VIII esterases, which have similarities with class C β-lactamases and penicillin-binding proteins, remains largely unknown. In this study, a novel family VIII esterase with β-lactamase activity (PsEstA) from Paenibacillus sp. was characterized using several biochemical and biophysical methods. PsEstA was effective on a broad range of substrates including tertiary butyl acetate, glyceryl tributyrate, glucose pentaacetate, olive oil, and p-nitrophenyl esters. Additionally, PsEstA hydrolyzed nitrocefin, cefotaxime, and 7-aminocephalosporanic acid. Interestingly, two forms of immobilized PsEstA (CLEAs-PsEstA and mCLEAs-PsEstA) showed high recycling property and enhanced stability, but hybrid nanoflowers (hNFs) of PsEstA require improvement. This study provides a molecular understanding of substrate specificities, catalytic regulation, and immobilization of PsEstA, which can be efficiently used in biotechnological applications.

Introduction
Bacterial β-lactamases hydrolyze chemical compounds containing a β-lactam ring and are the primary cause of bacterial resistance against different classes of β-lactam antibiotics. Therefore, the production of one or more types of β-lactamase is the most effective strategy through which clinically important gram-negative bacteria effectively hydrolyze the β-lactam rings of antibiotics, such as penicillins, cephalosporins, or carbapenems [1][2][3]. Based on amino acid motifs, protein sequence, molecular size, and 3D structure, β-lactamases are classified into four molecular classes termed A, B, C, and D [4]. Interestingly, classes A, C, and D hydrolyze their substrates through the formation of an acyl-enzyme with a catalytic serine, whereas class B utilizes a zinc ion to facilitate a hydrolytic reaction. Among β-lactamases, class C enzymes are of high importance, because they pose significant threats for antibiotic treatments due to their occurrence in many gram-negative pathogens, such as Enterobacteriaceae and Pseudomonas spp. [5,6]. Class C β-lactamases play a role in the resistance of these pathogens to cephalosporins, cephamycins, and carbapenems, and are not inhibited by clavulanic acid [7,8]. The molecular structure of class C β-lactamases consists of two domains of different sizes. The bigger domain contains a central antiparallel beta sheet flanked by alpha helices at each side, was performed in the positive ion mode on a Voyager™ BioSpectrometry™. Urea-induced unfolding was performed following 1 h incubation of PsEstA with urea at 25 • C. The emission spectra from 300 nm to 400 nm were recorded after excitation at 295 nm. Thermal unfolding was monitored from 15 • C to 90 • C at 222 nm using circular dichroism (CD) signals with a thermostatic cell holder.

β-Lactamase Assay
The β-lactamase activity of PsEstA was determined using the chromogenic β-lactam substrate nitrocefin [17]. PsEstA (0.1 mg/mL) was reacted with a 484 µM nitrocefin solution in 20 mM Tris-HCl pH 8.5 at 25 • C, and the color change in the reaction mixture was observed. In addition, pH indicator-based colorimetric assay was performed to investigate the activity of PsEstA on 1 mM cefotaxime (CTX) and 7-aminocephalosporanic acid (7-ACA) at 25 • C for 1 h.

Immobilization of PsEstA
For preparing crosslinked enzyme aggregates (CLEAs), 0.5 mg·mL −1 of PsEstA was coprecipitated by 80% (w/v) ammonium sulfate at 4 • C. Crosslinking was then performed by the dropwise addition of glutaraldehyde to the final concentration of 25 mM at room temperature. After an overnight incubation followed by centrifugation at 15,000 g for 10 min, the pellet (CLEA-PsEstA) was resuspended and repeatedly washed until no activity was observed in the supernatant. The preparation and addition of the magnetic Fe 3 O 4 nanoparticles were carried out as previously described [27,31]. Nanoparticles were mixed with PsEstA solution containing ammonium sulfate and glutaraldehyde. After gentle agitation for 12 h, magnetic CLEAs of PsEstA (mCLEA-PsEstA) were precipitated by centrifugation at 15,000× g for 10 min and stored in 50 mM sodium phosphate buffer (pH 7.5) at 4 • C. Scanning electron microscope images were obtained at various magnifications (50,000-100,000×) using a Carl Zeiss SUPRA 55VP microscope. The chemical stabilities of the three forms of PsEstA (free PsEstA, CLEA-PsEstA, and mCLEA-PsEstA) were determined by measuring the residual activity toward p-nitrophenyl butyrate (p-NB) after 1 h incubation in the presence of the chemical compounds [ethanol, iso-propanol, sodium dodecyl sulfate (SDS), urea] at 25 • C. For the recycling process, the two immobilized forms of PsEstA (CLEA-PsEstA and mCLEA-PsEstA) were extensively washed and then reused in the next cycle. For preparing enzyme-inorganic hybrid nanoflower (hNF), PsEstA (0.05, 0.1, and 0.5 mg·mL −1 ) was added into 800 µM of the Cu 2+ metal solutions for a total volume of 3 mL [32]. The resulting mixtures were incubated at 25 • C and centrifuged at 15,000× g for 10 min. The pellets were resuspended and transferred to 1.5 mL microtubes. In the reusability tests, hNF-PsEstA was recovered by centrifugation and extensively washed before being reused in the next cycle. Then, fresh substrate of 1 mM p-nitrophenyl butyrate (p-NB) was reacted for another cycle and activity was measured. In these experiments, the enzyme activity of free PsEstA was defined as 100%.

Bioinformatic Analysis
For phylogenetic tree analysis of PsEstA, the 26 representative bacterial lipases/esterases sequences were analyzed. As shown in Figure 1, PsEstA belongs to family VIII esterases, which include EstC [12], Est-Y29 [33], EstB [34], and EstU1 [35]. solutions for a total volume of 3 mL [32]. The resulting mixtures were incubated at 25 °C and centrifuged at 15,000× g for 10 min. The pellets were resuspended and transferred to 1.5 mL microtubes. In the reusability tests, hNF-PsEstA was recovered by centrifugation and extensively washed before being reused in the next cycle. Then, fresh substrate of 1 mM p-nitrophenyl butyrate (p-NB) was reacted for another cycle and activity was measured. In these experiments, the enzyme activity of free PsEstA was defined as 100%.

Bioinformatic Analysis
For phylogenetic tree analysis of PsEstA, the 26 representative bacterial lipases/esterases sequences were analyzed. As shown in Figure 1, PsEstA belongs to family VIII esterases, which include EstC [12], Est-Y29 [33], EstB [34], and EstU1 [35]. As shown in Figure 2A, three motifs were found to be highly conserved among class C βlactamases based on multiple sequence alignment. These three motifs (motif I, II, and III) have also been identified in family VIII esterases. In motif I, a nucleophilic Ser 58 is located in the characteristic S 58 -x-x-K 61 tetrapeptide, and the alkoxide ion formation at Ser 58 is believed to be enhanced by Lys 61 . The Tyr 172 residue in motif II is involved in the substrate recognition and specificity regulation. Although motif III of class C β-lactamases is composed of the highly conserved K-T-G sequence, a W-x-G motif is observed instead in PsEstA and other family VIII esterases. The third residue of motif III has to be glycine because otherwise, other amino acids would be sterically unfavorable for the binding of the substrate. In PsEstA, the W-S-G sequence was observed in motif III. In sequence analysis, PsEstA was found to contain a higher percentage of acidic (Asp and Glu, 12.0%) than basic (Lys and Arg, 9.3%) amino acids. As shown in Figure 2A, three motifs were found to be highly conserved among class C β-lactamases based on multiple sequence alignment. These three motifs (motif I, II, and III) have also been identified in family VIII esterases. In motif I, a nucleophilic Ser 58 is located in the characteristic S 58 -x-x-K 61 tetrapeptide, and the alkoxide ion formation at Ser 58 is believed to be enhanced by Lys 61 . The Tyr 172 residue in motif II is involved in the substrate recognition and specificity regulation. Although motif III of class C β-lactamases is composed of the highly conserved K-T-G sequence, a W-x-G motif is observed instead in PsEstA and other family VIII esterases. The third residue of motif III has to be glycine because otherwise, other amino acids would be sterically unfavorable for the binding of the substrate. In PsEstA, the W-S-G sequence was observed in motif III. In sequence analysis, PsEstA was found to contain a higher percentage of acidic (Asp and Glu, 12.0%) than basic (Lys and Arg, 9.3%) amino acids. The structural model of PsEstA consisted of a large α/β domain and a small α-helical domain, and these features are frequently observed in other β-lactamases and family VIII esterases [14,17,34,35]. The nucleophilic serine was found to be located near the surface of the tunnel formed The structural model of PsEstA consisted of a large α/β domain and a small α-helical domain, and these features are frequently observed in other β-lactamases and family VIII esterases [14,17,34,35]. The nucleophilic serine was found to be located near the surface of the tunnel formed by two domains ( Figure 2B). The putative catalytic triad made of Ser 58 , Lys 61 , and Tyr 169 is positioned within a catalytic pocket close to the surface ( Figure 2B, rectangular region). The substrate-binding pocket is mainly delineated by the aromatic amino acids Tyr 127 , Tyr 169 , and Trp 333 , presumably regulating the entrance of substrates via hydrophobic interactions. These residues are also highly conserved in most family VIII esterases. In molecular docking analysis, Tyr 169 and Trp 333 were shown to stabilize the p-nitrophenol ring, while side chains of Asp 302 and Ser 334 formed hydrogen bonds with the nitro moiety (-NO 2 ) of p-nitrophenyl butyrate ( Figure 2C). In addition, backbone nitrogen of Ala 336 is involved in the formation of an oxyanion hole. This model analysis suggested a number of active-site residues seem to be involved in hydrogen bonding networks among other conserved residues or substrate [34]. In addition to conservation of primary sequences, spatial orientation of catalytic residues between class C β-lactamases and family VIII esterases are very similar. As shown in Figure 3, catalytic residues of PsEstA have comparable conformation to those of AmpC [36] and blaMOX-1 [37], as well as EstU1 and Est-Y29.
Biomolecules 2019, 9, x FOR PEER REVIEW 6 of 15 by two domains ( Figure 2B). The putative catalytic triad made of Ser 58 , Lys 61 , and Tyr 169 is positioned within a catalytic pocket close to the surface ( Figure 2B, rectangular region). The substrate-binding pocket is mainly delineated by the aromatic amino acids Tyr 127 , Tyr 169 , and Trp 333 , presumably regulating the entrance of substrates via hydrophobic interactions. These residues are also highly conserved in most family VIII esterases. In molecular docking analysis, Tyr 169 and Trp 333 were shown to stabilize the p-nitrophenol ring, while side chains of Asp 302 and Ser 334 formed hydrogen bonds with the nitro moiety (-NO2) of p-nitrophenyl butyrate ( Figure 2C). In addition, backbone nitrogen of Ala 336 is involved in the formation of an oxyanion hole. This model analysis suggested a number of activesite residues seem to be involved in hydrogen bonding networks among other conserved residues or substrate [34]. In addition to conservation of primary sequences, spatial orientation of catalytic residues between class C β-lactamases and family VIII esterases are very similar. As shown in Figure  3, catalytic residues of PsEstA have comparable conformation to those of AmpC [36] and blaMOX-1 [37], as well as EstU1 and Est-Y29.

