
==== Front
J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02148-3
10.1016/j.jbc.2024.107647
107647
Research Article
Curcumin degradation in a soil microorganism: Screening and characterization of a β-diketone hydrolase
Hashimoto Yoshiteru hashimoto.y.gu@u.tsukuba.ac.jp
12∗‡
Ishigami Kana 1‡
Hassaninasab Azam 1‡
Kishi Katsuhiro 1
Kumano Takuto 12
Kobayashi Michihiko kobayashi.m.fe@u.tsukuba.ac.jp
123∗
1 Institute of Applied Biochemistry and Graduate School of Life and Environmental Sciences, The University of Tsukuba, , Tsukuba, Ibaraki, Japan
2 Microbiology Research Center for Sustainability (MiCS), The University of Tsukuba, Tsukuba, Ibaraki, Japan
3 Center for Quantum and Information LifeSciences, The University of Tsukuba, Tsukuba, Ibaraki, Japan
∗ For correspondence: Michihiko Kobayashi; Yoshiteru Hashimoto hashimoto.y.gu@u.tsukuba.ac.jpkobayashi.m.fe@u.tsukuba.ac.jp
‡ These authors contributed equally to this work.

08 8 2024
9 2024
08 8 2024
300 9 10764711 6 2024
24 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Curcumin is a plant-derived secondary metabolite exhibiting antitumor, neuroprotective, antidiabetic activities, and so on. We previously isolated Escherichia coli as an enterobacterium exhibiting curcumin-converting activity from human feces, and discovered an enzyme showing this activity (CurA) and named it NADPH-dependent curcumin/dihydrocurcumin reductase. From soil, here, we isolated a curcumin-degrading microorganism (No. 34) using the screening medium containing curcumin as the sole carbon source and identified as Rhodococcus sp. A curcumin-degrading enzyme designated as CurH was purified from this strain and characterized, and compared with CurA. CurH catalyzed hydrolytic cleavage of a carbon-carbon bond in the β-diketone moiety of curcumin and its analogs, yielding two products bearing a methyl ketone terminus and a carboxylic acid terminus, respectively. These findings demonstrated that a curcumin degradation reaction catalyzed by CurH in the soil environment was completely different from the one catalyzed by CurA in the human microbiome. Of all the curcumin analogs tested, suitable substrates for the enzyme were curcuminoids (i.e., curcumin and bisdemethoxycurcumin) and tetrahydrocurcuminoids. Thus, we named this enzyme curcuminoid hydrolase. The deduced amino acid sequence of curH exhibited similarity to those of members of acetyl-CoA C-acetyltransferase family. Considering results of oxygen isotope analyses and a series of site-directed mutagenesis experiments on our enzyme, we propose a possible catalytic mechanism of CurH, which is unique and distinct from those of enzymes degrading β-diketone moieties such as β-diketone hydrolases known so far.

Keywords

microbial screening
bacteria
secondary metabolites
curcuminoid
enzyme
enzyme purification
enzyme reaction
hydrolase
metabolism
bioconversion
Abbreviations

DHC dihydrocurcumin

THC tetrahydrocurcumin

Reviewed by members of the JBC Editorial Board. Edited by Chris Whitfield
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pmcWe have studied microbial metabolism of biologically active naturally occurring compounds [e.g., sesamin (1), harmaline (2) and C-glycosides (3, 4)] as well as man-made compounds (5, 6). The former compounds have been often invaluable for foods and clinical medicines. Curcumin is the most representative secondary metabolite component extracted from the rhizomes of a perennial plant Curcuma longa Linn (known as turmeric) and other Curcuma spp (7, 8). In Asian countries, ground turmeric powder containing 2 to 9% three major curcuminoids (i.e., curcumin, demethoxycurcumin, and bisdemethoxycurcumin) (9) has been used as a food-flavoring spice for centuries (10). Curcuminoids in turmeric are important ingredients in curry and gives curry powder a distinctive yellow color. Because of the intense color, turmeric has been widely used as an yellow pigment for foods, such as curry powders and mustard sauces, throughout the world. Curcumin, which is a plant-derived natural polyphenol, possesses a wide range of physiological and pharmacological properties (such as antioxidant, antitumor, neuroprotective, and antidiabetic activities) (7, 10, 11, 12, 13). Due to these effects, it is also available for a dietary supplement in several forms including capsules, tablets, energy drinks, ointments, and cosmetics. Orally delivered curcumin shows several biological effects such as antioxidant, antiinflammatory, and anticancer activities. In animals, curcumin in the diet is reduced to dihydrocurcumin (DHC), tetrahydrocurcumin (THC), hexahydrocurcumin, and octahydrocurcumin (14, 15, 16, 17, 18, 19). Curcumin and the reduced products are subjected to glucuronidation to yield curcumin glucuronide, DHC-glucuronide, THC-glucuronide, and curcumin sulfate (14, 15, 16, 17, 18, 19). Studies on purification and characterization of enzymes involved in the metabolism of curcumin, and cloning of their genes had not been reported for a long time. Therefore, we embarked on screening of microorganisms displaying curcumin-converting activity from human feces, and succeeded in the isolation of an Escherichia coli strain exhibiting the highest activity among enterobacteria (20). From the genome-sequenced E. coli str. K-12 substr. DH10B, which possessed higher curcumin-converting activity than the isolated strain, we purified an enzyme that acts on curcumin as a substrate and identified the structural gene. Based on its substrate specificity, reaction mode and NADPH-dependency, we named this enzyme NADPH-dependent curcumin/DHC reductase (CurA). Until now, findings regarding a purified enzyme involved in curcumin conversion have been limited to CurA.