Characterization of PsEstA
The recombinant PsEstA was highly purified using an immobilized metal-affinity column, and a single band was observed ( Figure 4A). The molecular mass of PsEstA was estimated to be approximately 45 kDa by SDS-PAGE, and this value is similar to those of Est-Y29 [33] and EstB [34], but smaller than EstU1 [35]. An overlay activity assay showed high fluorescence at the position where purified PsEstA was located in native PAGE ( Figure 4B). Furthermore, strong fluorescence intensity was observed for 4-MU acetate and PsEstA, but not for 4-MU phosphate and PsEstA ( Figure 4C,D). In native PAGE, the molecular mass of active PsEstA was found to be substantially greater than 200 kDa, implying that PsEstA is in an oligomeric conformation. A similar behavior indicating an oligomeric state of PsEstA was also observed in gel filtration chromatography ( Figure 4E), which has been reported for other family VIII esterases [38][39][40]. The mass spectrometric analysis showed a major peak (m/z) at 47.7 kDa with all additional amino acids including the His-tag ( Figure 4F). was observed for 4-MU acetate and PsEstA, but not for 4-MU phosphate and PsEstA ( Figure 4C,D). In native PAGE, the molecular mass of active PsEstA was found to be substantially greater than 200 kDa, implying that PsEstA is in an oligomeric conformation. A similar behavior indicating an oligomeric state of PsEstA was also observed in gel filtration chromatography ( Figure 4E), which has been reported for other family VIII esterases [38][39][40]. The mass spectrometric analysis showed a major peak (m/z) at 47.7 kDa with all additional amino acids including the His-tag ( Figure 4F).

Biochemical Assay
The hydrolytic activity of PsEstA was analyzed using p-nitrophenyl (p-NP) esters with acyl chain lengths of different length [27][28][29]41]. As shown in Figure 5A, PsEstA had a strong substrate preference for p-NA or p-NB. However, almost no enzyme activity was observed for the long-chain substrates p-NO and p-ND. Similarly, other members of family VIII esterases such as Est22 [13], EstU1 [42], or EstM-N1 [43] showed substrate preference for p-NB. When naphthyl esters were used as the substrates, the highest enzymatic activities were observed with 1-naphthyl acetate (1-NA), followed by 1-naphthyl butyrate (1-NB) and 2-naphthyl acetate (2-NA). However, mutation of Ser 58 residue abolished most of the hydrolytic activity ( Figure 5B). As shown in Figure 5B, PsEstA exhibited

Biochemical Assay
The hydrolytic activity of PsEstA was analyzed using p-nitrophenyl (p-NP) esters with acyl chain lengths of different length [27][28][29]41]. As shown in Figure 5A, PsEstA had a strong substrate preference for p-NA or p-NB. However, almost no enzyme activity was observed for the long-chain substrates p-NO and p-ND. Similarly, other members of family VIII esterases such as Est22 [13], EstU1 [42], or EstM-N1 [43] showed substrate preference for p-NB. When naphthyl esters were used as the substrates, the highest enzymatic activities were observed with 1-naphthyl acetate (1-NA), followed by 1-naphthyl butyrate (1-NB) and 2-naphthyl acetate (2-NA). However, mutation of Ser 58 residue abolished most of the hydrolytic activity ( Figure 5B). As shown in Figure 5B, PsEstA exhibited approximately 75% activity on 2-NA. In contrast, it showed almost no activity on 1-naphthyl phosphate (1-NP). PsEstA was also shown to be highly stable in the presence of NaCl and glycerol. Specifically, it was shown to retain approximately 100% and 70% of its enzymatic activity in the presence of 3.0 M and 4.0 M NaCl, respectively ( Figure 5C). In addition, PsEstA exhibited strong tolerance to glycerol with almost no loss of enzymatic activity after incubating with up to 30% glycerol.
The chemical stability of PsEstA was investigated by monitoring the intrinsic fluorescence spectra. In its native form, PsEstA exhibited a λ max at 334 nm, implying that all the tryptophan residues of PsEstA were located in the hydrophobic interior. However, a red shift of λ max to 344 nm was observed with a remarkable increase in fluorescence intensity in the presence of 5 M urea, suggesting that the tryptophan residues were mostly exposed to the solvent ( Figure 5D). The chemical stability of PsEstA in urea was also investigated by measuring the enzymatic activity in the presence of increasing concentrations of urea. At 4.0 M,~50% of the initial activity was retained ( Figure 5E). Next, the thermal stability of PsEstA was investigated by monitoring its thermal denaturation from 15 to 90 • C using far-UV CD at 222 nm. PsEstA showed only minor changes up to 50 • C, and its melting temperature was determined as 63 • C ( Figure 5F). approximately 75% activity on 2-NA. In contrast, it showed almost no activity on 1-naphthyl phosphate (1-NP). PsEstA was also shown to be highly stable in the presence of NaCl and glycerol. Specifically, it was shown to retain approximately 100% and 70% of its enzymatic activity in the presence of 3.0 M and 4.0 M NaCl, respectively ( Figure 5C). In addition, PsEstA exhibited strong tolerance to glycerol with almost no loss of enzymatic activity after incubating with up to 30% glycerol. The chemical stability of PsEstA was investigated by monitoring the intrinsic fluorescence spectra. In its native form, PsEstA exhibited a λmax at 334 nm, implying that all the tryptophan residues of PsEstA were located in the hydrophobic interior. However, a red shift of λmax to 344 nm was observed with a remarkable increase in fluorescence intensity in the presence of 5 M urea, suggesting that the tryptophan residues were mostly exposed to the solvent ( Figure 5D). The chemical stability of PsEstA in urea was also investigated by measuring the enzymatic activity in the presence of increasing concentrations of urea. At 4.0 M, ~50% of the initial activity was retained ( Figure 5E). Next, the thermal stability of PsEstA was investigated by monitoring its thermal denaturation from 15 to 90 °C using far-UV CD at 222 nm. PsEstA showed only minor changes up to 50 °C, and its melting temperature was determined as 63 °C ( Figure 5F).

Substrate Analysis
The hydrolytic properties of PsEstA for acetylated carbohydrates, tertiary alcohol esters, and lipids were evaluated using a colorimetric assay [28,29,41]. The tertiary alcohol esters tert-butyl acetate, α-terpinyl acetate, and linalyl acetate were used. As shown in Figure 6A, PsEstA could effectively hydrolyze tert-butyl acetate, but not linalyl acetate or α-terpinyl acetate. Additionally, a significant hydrolytic activity of PsEstA was detected only for glyceryl tributyrate based on the yellow color of the solution ( Figure 6B). Furthermore, PsEstA displayed a high activity for glucose pentaacetate, although no significant activity was detected for cellulose acetate or glucosamine acetate ( Figure 6C). In addition, a high level of enzymatic activity of PsEstA was observed toward olive oil, judging from the fluorescence spectra ( Figure 6D,E). For enantioselectivity analysis, (R)-and (S)-methyl-β-hydroxyisobutyrate were used [27,32]. Following incubation with PsEstA, only the reaction mixture containing the (S)-enantiomer turned yellow, showing the (S)-selectivity of PsEstA ( Figure 6F). This indiscriminate nature of PsEstA could be useful for industrial applications as a biocatalyst. acetate ( Figure 6C). In addition, a high level of enzymatic activity of PsEstA was observed toward olive oil, judging from the fluorescence spectra ( Figure 6D,E). For enantioselectivity analysis, (R)-and (S)-methyl-β-hydroxyisobutyrate were used [27,32]. Following incubation with PsEstA, only the reaction mixture containing the (S)-enantiomer turned yellow, showing the (S)-selectivity of PsEstA ( Figure 6F). This indiscriminate nature of PsEstA could be useful for industrial applications as a biocatalyst.

β-Lactamase Activity of PsEstA
To investigate the β-lactamase activity of PsEstA, a chromogenic β-lactamase substrate (nitrocefin) was used as a substrate [12,17]. As shown in Figure 7, PsEstA apparently showed β-lactamase activity toward nitrocefin. The β-lactamase activity of PsEstA was further investigated using 7-ACA and CTX. PsEstA showed significant catalytic activities toward cefotaxime (CTX) and 7-aminocephalosporanic acid (7-ACA) in a pH shift assay ( Figure 7E,F). In accordance with PsEstA, Est22 and EstU1 were shown to possess a noteworthy β-lactam hydrolytic activity [13,35]. In contrast, EstC showed a significant hydrolyzing activity for nitrocefin, but none of these esterases displayed any activity for other β-lactam substrates [12]. EstC showed a significant hydrolyzing activity for nitrocefin, but none of these esterases displayed any activity for other β-lactam substrates [12].

Immobilization of PsEstA
Enzyme immobilization has been explored in a large variety of industrial applications [44,45]. Crosslinked enzyme aggregates (CLEAs) constitute one of the highly studied methods to generate highly efficient immobilized biocatalysts [46]. Toward this end, CLEAs-PsEstA was prepared by precipitating PsEstA with ammonium sulfate and glutaraldehyde. The SEM images of CLEAs-PsEstA showed formation of globular structures with a diameter of 1-2 nm ( Figure 8A). The stabilities of both free PsEstA and CLEAs-PsEstA in the presence of several chemical compounds were compared ( Figure 8B). For 5 M urea and 0.1% (v/v) SDS, there were no significant differences between these two forms. However, for alcohols such as ethanol and iso-propanol, CLEAs-PsEstA was highly active compared with the free form of the enzyme. Specifically, with 30% ethanol, free PsEstA retained only ~3% of its original activity, whereas CLEAs-PsEstA showed ~35% of the original activity. The operational stability of CLEAs-PsEstA was studied up to 20 cycles. As shown in Figure 8C, CLEAs-PsEstA was highly stable for 20 cycles, retaining at the end ~75% of the original activity.
Furthermore, PsEstA was immobilized as CLEAs on magnetite nanoparticles for an efficient separation during industrial applications [27,32]. To obtain magnetic CLEA form of PsEstA (mCLEA-PsEstA), PsEstA was coaggregated with nanoparticles, and then chemically crosslinked using glutaraldehyde. Transmission electron microscopy showed that the magnetic nanoparticles had a diameter of 5-8 nm ( Figure 8D). The effects of the chemical compounds on the activities of mCLEA-

Immobilization of PsEstA
Enzyme immobilization has been explored in a large variety of industrial applications [44,45]. Crosslinked enzyme aggregates (CLEAs) constitute one of the highly studied methods to generate highly efficient immobilized biocatalysts [46]. Toward this end, CLEAs-PsEstA was prepared by precipitating PsEstA with ammonium sulfate and glutaraldehyde. The SEM images of CLEAs-PsEstA showed formation of globular structures with a diameter of 1-2 nm ( Figure 8A). The stabilities of both free PsEstA and CLEAs-PsEstA in the presence of several chemical compounds were compared ( Figure 8B). For 5 M urea and 0.1% (v/v) SDS, there were no significant differences between these two forms. However, for alcohols such as ethanol and iso-propanol, CLEAs-PsEstA was highly active compared with the free form of the enzyme. Specifically, with 30% ethanol, free PsEstA retained only~3% of its original activity, whereas CLEAs-PsEstA showed~35% of the original activity. The operational stability of CLEAs-PsEstA was studied up to 20 cycles. As shown in Figure 8C, CLEAs-PsEstA was highly stable for 20 cycles, retaining at the end~75% of the original activity.
Furthermore, PsEstA was immobilized as CLEAs on magnetite nanoparticles for an efficient separation during industrial applications [27,32]. To obtain magnetic CLEA form of PsEstA (mCLEA-PsEstA), PsEstA was coaggregated with nanoparticles, and then chemically crosslinked using glutaraldehyde. Transmission electron microscopy showed that the magnetic nanoparticles had a diameter of 5-8 nm ( Figure 8D). The effects of the chemical compounds on the activities of mCLEA-PsEstA and free PsEstA were investigated by measuring the residual activities in the presence of each chemical. As shown in Figure 8E, the enzymatic activities of mCLEA-PsEstA were largely remarkably higher than those of the free form of PsEstA in the presence of the alcohols tested. Specifically, free PsEstA showed a relative activity of 21% in the presence of 10% EtOH, whereas mCLEA-PsEstA retained 47% of the initial activity. Moreover, although the free PsEstA retained less than 5% of its original activity in the presence of 30% EtOH, mCLEA-PsEstA showed 62% activity at the end. However, even 0.1% (v/v) SDS was sufficient to inactivate both free PsEstA and mCLEA-PsEstA almost completely. Similar behavior was also observed in the presence of 5 M urea. In reusability analysis, mCLEA-PsEstA retained~75% of its original activity after the 4th cycle, but lost a substantial level of activity at the 5th cycle ( Figure 8F). Taken together, immobilization of PsEstA (CLEAs-PsEstA and mCLEAs-PsEstA) was effectively performed, and these immobilized forms could function more effectively than the free form of PsEstA.
Biomolecules 2019, 9, x FOR PEER REVIEW 11 of 15 PsEstA and free PsEstA were investigated by measuring the residual activities in the presence of each chemical. As shown in Figure 8E, the enzymatic activities of mCLEA-PsEstA were largely remarkably higher than those of the free form of PsEstA in the presence of the alcohols tested. Specifically, free PsEstA showed a relative activity of 21% in the presence of 10% EtOH, whereas mCLEA-PsEstA retained 47% of the initial activity. Moreover, although the free PsEstA retained less than 5% of its original activity in the presence of 30% EtOH, mCLEA-PsEstA showed 62% activity at the end. However, even 0.1% (v/v) SDS was sufficient to inactivate both free PsEstA and mCLEA-PsEstA almost completely. Similar behavior was also observed in the presence of 5 M urea. In reusability analysis, mCLEA-PsEstA retained ~75% of its original activity after the 4th cycle, but lost a substantial level of activity at the 5th cycle ( Figure 8F). Taken together, immobilization of PsEstA (CLEAs-PsEstA and mCLEAs-PsEstA) was effectively performed, and these immobilized forms could function more effectively than the free form of PsEstA.

Formation of Organic-Inorganic Hybrid PsEstA Nanoflower
Recently, syntheses of organic-inorganic hybrid nanoflowers (hNFs) with greatly enhanced catalytic activities and stabilities have been reported [47,48]. Herein, the formation and catalytic activity of hybrid nanoflowers containing PsEstA were pursued. Scanning electron microscopy showed that the hybrid nanoflowers of PsEstA were hierarchical peony-like structures assembled from interlaced nanoplates ( Figure 9A). Reusability tests were also carried out for 10 cycles showing that the residual activity was~25% by the 3rd cycle ( Figure 9B).
Recently, syntheses of organic-inorganic hybrid nanoflowers (hNFs) with greatly enhanced catalytic activities and stabilities have been reported [47,48]. Herein, the formation and catalytic activity of hybrid nanoflowers containing PsEstA were pursued. Scanning electron microscopy showed that the hybrid nanoflowers of PsEstA were hierarchical peony-like structures assembled from interlaced nanoplates ( Figure 9A). Reusability tests were also carried out for 10 cycles showing that the residual activity was ~25% by the 3rd cycle ( Figure 9B).

Conclusions
Although family VIII esterases with β-lactamase activity have attracted considerable research interest, there is still limited information available regarding this enzyme family. Here, a novel family VIII esterase with β-lactamase activity (PsEstA) from Paenibacillus sp. was characterized using molecular modeling, spectroscopic methods, biochemical assays such as β-lactamase assay, and immobilization strategies. This structural and functional characterization of PsEstA is expected to provide a molecular platform for the comprehensive understanding of family VIII esterases as well as β-lactamase, although the physiological role of PsEstA still needs to be explored. In addition, considering the fact that very few enzymes of this family have been biochemically characterized and immobilized for biotechnological applications, PsEstA could be a promising target for biotechnological applications [49][50][51]. Further studies on PsEstA including mutagenesis of the key residues, in-depth kinetic analysis, and evaluation of enzyme-substrate complex formation will be necessary for a thorough understanding of this enzyme at the molecular level.

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

Conclusions
Although family VIII esterases with β-lactamase activity have attracted considerable research interest, there is still limited information available regarding this enzyme family. Here, a novel family VIII esterase with β-lactamase activity (PsEstA) from Paenibacillus sp. was characterized using molecular modeling, spectroscopic methods, biochemical assays such as β-lactamase assay, and immobilization strategies. This structural and functional characterization of PsEstA is expected to provide a molecular platform for the comprehensive understanding of family VIII esterases as well as β-lactamase, although the physiological role of PsEstA still needs to be explored. In addition, considering the fact that very few enzymes of this family have been biochemically characterized and immobilized for biotechnological applications, PsEstA could be a promising target for biotechnological applications [49][50][51]. Further studies on PsEstA including mutagenesis of the key residues, in-depth kinetic analysis, and evaluation of enzyme-substrate complex formation will be necessary for a thorough understanding of this enzyme at the molecular level.


extfieldsofstudy: Chemistry Medicine

text:
RNA binding activates RIG-I by releasing an autorepressed signaling domain

Direct monitoring of RIG-I activation by FRET reveals reversible activation by RNA alone.