To the best of our knowledge, on the other hand, metabolism of curcuminoids in nature has never been reported. Here, we for the first time describe the isolation and identification of a soil microorganism with curcumin degradation activity together with purification and characterization of a thus-far unknown enzyme (designated as CurH) showing this activity. We also demonstrated that the unique catalytic reaction catalyzed by this enzyme was quite different from the reduction reaction catalyzed by CurA.

Results

Isolation and identification of a curcumin-degrading microorganism

After about 3 months from the start of the screening, using the acclimatization culture method described under “Isolation of curcumin-degrading bacteria”, one microorganism, No. 34 strain, was obtained from soil at Mt. Tsukuba (Ibaraki, Japan).

The 16S rRNA gene sequence (Fig. S1) of the isolated microorganism was used as a query to search for homologous sequences in the Ribosomal Database Project (21). As a result, the nucleotide sequence of its 16S rRNA gene was found to show 99.39, 99.38, and 99.38% similarities to those of the closest type strains, Rhodococcus qingshengii djl-6T, Rhodococcus jialingiae djl-6-2T, and Nocardia coeliaca DSM44595T. Phylogenetic analysis revealed that the strain was closely related to the genus Rhodococcus (Fig. 1).Figure 1 Phylogenetic tree of 16S rRNA genes of Rhodococcus sp. strain No. 34 and the type strains of related species. The branching pattern was generated by the neighbor-joining method (45). The numbers at the nodes indicate the levels of bootstrap support based on a neighbor-joining analysis of 1000 resampled data sets. Rhodococcus sp. strain No. 34 is indicated in red. The scale bar represents 0.005 substitutions per a nucleotide position.

Morphologically, strain No. 34 was Gram stain-variable, nonmotile, and nonspore forming. Colonies on nutrient agar (Nissui Pharmaceutical Co., Ltd) were circular with whole margins and pale salmon-pink pigment, and reached 0.7 to 1.0  mm in diameter when cultivated at 30 °C for 48  h. The microscopic morphology varied from bacillary to coccoid depending on the conditions and phase of growth; 1.0 × 3.0 to 4.0 μm and 0.7 to 1.0 μm in diameter when cultivated at 30 °C for 24 h and 48 h, respectively. The strain grew at 15 and 30 °C, and exhibited slow growth at 37 °C, but its growth was inhibited at 45 °C. Its physiological characteristics were as follows: catalase, positive; nitrate reduction, negative; oxidase, negative; pyrazinamidase, negative; pyrrolidonyl arylamidase, negative; alkaline phosphatase, positive; β-glucuronidase, negative; β-galactosidase, negative; α-glucosidase, positive; esculin hydrolysis (β-glucosidase), positive; N-acetyl-β-glucosaminidase, negative; urease, positive; gelatin liquefaction, negative; and tyrosine hydrolysis, negative. Acid was not produced from d-glucose, d-ribose, d-xylose, d-mannitol, maltose, lactose, saccharose, or glycogen. Based on its physiological characteristics together with its 16S rRNA sequence, strain No. 34 was identified as Rhodococcus sp.