INTRODUCTION
Retinoic acid-inducible gene I (RIG-I) is a human pattern recognition receptor (PRR) that is a promising therapeutic target (1). Like most PRRs, its activation leads to a potent immune response that may be useful if harnessed for immunooncology, vaccination, or antiviral therapy (2)(3)(4). Conversely, inappropriate activation of RIG-I can lead to autoimmune disorders (5). The pathogen-associated molecular pattern (PAMP) recognized by RIG-I has been identified as a 5′-triphosphorylated blunt duplex RNA (6). However, many lesseffective ligands have also been reported due to the complexity of the cellular immune response (7). PAMPs can have cell type-specific activity, cross-talk with other PRRs, contain impurities, or fall prey to a number of other caveats that produce misleading results in cell culture. Careful in vitro investigation of PRRs and their ligands is necessary to complement cell-based results and has led to notable advances in our understanding of RIG-I (8,9).
Many of the advances in our understanding of RIG-I have come from in vitro structural work. This includes structures of RIG-I bound to a minimal RNA ligand, confirming its recognition of a 5′-triphosphorylated blunt duplex RNA (10,11). The apo structure of RIG-I has been of particular interest because it revealed an autoinhibited conformation where the critical signaling domain (a pair of caspase recruitment domains, known as 2CARD) forms an interaction interface with the hel2i domain, effectively tucking it into the body of the protein (12). Upon activation of RIG-I, 2CARD is known to interact with downstream proteins (such as MAVS) to transmit the immunostimulatory signal (13)(14)(15)(16)(17). Therefore, it was predicted that the 2CARD/hel2i contact is broken upon RIG-I activation and that 2CARD is flipped into solution, making it sterically accessible for interaction with other proteins (12).
In vitro biochemical work largely agrees with the structural model for RIG-I activation, but there are issues that stem from shortcomings in available methodologies. Traditional RNA binding and adenosine triphosphatase (ATPase) techniques have been applied to RIG-I with great success (18)(19)(20)(21)(22). However, there are many steps in the RIG-I signaling cascade between these commonly measured in vitro activities and the signaling outputs that are measured in cell culture experiments. This has led to some inconsistencies between results from RNA binding, adenosine triphosphate (ATP) hydrolysis, and cell signaling experiments. For example, RNAs that bind RIG-I or stimulate ATP hydrolysis do not always signal in cells (18,20). Inconsistencies between these results hinder the identification and validation of proposed agonists and antagonists of RIG-I. In addition, these inconsistencies prevent a complete understanding of RIG-I function and its response to the binding of specific ligands. For example, ATP has been proposed to be involved in RIG-I proofreading, conformational rearrangement, translocation, and multimerization, and all these processes have been suggested to affect signaling by the protein (18,(23)(24)(25). However, the physical act of signal presentation by RIG-I (2CARD ejection) has not been directly measured and therefore cannot be differentiated from the many downstream events required for signal transduction.
2CARD ejection is necessarily the first step in RIG-I activation, but existing methods for measuring this conformational change have limitations. Hydrogen/deuterium exchange (HDX) and small-angle x-ray scattering (SAXS) have been used to monitor the relative position of 2CARD, but these methods require artificially high concentrations of reagents and have limited throughput (26)(27)(28). In addition, HDX reports subtle conformational changes that alter solvent accessibility, and one cannot distinguish these from large conformational changes. SAXS is sensitive to large conformational changes but is limited by its low resolution.
We set out to bridge the methodological gap between existing in vitro biochemical experiments and cell culture experiments. These efforts led to the creation of a validated assay for RIG-I activation using fluorescence resonance energy transfer (FRET). This FRET assay measures 2CARD ejection, which better represents the first stages of RIG-I activation than other in vitro techniques and is more readily controlled than cell-based techniques. The quantitative and facile readout of the assay also provides critical advantages over existing methods for monitoring the position of the 2CARD domain. Here, we have used the FRET assay to answer questions about the relative contributions of RNA and ATP to RIG-I activation.

Design of a FRET reporter for RIG-I activation
We designed a reporter for RIG-I activation that measures the position of the 2CARD domain relative to the helicase core (Fig. 1A). 2CARD directly contacts the hel2i domain in the autoinhibited state, but 2CARD is predicted to be moved significantly further away in the active conformation, allowing it to interact with downstream partners. While a complete structure of active RIG-I containing the 2CARD domain is lacking, we know that 2CARD is separated from hel2i by 275 amino acids, including a flexible 50-amino acid linker between 2CARD and the helicase core. Thus, RIG-I activation should lead to a change in distance between the 2CARD and hel2i domains that is both functionally relevant to signaling and large enough to be measured by FRET.
The design of the FRET construct involves incorporation of small organic fluorophores unlikely to perturb RIG-I function, and it requires the insertion of chemically orthogonal tags at internal positions of the protein. We FlAsH-labeled 2CARD by inserting a small tetracysteine motif (CCPGCC) in the disordered and poorly conserved linker immediately C-terminal to 2CARD ( Fig. 1 and fig. S1). This tetracysteine motif specifically coordinates the biarsenical fluorescein derivative FlAsH (29). The location of the tetracysteine tag was selected because it is solvent accessible, insertion of the tag is unlikely to perturb RIG-I function, and the distance to hel2i is minimal in the autoinhibited state. For attachment of the second fluorophore, we mutated amino acid Glu 494 in the hel2i domain to the unnatural amino acid azido-phenylalanine (AzF). This azide functionality allows specific labeling with alkyne-containing fluorophores via click chemistry. Like the tetracysteine tag, Glu 494 is in a disordered and evolutionarily divergent region and is proximal to 2CARD ( Fig. 1 and fig. S1). Thus, the incorporated tags are both solvent-exposed to facilitate labeling; they provide orthogonal chemical functionality; and their incorporation is unlikely to perturb RIG-I function.

Creation and labeling of the RIG-I FRET reporter
We encountered several challenges in the creation of the dual-labeled FRET reporter that required innovative steps to overcome. The azide group of AzF is prone to reduction to a nonreactive amine, so we decreased protein expression time and omitted reducing agents during purification (30). In addition, we performed click labeling before FlAsH labeling because the FlAsH labeling reaction requires reducing agents ( Fig. 2A). Traditional click labeling requires a copper catalyst that negatively affected RIG-I activity. However, a strained alkyne moiety, such as dibenzylcyclooctyne (DIBO), eliminates the need for copper in the click reaction. Several DIBO-modified fluorophores are commercially available, including Alexa Fluor 594, which has good spectral overlap with FlAsH. Copper-free click labeling with DIBO Alexa Fluor 594 was initially successful but was not specific for AzF-containing protein (Fig. 2B). We hypothesized that the nonspecific reaction was occurring via thiol-yne addition to cysteines, so we added a small amount of reducing agent and saw a marked increase in specificity without a significant reduction in labeling efficiency ( Fig. 2B) (31,32).
FlAsH labeling was also inefficient before the addition of precise amounts of 2-mercaptoethanol (ME) (Fig. 2C). FlAsH has the unique feature of being quenched in solution when bound to ethanedithiol (EDT), significantly reducing background signal from free fluorophore (33). This quenching is alleviated when the tetracysteine tag displaces EDT. However, ME also displaces EDT and increases background signal from free FlAsH (Fig. 2C). Thus, precise concentrations of ME and EDT were required to achieve specific and efficient FlAsH labeling.
Optimized labeling reactions produced active, dual-labeled RIG-I that exhibits a specific FRET signal in the autoinhibited state. After dual-labeling RIG-I, excess free fluorophore was removed ( Fig. 2A), and RNA binding activity was measured. Dual-labeled RIG-I binds RNA with a similar affinity to untagged wild-type protein (K D,WT = 2.2 ± 0.2 nM and K D,labeled = 1.2 ± 0.1 nM), suggesting that neither the tags nor fluorophores perturb RIG-I function ( fig. S2A). SDSpolyacrylamide gel electrophoresis (PAGE) analysis demonstrated that dual labeling requires incorporation of the tetracysteine and AzF tags, allowing us to attribute any FRET signal to the specifically tagged locations (Fig. 2D). The fluorescent emission spectrum upon excitation of FlAsH (donor) revealed a peak corresponding to Alexa Fluor 594 (acceptor) emission (Fig. 2E). This Alexa Fluor 594 signal was substantially higher than that observed for singly labeled proteins, suggesting that it derives from FRET, rather than spectral bleedthrough from direct excitation/emission of individual fluorophores. Unfortunately, some noncovalently bound Alexa Fluor 594 copurified with the dual-labeled protein, but controls using a protein with only the tetracysteine tag showed that the FRET signal does not derive from the copurified Alexa Fluor 594 ( fig. S2, B and C). Thus, we observe a FRET signal for active, dual-labeled RIG-I that specifically corresponds to the distance between the 2CARD and hel2i domains.

RNA binding ejects the 2CARD domain
RNA binding causes a reduction in FRET consistent with the ejection of 2CARD from the body of the protein (Fig. 2F). FRET was measured by quantifying the acceptor fluorophore emission upon excitation of the donor fluorophore. After correcting for bleed-through and loading, this FRET signal was normalized to maximum and minimum FRET states corresponding to the autoinhibited state and a denatured sample, respectively. Thus, a value of 1 corresponds to the high-FRET auto-inhibited state, and a value of 0 corresponds to a low-FRET state where the fluorophores are as far apart as possible. When an optimal RIG-I RNA ligand (SLR10) was added, the FRET signal decreased significantly, suggesting an increase in distance between the 2CARD and hel2i domains (Fig. 2F), consistent with the exceptional potency of this ligand (3). RNA binding did not significantly affect the fluorescence intensity of individually labeled proteins, suggesting that the FRET change derives from an increase in distance between fluorophores, rather than an artifact of fluorescence quenching or enhancement ( fig. S2D). The observed decrease in FRET occurred faster than could be detected with the current experimental setup (~5-s resolution), consistent with the on-rate (t 1/2 ≈ 0.1 s) that would be predicted for a highaffinity RNA such as SLR10 [K D ≈ 160 pM (19) and k off = 0.028 s −1 (18)] at a concentration of 100 nM with 50 nM protein. Nuclease digestion of the RNA caused an increase in FRET, back to the original level, demonstrating that the FRET change is RNA dependent and reversible. This is the first time that RIG-I activation has been shown to be reversible and that the protein can "reset" by restoring 2CARD to its original position in the inactive state. Next, we sought to determine the structural requirements for RNA-mediated 2CARD ejection. Other optimal RIG-I ligands (SLR14  Labeled protein was analyzed by SDS-polyacrylamide gel electrophoresis (PAGE), followed by fluorescent imaging (excitation/emission: 532 nm/LP575). In the absence of TCEP, negative control protein lacking AzF (−AzF) is nonspecifically labeled. (C) ME is required for efficient FlAsH labeling (top), but large amounts of ME cause nonspecific fluorescence by free FlAsH (bottom). One micromolar ME was identified as an ideal concentration that is sufficient for labeling without causing background fluorescence. and SLR30) eject 2CARD to similar extents, whereas RNAs that do not bind RIG-I (U55 and pppNS) do not eject 2CARD, further demonstrating the specificity of the assay (Fig. 3). SLR10, SLR14, and SLR30 are 5′-triphosphorylated blunt duplex RNA hairpins that contain 10, 14, and 30 base pairs (bp), respectively. A slightly shorter RNA hairpin, SLR8, binds with high affinity, but does not signal in cell culture, exemplifying the limitations of existing in vitro binding assays (20). SLR8 would be expected to partially clash with 2CARD, if bound in the same manner as longer RNAs. The FRET assay reveals that SLR8 inefficiently ejects 2CARD, suggesting the formation of a protein-RNA complex distinct from the structurally characterized conformations and providing a mechanistic explanation for the inability of this ligand to signal in cells. Thus, an RNA stem longer than 8 bp is required for 2CARD ejection, but additional increases beyond 10 bp do not affect 2CARD ejection.
We also tested a 5′-triphosphorylated RNA equal in length to SLR10 but lacking the duplex structure (pppNS). pppNS binds RIG-I weakly with a low-micromolar K D and does not activate RIG-I in vivo (3,19). The FRET assay uses nanomolar concentrations of protein and RNA, meaning that pppNS would not be expected to bind, and we see that pppNS does not cause 2CARD ejection (Fig. 3). However, pppNS can cause 2CARD ejection at low-micromolar concentrations, consistent with its K D (fig. S3). These results demonstrate the utility of the FRET assay over other assays that require higher concentrations of reagents. The FRET assay is uniquely able to differentiate between high-and low-affinity RIG-I ligands because of the low concentrations of reagents it requires. These observations further suggest that, above a certain RNA length, the structural requirements for 2CARD ejection are lax and that RIG-I activation in vivo is largely dictated by its relative affinity for a given RNA.
Last, we tested how changes to the 5′ end of the RNA affect 2CARD ejection. OH-SLR14 has the same blunt duplex as SLR14, but it has a 5′-OH in place of the triphosphate. This RNA has decreased affinity for RIG-I and does not activate RIG-I in vivo (3,19). However, RIG-I binds OH-SLR14 with moderate affinity (low-nanomolar K D ), and OH-SLR14 can activate RIG-I mutants that are deficient in their ability to dissociate from RNA (34). We observe that OH-SLR14 ejects 2CARD when bound to RIG-I (Fig. 3). It is important to note that the concentrations of protein and RNA used in this assay are greater than the K D for OH-SLR14, ensuring that RIG-I is fully bound to OH-SLR14. Thus, the 5′ triphosphate is not strictly required for 2CARD ejection, as long as RNA binding can still occur. Similarly, polyinosinic:polycytidylic acid [poly(I:C)] lacks a 5′ triphosphate and is capable of ejecting 2CARD (Fig. 3). However, by mass, more poly(I:C) than SLR10 is required to eject 2CARD, consistent with the fact that poly(I:C) has a lower concentration of blunt duplex ends than SLRs (Fig. 3) (20). This further suggests that the internal duplex binding sites present in poly(I:C) are not capable of binding RIG-I and ejecting 2CARD at the concentrations used in this assay. In summary, RNA duplexes greater than 8 bp are sufficient for 2CARD ejection, and additional structural features, such as a blunt terminus and 5′ triphosphate moiety, serve to increase binding affinity.
ATP has little effect on 2CARD ejection ATP binding also plays a role in RIG-I signaling; thus, we tested the effects of ATP and various analogs on the conformation of the 2CARD domain. The ability of RIG-I to bind ATP correlates with its ability to signal in cells (18). In addition, the inability of RIG-I to hydrolyze ATP leads to its overactivation (5,22). These observations led to the hypothesis that, following RNA binding, ATP binding promotes signaling by enhancing 2CARD ejection (18,35). However, we observed no major change in 2CARD ejection upon the addition of nonhydrolyzable ATP analogs (AMP-PNP and ATPS), which should mimic an ATP-bound conformation of RIG-I (Fig. 4). Adenosine diphosphate (ADP) and ATP do not influence 2CARD ejection, but the transition-state analog ADP-AlF x appears to cause a subtle decrease in FRET. ATP analogs have similarly minor effects in the presence of SLR30 and the absence of RNA, further demonstrating that RNA length does not affect 2CARD ejection ( fig. S4, A and B). After repeating the ADP-AlF x experiment, the decrease in FRET is significant but subtle ( fig. S4C). The normalized FRET value with ADP-AlF x is greater than 0 (the level of denatured protein), suggesting that the FRET assay is still within its dynamic range. Therefore, ADP-AlF x causes a reproducible but small change in the position of the 2CARD domain.

DISCUSSION
Here, we have used a novel assay to reveal key features about the interplay between RNA and ATP in RIG-I activation. First, we have provided important insight into how RIG-I avoids unwanted activation by self-RNA and how this might become misregulated in autoimmune diseases, such as Singleton-Merten syndrome (SMS) (5). We have shown that activation of RIG-I is not strictly dependent on the 5′ triphosphate moiety, suggesting that self-RNA is capable of activating RIG-I under misregulated conditions. Such misregulation could occur for a variety of reasons, including an increase in cellular RNA concentration or misregulation of RIG-I binding. For example, patients with SMS have mutations in RIG-I that block its ATPase activity, preventing dissociation of RIG-I from self-RNA (5,18,22,36). However, there was previously no direct evidence that this dissociation from RNA prevented signaling. Here, we provide the first direct evidence that the 2CARD signaling domain rapidly returns to its autoinhibited state after RNA dissociation, supporting the current model that SMS is caused by impaired ATP hydrolysis, impaired dissociation of RIG-I from self RNA, and, ultimately, impaired sequestration of the 2CARD domain (18,22,36). The ability of 2CARD to reset after dissociation from RNA has implications for mechanisms of downstream signaling. One model for RIG-I signaling involves the formation of a homotetramer via the 2CARD domain (37). In the case of SLR14, this would require the simultaneous encounter of four individual protein-RNA complexes. This process would be extremely inefficient if RIG-I were also dissociating from RNA and reverting to its autoinhibited state, yet SLR14 is a potent RIG-I agonist in vivo (3). There is also evidence that ubiquitination of the 2CARD domain promotes RIG-I signaling, especially on short RNAs (16,38). Thus, it is tempting to speculate that ubiquitination and, perhaps, other modifications of the 2CARD domain prevent 2CARD reset, trapping RIG-I in an active state and facilitating the formation of a tetramer.
It is particularly significant that RNA alone is sufficient to cause 2CARD ejection and that presentation of 2CARD does not require ATP. These observations are important in light of the many different functions that have been ascribed to ATP binding and hydrolysis by RIG-I (18,(23)(24)(25), and they underscore the primary role of RNA binding during activation of the RIG-I sensor. They explain the observation that short (10 to 14 bp) triphosphorylated RNA ligands are particularly potent RIG-I agonists and that these small RNAs can stimulate signaling even by ATPase-deficient RIG-I mutants (18). However, they do not explain the fact that, although RIG-I can bind tightly to the terminus of any triphosphorylated RNA duplex, regardless of length (20), it can only signal from longer double-stranded RNAs (dsRNAs) if ATP-binding activity is maintained (18). One explanation is that the 2CARD ejection caused by RNA alone might be insufficient for signaling. In the presence of ADP-AlF x , there is a small but observable decrease in FRET signal, and it is known that ADP-AlF x can mimic a transient transition state during ATP hydrolysis that may somehow populate a conformation that assists in CARD presentation. However, the dispensability of ATP hydrolysis for RIG-I signaling in cells undercuts this model (18,22). An alternative explanation is that RNA alone is sufficient to eject 2CARD and activate RIG-I in a pure system and on short RNAs in cells but that longer RNAs such as SLR30 are coated by cellular factors that repress signaling by ATPase-deficient RIG-I mutants. In this case, the ATPase activity of RIG-I may have a second function, enabling RIG-I to displace competing cellular factors to signal from the dsRNA terminus (39).
Shorter RIG-I agonists, such as SLR10 and SLR14, are still able to induce cell signaling in ATPase-deficient RIG-I mutants because they may lack binding sites for additional cellular factors and/or the tightly bound RIG-I molecule that caps the short duplex terminus does not leave any room for competing factors to bind. On the basis of the available data from all laboratories, we favor a model in which ATP plays no substantive role in 2CARD presentation or for the conformational changes needed for ejecting 2CARD from its Hel2i-binding site, but that ATP may be required for displacement of competing proteins and (as has now been demonstrated by multiple laboratories) for proofreading of inappropriate RIG-I targets such as host RNA (18,22,23,26,34,35).
That said, if RIG-I is capable of displacing proteins from long RNA targets, it is expected to proceed through the winch-like mechanism that is commonly observed for protein displacement by DEAD-box proteins (40), as RIG-I and related ATPases are structurally and phylogenetically related to DEAD-box proteins (41) rather than processive viral helicases (such as NPH-II) (42). Evolution of the pincer domain in RIG-I-like proteins resulted in a conformational alteration that prevents translocation via the mechanism established for SF2 proteins such as NS3 (43), perhaps explaining slow rates of apparent RIG-I motion on RNA (23) and suggesting a nonprocessive mechanism much like that reported for related DEAD-box RNPases (40).
Recent publications have also reported 2CARD ejection by RNA and ATP analogs using HDX and SAXS, but the FRET assay adds important quantitative information that leads us to a slightly different conclusion (26)(27)(28). Using HDX, researchers observed an increase in 2CARD solvent exposure upon binding RNA (28). Like the FRET assay, HDX reports an additional increase in solvent exposure upon the addition of ADP-AlF x (28). However, the extent of the conformational change upon the addition of ADP-AlF x was unclear, and it is curious that no other ATP analog showed this effect. On the basis of these data, the authors concluded that ATP plays a significant role in RIG-I activation (28). SAXS, on the other hand, reports a quantitative increase in molecular size upon RNA and ADP-AlF x binding, but the domains responsible for this increase are ambiguous, and again, the effect was only observed with ADP-AlF x (27). These results led to the conclusion that ATP is important for CARD ejection, but the conformational changes were small and cannot be unambiguously attributed to motion of the 2CARD domain. FRET provides a more specific and quantitative metric for 2CARD ejection. This new metric reveals substantial 2CARD ejection upon RNA binding and a relatively small change caused by ADP-AlF x . In combination with the body of work described above, RNA alone appears sufficient for 2CARD ejection and activation of RIG-I.
The FRET assay has certain technical advantages over other 2CARD ejection assays that aid interpretation of results and will benefit future studies. The FRET assay uses low-to mid-nanomolar concentrations of protein and RNA, whereas structural studies such as HDX and SAXS use much higher concentrations in the low-to mid-micromolar range. The lower concentrations required for FRET are more biologically relevant and less prone to artifacts, such as homodimerization of RNA hairpins and protein multimerization. The low concentrations made it possible to differentiate between high-and low-affinity binding events, such as the difference between SLR and pppNS binding. The FRET assay can also be adapted for future studies of RIG-I that would be intractable with HDX or SAXS. First, FRET is more accessible than HDX and SAXS as it requires only a fluorometer, rather than a mass spectrometer or x-ray source. Second, the readout for the FRET assay is rapid and facile, which allows for use in screening and kinetic experiments. Last, this FRET construct could be used in single-molecule experiments to identify transient or lowly populated states involved in RIG-I activation (44). Unlike HDX and SAXS, FRET requires careful design and validation of a construct, which we have performed here. We placed small nonperturbing fluorophores at locations as proximal as possible in the high-FRET state. There are relatively few methods able to attach small fluorophores at specific internal protein locations, and these methods are further restricted for large and unstable proteins such as RIG-I. Several labeling strategies require pH, temperatures, or reagents that adversely affected RIG-I activity. FlAsH and AzF were ultimately successful because they require mild conditions and commercially available reagents. Here, we have optimized these strategies for use together, creating a dual-labeling strategy that is applicable to other large proteins.
We have created and used a novel method to address the unmet need for a biochemical assay that measures RIG-I activation in vitro. Previous biochemical studies of RIG-I have focused on RNA binding and ATP hydrolysis activities, but these activities do not necessarily correlate with RIG-I activation. There are many steps between RNA binding/ATP hydrolysis and signaling in a cell; thus, it is expected that the correlations between these activities are imperfect. Our 2CARD ejection assay measures a structural change more closely related to signaling and, therefore, more closely reports activation of RIG-I.Our FRET assay can be adapted for a variety of experiments on RIG-I, and similar methodology could be applied to homologous proteins, such as MDA5. In addition, we have provided a roadmap for the site-specific dual labeling of large proteins that will facilitate similar studies in many biological systems.

Cloning and protein construct creation
Full-length human RIG-I was previously cloned into a pET SUMO expression vector (19). This construct contains an N-terminal 6-His tag, followed by a SUMO tag and the complete human RIG-I protein sequence. A serine residue was also added between the SUMO tag and RIG-I to ensure efficient tag cleavage.
In this study, we added the amino acid sequence MPCCPGCCGS following RIG-I residue 190. This sequence was added using Q5 mutagenesis (New England Biolabs) according to the manufacturer's instructions. AzF was incorporated by amber stop codon suppression. Thus, the wild-type amino acid Glu 494 was mutated to an amber stop codon (UAG) by the QuikChange Site-Directed Mutagenesis Kit (Agilent) according to the manufacturer's recommendations.

RIG-I expression and purification
RIG-I was expressed and purified essentially as previously described (19). Plasmid was transformed into Rosetta (DE3) competent cells (Novagen). Cells were grown at 37°C in Luria broth (Research Products International) to an optical density at 600 nm (OD 600 ) of 0.6. Upon reaching 0.6, cells were cold-shocked on ice for 20 min and then returned to a 16°C incubator. Protein expression was induced with 0.3 mM isopropyl--d-thiogalactopyranoside (IPTG), and cells were grown for 16 to 24 hours. Cells were harvested by centrifugation and resuspended in 10-ml lysis buffer per gram of cell paste [25 mM K-Hepes, 300 mM NaCl, 5 mM ME, 5 mM imidazole, 10% glycerol (pH 8.0)]. One EDTA-free protease inhibitor tablet (Roche) was added, and cells were lysed by passing three to five times through a microfluidizer. Lysate was clarified by centrifugation at 15,000g for 30 min. Clarified lysate was batchincubated with Ni-NTA agarose beads for 1 hour, washed with 5 column volumes wash buffer [25 mM K-Hepes, 1 M NaCl, 30 mM imidazole, and 10% glycerol (pH 8.0)], and eluted [25 mM K-Hepes, 300 mM NaCl, 5 mM ME, 250 mM imidazole, and 10% glycerol (pH 8.0)]. Eluant was diluted 1:1 with elution buffer lacking imidazole and incubated with Ulp1 SUMO protease overnight at 4°C. Cleaved protein was further diluted 1:1 with 25 mM K-Hepes, 5 mM ME, and 10% glycerol (pH 8.0) and loaded onto a HiTrap Heparin HP column (GE Healthcare). The column was washed with 25 mM K-Hepes, 150 mM NaCl, 10% glycerol, and 5 mM ME (pH 8.0) and eluted with 25 mM K-Hepes, 650 mM NaCl, 10% glycerol, and 5 mM ME (pH 8.0). Eluent from the heparin column was concentrated and further purified on a HiLoad Superdex 200-pg column (GE Healthcare) equilibrated in 25 mM K-Hepes, 150 mM NaCl, and 5% glycerol (pH 7.5). Monomeric protein was collected, concentrated, flash-frozen in liquid nitrogen, and stored at −80°C.
RIG-I containing AzF was expressed and purified in a similar manner with several alterations. BL21(DE3) Escherichia coli were transformed with 200 ng of pSUMO-RIG-I containing an amber stop codon and 200 ng of pEVOL-pAzF. pEVOL-pAzF was a gift from P. Schultz (Addgene plasmid #31186; http://n2t.net/addgene:31186; RRID:Addgene_31186) (45). Cells were grown to an OD 600 of 0.4, the temperature was decreased to 16°C, and 4-azido-l-phenylaline (0.25 g/liter) was added. l-Arabinose (0.02%) was added after 5 min. Cells were further grown to an OD 600 of 0.6 when 0.3 mM IPTG was added, and cells were grown for 8 to 9 hours. Cells were harvested, and protein was purified as described for wild-type RIG-I except that exposure to light was limited and ME was omitted to prevent reduction of the azide.
SLR30 was transcribed in vitro from a double-stranded synthetic DNA template (Integrated DNA Technologies) containing 2′-OMe modifications on the last two nucleotides of the template strand. One microgram of annealed template was used in a 100-l transcription reaction containing 5 mM each ribonucleoside triphosphate, 22 mM magnesium chloride, 40 mM tris-HCl (pH 8.0), 2 mM spermidine, 10 mM dithiothreitol (DTT), 0.01% Triton X-100, 40 U of ribonuclease (RNase) inhibitor (New England Biolabs), and 5 l of T7 RNA polymerase. Transcription reactions were incubated at 37°C for 2 to 12 hours.
All RNA was gel-purified by denaturing gel electrophoresis. Bands were excised, and RNA was extracted by crush soak in 10 mM Mops (pH 6.0), 300 mM NaCl, and 1 mM disodium-EDTA. RNA was ethanolprecipitated, washed once with 70% ethanol, and dissolved in water. RNA purity was confirmed by mass spectrometry (Novatia LLC). Low-molecular weight poly(I:C) was purchased from Invivogen and used without further purification.

Fluorophore conjugation
RIG-I containing the tetracysteine tag and E494AzF mutation was diluted to 25 M in 25 mM K-Hepes and 150 mM NaCl (pH 7.5).
Unless otherwise indicated, 1 mM TCEP-HCl (pH 7.5) was added, immediately followed by 50 M Click-IT Alexa Fluor 594 DIBO Alkyne (Thermo Fisher Scientific) dissolved in dimethyl sulfoxide (DMSO). The final DMSO concentration in the click labeling reaction was less than 5%. The reaction was incubated at 4°C overnight.
FlAsH labeling was performed after overnight click labeling. One millimolar TCEP-HCl, 1 mM ME, 1 mM EDT, and 70 M FlAsH-EDT 2 were added, and the reaction was incubated for 2 hours at 4°C. Excess free fluorophore was removed using a 40,000 molecular weight cutoff Zeba desalting column (Thermo Fisher Scientific) equilibrated with 25 mM K-Hepes, 150 mM NaCl, and 1 mM TCEP-HCl (pH 7.5).
After labeling and desalting, concentrations of protein and fluorophores were calculated by spectrophotometry using the extinction coefficients of  515 = 41,000 M −1 cm −1 and  280 = 4100 M −1 cm −1 for FlAsH,  590 = 73,000 M −1 cm −1 and  280 = 40,900 M −1 cm −1 for Alexa Fluor 594, and  280 = 99,700 M −1 cm −1 for RIG-I. Samples were also analyzed by SDS-PAGE using a 12% acrylamide tris-glycine gel and loading dye that contained 50 mM TCEP, rather than ME, to prevent dissociation of the FlAsH fluorophore. Fluorescent imaging of the gels was performed on a Typhoon FLA 9500 biomolecular imager (GE Healthcare), followed by Coomassie staining. The intensities of bands were quantified manually in ImageQuant (GE Healthcare), and labeling specificity was calculated as the ratio of fluorescence intensity for tagged protein over untagged protein. Labeling efficiency was calculated as the concentration of fluorophore divided by the concentration of protein, as measured by spectrophotometry. Copurified free Alexa Fluor 594 was corrected for by measuring the fraction of total fluorescent signal on the SDS-PAGE gel derived from the protein band.

RIG-I CARD ejection assay
Dual-labeled RIG-I was mixed with RNA in a black 96-well nonbinding plate (Corning). ATP analogs were added, as indicated, and reactions were equilibrated briefly on ice. ADP, ATP, and AMP-PNP were purchased from Sigma-Aldrich, and ATPS was purchased from Cayman Chemical. A 3× stock of ADP-AlF x was created by mixing 6 mM ADP, 30 mM NaF, and 6 mM AlCl 3 , followed by 0.2-m filtration. Final reactions contained 25 mM K-Hepes (pH 7.5), 75 mM NaCl, 75 mM KCl, bovine serum albumin (0.1 mg/ml), 0.3 mM TCEP-HCl, 50 nM RIG-I, 100 nM RNA, and 2 mM MgCl 2 /ATP analog. Reactions were set up at least in triplicate, and measurements were collected 5, 15, 30, 45, and 60 min after mixing. We observed no change in fluorescence over time; thus, readings for individual wells were averaged across time points. Reported error corresponds to the SE between replicate wells. RNase digestion of RNA was achieved by adding 500 U of benzonase (EMD Millipore) and 2 mM MgCl 2 , followed by incubation on ice for 30 min. Fluorescence was measured using a BioTek Synergy H1 plate reader with excitation/emission wavelengths of 479/615 nm ("FRET signal"), 479/532 nm ("donor signal"), and 580/615 nm ("acceptor signal").
Proteins labeled with donor only or acceptor only were used to calculate the contribution of bleed-through to the FRET signal. Protein labeled with donor only had a bleed-through FRET signal that was 2.5% of the donor signal. Protein labeled with acceptor only had a bleed-through FRET signal that was 3.2% of the acceptor signal. Using these calibration values, a "subtracted FRET signal" was calculated using Eq. 1 Subtracted FRET = FRET signal − 0.025 * donor signal − 0.032 * acceptor signal (1) Variations in loading were accounted for by normalizing this subtracted FRET signal to the acceptor signal according to Eq. 2 Relative FRET = subtracted FRET ─ acceptor signal (2) Last, the "relative FRET" value for each sample was normalized to a sample without RNA (maximal FRET) and a sample treated with 6 M urea (minimal FRET) to calculate a "normalized FRET" value according to Eq. 3 Normalized FRET = sample relative FRET − urea relative FRET ──────────────────── no RNA relative FRET − urea relative FRET (3)

Fluorescence anisotropy binding assay
Binding experiments were performed as described previously (18). The RNA used in these experiments consists of OH-SLR10 with Alexa Fluor 488 attached to an amine-modified uracil in the tetraloop (table S1). RNA was diluted in binding buffer [25 mM Mops (pH 7.4), 150 mM NaCl, 5 mM DTT, 2 mM MgCl, and 0.01% Triton X-100] to a concentration of 500 pM. RIG-I protein was serially diluted in binding buffer and mixed 1:1 with RNA in a 384-well nonbinding plate (Corning). Volumes were at least 20 l. Plates were covered and incubated on ice for at least 2 hours until reactions reached equilibrium. Fluorescence polarization was measured using a Synergy H1 plate reader (BioTek). Samples were excited using a bandpass filter at 485/20 nm, and fluorescence emission was measured using a bandpass filter at 528/20 nm. Polarization was calculated using Eq. 4 Polarization = I II − G * I ⊥ ─ I II + G * I ⊥ (4) where I II is the intensity of light parallel to the excitation plane, I ⊥ is the intensity of light perpendicular to the excitation plane, and G is a correction factor empirically determined to be 0.87. Protein dilutions were performed in triplicate, and averages were plotted against protein concentration. These data were fit to Eq. 4 using GraphPad Prism to calculate the dissociation constant where y is the polarization value, y 0 is the polarization value without protein, y max is the polarization value at saturating RIG-I concentration, K D is dissociation constant, and x is the concentration of RIG-I.

SUPPLEMENTARY MATERIALS
Supplementary material for this article is available at http://advances.sciencemag.org/cgi/ content/full/5/10/eaax3641/DC1 Fig. S1. Tags were placed in locations designed to facilitate labeling and avoid perturbation of RIG-I function. Fig. S2. Controls demonstrate that dual labeling of RIG-I does not perturb its function, and FRET reports the distance between hel2i and 2CARD. Fig. S3. pppNS ejects 2CARD at high concentrations. Fig. S4. ADP-AlFx subtly but reproducibly increases 2CARD ejection. Table S1. RNA sequences used in this study. References (50,51)


The effects of the metformin on inhibition of UVA-induced expression of MMPs and COL-I in human skin fibroblasts

This study was to investigate the effects of metformin (MF) on ultraviolet A (UVA)-induced expression of matrix metalloproteinases (MMPs) and type I collagen (COL-I) in human skin fibroblasts (HSFs). HSFs were cultured in vitro and divided into control group, UVA group, and UVA + MF group. Cell proliferation was detected by CCK-8 method, and intracellular reactive oxygen species (ROS) level was detected by flow cytometry with fluorescent probe 2′,7′-dichlorofluorescein diacetate (DCF-DA) staining. Meanwhile, real-time polymerase chain reaction (PCR) was used to examine the relative messenger RNA (mRNA) expression of aging-related genes, including MMP1, MMP3, and COL-I. Moreover, the expression of MMP1, MMP3, and COL-I proteins was further detected by western blot. Compared with the control group, the ROS content in the UVA group was increased significantly (P < 0.05), while the ROS content in the UVA + MF group was evidently lower than that in the UVA group (P < 0.05). In addition, the MMP1 and MMP3 mRNA levels were significantly elevated, while the COL-I mRNA was significantly decreased in UVA-induced HSF cells compared with the control cells. However, MF could significantly inhibit the improved MMP1 and MMP3 mRNA level and increase the COL-I mRNA level. Moreover, MF could significantly reverse the increasing MMP1 and MMP3 protein level and decreasing COL-I protein level induced by UVA. In conclusion, MF can increase the antioxidant capacity of cells and increase the synthesis of COL-I by inhibiting the level of intracellular ROS and the expression of related MMPs, thereby inhibiting the UVA-induced photoaging effect of HSF.

Introduction
It was demonstrated in the related data 1 that the extracellular matrix (ECM) of the skin is mainly composed of type I collagen (COL-I) secreted by human skin fibroblasts (HSFs), which plays an important role in maintaining skin tension and tolerance. Sunlight is an important factor in inducing photoaging of the skin. A large amount of ultraviolet A (UVA) can penetrate the dermis layer of the skin, causing photoaging phenomenon of skin such as skin damage, cutis laxa, and wrinkles. 2 The photoaging of the skin induced by ultraviolet radiation is mainly manifested in the increase of skin wrinkles, decreased elasticity, and pigmentation, which is associated with increased secretion of matrix metalloproteinases (MMPs) and decreased synthesis of COL-I caused by ultraviolet B (UVB) irradiation. 3 Metformin (MF) is a widely used anti-diabetic drug, and it is further verified in the recent studies that MF possessed the effects of reducing cancer incidence, anti-aging, and prolonging life. 4 MF can inhibit the activation of I kappa B kinase (IKK) kinase and reduce the occurrence of chronic inflammatory reactions by reducing the production of mitochondrial reactive oxygen species (ROS), thereby inhibiting aging. 5 This study was to observe whether MF can protect against UVA-induced photoaging of HSF, so as to provide a basis for the application of MF in photoaging prevention.

CCK-8 assay for cell proliferation
HSF was provided by Shanghai Aolu Biological Technology Co., Ltd. and cultured in a high-glucose DMEM (Dulbecco's Modified Eagle Medium; Beijing Qingda Tianyi Technology Co., Ltd.) containing 10% fetal bovine serum (ScienCell Co., Ltd., Carlsbad, CA, USA). This study was approved by the Ethics Committee of Shunde Hospital Affiliated to Guangzhou University of Traditional Chinese Medicine. The cells were plated at a density of 2000 cells/well in a 96-well plate and cultured at 37°C under 5% CO 2 condition. After adherence, a concentration of 0.01 mmol/L of MF (Taize Ruida Technology Co., Ltd., Beijing, China) was added, and after 72 h of culture, CCK-8 reagent was added and operated according to the kit instructions (Aviva Systems Biology, Beijing, China). The absorbance (A) value was measured with a microplate reader at an excitation wavelength of 450 nm. The experiment was repeated three times separately.

Detection of ROS level by flow cytometry
The cell culture conditions are the same as "CCK-8 assay." And the cells were seeded at a density of 10 5 cells/well in a six-well plate and divided into control group, UVA group, and UVA + MF group. The UVA + MF group was treated with 0.01 mmol/L MF, while the control group and UVA group were treated with DMEM medium instead. After 24 h of drug treatment, the culture solution was removed, and then the cells were washed three times with phosphate-buffered saline (PBS). Finally, the UVA group and UVA + MF group were added with 500 μL of PBS and irradiated at a dose of 9 J/cm 2 . 6 The control group did not receive any dose of irradiation. After the irradiation, the UVA + MF group was further cultured by adding 0.01 mmol/L of MF. After 72 h of culture, the cells were collected by trypsinization, then ROSsensitive fluorescent probe, 2′,7′-dichlorofluorescein diacetate (DCF-DA; Sigma Aldrich, Germany), was added, followed by incubation at 37°C for 30 min. It was washed three times with PBS. A FACScan flow cytometer and FACSCanto II version 4.1 (BD Biosciences, San Jose, CA, USA) were used to perform the detection. The experiment was repeated three times separately.

Real-time polymerase chain reaction method for detecting relative expression of MMP1, MMP3, and COL-I mRNA
Cell culture and treatment are the same as "flow cytometry assay." After 72 h of culture, the cells were collected by trypsinization, and total RNA was extracted, purified, and quantified. The RNA was reverse transcribed into complementary DNA (cDNA) according to the instructions of the reverse transcription kit, pre-denatured at 95°C for 5 min, denatured at 95°C for 15 s, annealed at 60°C for 30 s, extended at 72°C for 30 s, 40 cycles, and maintained at 72°C for 10 min. β-actin was served as the internal control, and the expression levels of MMP1, MMP3, and COL-I genes were calculated based on the cycle threshold (Ct). The primer sequences are shown in Table 1 (GenePharma Co., Ltd, Shanghai, China). The experiment was repeated three times separately.

Western blot analysis of MMP1, MMP3, and COL-I protein expression
Cell culture and treatment are the same as "flow cytometry assay." After 72 h of culture, the culture medium was discarded. And the radioimmunoprecipitation assay (RIPA) lysis buffer was added after washing with PBS. Then, the cells were placed on the ice for 30 min, were centrifuged at 4°C, 12,000 r/min for 15 min, and the total protein was extracted and quantified. For sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), the protein in the gel was transferred to the nitrocellulose (NC) membrane using a transfer electrophoresis chamber at a constant current of 280 mA for 90 min, and then blocked in 5% skim milk for 2 h, followed by incubation with a MMP1 antibody, MMP3 antibody (1:1000; Proteintech Group, Inc.) or COL-I antibody (1:1000; SinoMab BioScience Limited) at 4°C overnight. After washing three times with TBST, the secondary antibody was added and incubated at 37°C for 1 h. After washing 3 times with TBST (Tris-buffered saline with Tween 20), ECL (chemiluminescent agent) was added. The bands were exposed under a chemiluminescence detection kit (Nanjing KeyGen Biotech Co. Ltd., Jiangsu, China) and analyzed by Image J version 1.8.0 (National Institutes of Health, Bethesda, MD, USA) and normalized to β-actin. The experiment was repeated three times separately.

Statistical analysis
All data were analyzed by SPSS 21.0 analysis software. The data were expressed as mean ± standard deviation ( ) x SD ± . Comparison between the groups was analyzed using one-way analysis of variance (ANOVA) analysis. The P value of <0.05 indicated that the difference was statistically significant.

Effect of MF on proliferation of HSF cells
To choose the suitable concentration of MF in HSF cells, CCK-8 assay was adopted to detect the effect of different concentrations of MF on cell proliferation. As shown in Figure 1(a), the results showed that concentration of MF at 0.01 mmol/L exerted no significant cytotoxic effect after 72 h of treatment. However, concentrations of MF at 0.1 and 1 mmol/L resulted in a marked decrease in cell proliferation. Therefore, we finally chose 0.01 mmol/L as the concentration of MF in this study.

MF can reduce ROS level in HSF cells irradiated by UVA
Flow cytometry was used to determine whether MF can reduce UVA-induced ROS level. Compared with the control group, the ROS content in the UVA group was increased significantly. However, the upregulation of ROS level was significantly attenuated by MF, as shown in Figure 1(b).

Effects of MF on mRNA and proteins expression of MMP1, MMP3, and COL-I in HSF cells irradiated by UVA
To investigate the role of MF on the expression of MMP1, MMP3, and COL-I mRNA in UVA-treated HSF cells, real-time polymerase chain reaction (RT-PCR) method was performed in this study. The MMP1 and MMP3 mRNA levels were significantly elevated, while the COL-I mRNA was significantly decreased in UVA-induced HSF cells compared with the control cells. However, MF could significantly inhibit the improved MMP1 and MMP3 mRNA level and increase the COL-I mRNA level, as shown in Figure 1(c). Moreover, western blot analysis was carried out to further explore the function of MF on the expression of MMP1, MMP3, and COL-I proteins against UVA. Interestingly, MF could significantly reverse the increasing MMP1 and MMP3 protein level and decreasing COL-I protein level induced by UVA, as shown in Figure 1(d) and (e).

Discussion
MF, as a first-line oral drug for T2DM, has a safe and effective hypoglycemic effect. In addition, MF can regulate the mannose transport activity of dermal fibroblasts through AMPK (5′ AMP-activated protein kinase) 7 and inhibit the carcinogenic effect of benzopyrene on mouse skin. Moreover, it also inhibits the carcinoma cell proliferation of squamous cell by inhibiting nuclear factor kappa B (NF-κB) and mammalian target of rapamycin (mTOR) signaling pathways. Long-term chronic repeated exposure to ultraviolet light can cause skin photoaging, which was manifested as slack, dryness, roughness, and wrinkle increase. 8 The phenomenon of photoaging is related to the decrease of human dermal fibroblast (HDF) and the decline or abnormality of secretory and synthetic functions. Repeated UVA irradiation can accumulate ROS in HDF cells, which exceeds their clearance capacity and breaks the balance between oxidation and anti-oxidation. Consequently, it causes oxidative stress to occur and regulates a series of programmed cellular responses, even regulating aging-related signaling pathways, thereby promoting cellular senescence. The intracellular accumulation of ROS is an important part of the development of skin photoaging. 9 In this study, MF can eliminate intracellular ROS.
Collagen is the main component of skin protein and one of the most important parameters for evaluating the aging state of HDF. COL-I is the most important collagen component in the dermis. In the pathological state, the synthetic decomposition loses the balance and leads to collagen abnormality. MMPs are one of the most important factors involved in the decomposition mechanism. In the process of photoaging, MMPs can degrade COL-I, thereby attributing to the reconstruction of skin connective tissue structure and aggravating skin aging. 10 Ultraviolet radiation will increase the secretion of MMPs in the skin. UVB irradiation can cause the cells to produce a large amount of ROS, which can increase the phosphorylation of the membrane surface of the cell, thereby activating the mitogen-activated protein kinase (MAPK) pathway and activator protein-1 (AP-1) signal pathway, consequently leading to increased secretion of MMP-1, MMP-3, and elevated degradation of collagen. 11 In this experiment, MF could significantly reverse the increasing MMP1 and MMP3 mRNA and protein level and decreasing COL-I mRNA and protein level induced by UVA.
In conclusion, MF can improve the antioxidant capacity of cells and increase the synthesis of COL-I by inhibiting the expression of intracellular ROS and related MMPs, thereby inhibiting the photoaging effects of UVA-induced HDF cells. This study provides a certain experimental basis for the anti-aging effect of MF in delaying the photoaging of skin.

Declaration of conflicting interests
The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding
The author(s) received no financial support for the research, authorship, and/or publication of this article.