Purification of a curcumin-degrading enzyme (CurH)

When curcumin was used as a substrate for a cell-free extract prepared from the Rhodococcus cells grown in 2 × YT media containing 0.000005% (w/v) curcumin (see “Experimental procedures”), two possible product peaks were detected on HPLC with the consumption of curcumin. When the cell-free extract was not added in the reaction mixture, the substrate was not consumed. These findings indicated that strain No. 34 had an enzyme which probably degraded curcumin. The enzyme was purified through the eight purification steps described under “Experimental procedures” (Table 1). Although the purification was started from a large amount of cells (30 L), the yield of the purified enzyme (named CurH) was extremely low (0.0003%). The purified CurH gave only one band, corresponding to a molecular mass of 42 kDa, on SDS-PAGE (Fig. 2A).Table 1 Purification of the CurH

Step	Total protein	Total activity	Specific activity	Yield	
mg	units	units/mg	%	
Cell-free extract	15100	1140	0.0755	100	
DEAE-Sephacel	910	19.2	0.0211	1.69	
Hiprep Butyl	104	12.3	0.118	1.08	
Resource Q	17.4	1.70	0.0977	0.15	
TOYOPEARL PPG	2.68	1.31	0.489	0.12	
Mimetic Green	0.0881	0.502	5.70	0.04	
Type Q	0.0981	0.0433	0.441	0.004	
Resource Q	0.0138	0.00286	0.207	0.0003	

Figure 2 Discovery of curcumin-degrading enzyme, CurH.A, SDS-PAGE of purified CurH. Lane M, marker proteins. B–E, identification of reaction products of curcumin. Chromatograms of curcumin (B), the reaction mixture after incubation of curcumin with the purified recombinant CurH (C), authentic dehydrozingerone (D), and authentic ferulic acid (E). F–K, detection of reaction products of curcumin on LC/MS. Mass spectra of dehydrozingerone (F) and ferulic acid (G) in the reaction mixture, dehydrozingerone (F) and ferulic acid (I) in the reaction mixture containing H218O, dehydrozingerone (J), and ferulic acid (K) in the reaction mixture containing 18O2.

Cloning of the gene encoding CurH

To identify the gene encoding CurH, the N terminal partial amino acid sequences of the purified enzyme were determined to be PEAVIVSAVR. Using the sequence information, a local BLAST search was run on the draft genome database for strain No. 34 constructed in-house. We identified an ORF (designated as curH) of 1218 nucleotides (Fig. S2). The curH gene was found to code for a protein of 405 amino acids, ten of which corresponded to the above N-terminal sequence. The calculated molecular mass (42,244 Da) was consistent with that of the purified enzyme determined on SDS-PAGE.

Heterologous expression and purification of the recombinant CurH

In order to overproduce CurH, an expression plasmid was constructed and introduced into E. coli BL21-CodonPlus(DE3)-RIL, as described under “Experimental procedures”. By SDS-PAGE, we analyzed cell-free extracts prepared from the E. coli transformant and detected a remarkable protein band at the 42 kDa position (Fig. S3A). On the other hand, the remarkable band was not detected for cell-free extracts prepared from E. coli BL21-CodonPlus(DE3)-RIL carrying pET-24a(+). Cell-free extracts prepared from the transformant harboring pET-curH, which was cultured at 28 °C, exhibited significant specific activity (i.e., 0.475 units/mg) (Table S1). Therefore, overproduction of CurH in the active form was attained in the recombinant cells, which made it easy to purify the enzyme to homogeneity through ammonium sulfate fractionation and two-step column chromatography procedures (Table S1). The purified recombinant enzyme gave only one band on SDS-PAGE (Fig. S3B), and the apparent molecular mass of the purified enzyme was the same as that of the Rhodococcus sp. No. 34 enzyme.

Identification of the curcumin metabolites, elucidation of the CurH reaction by oxygen isotope analyses and its stoichiometry

We incubated the purified recombinant CurH with curcumin as a substrate. On HPLC, two peaks predicted to be reaction products were detected with the consumption of curcumin (Fig. 2, B and C). Next, we measured mass spectra of the probable reaction products by LC/MS, as described under “Experimental procedures”. The major mass peaks for the corresponding HPLC peaks were found at m/z 191 [M-H]- and 193 [M-H]- in the negative ion mode (Fig. 2, F and G), demonstrating that the molecular masses of the probable products were 192 and 194, respectively. Their retention times and UV-visible spectra were found to agree with those of authentic dehydrozingerone and ferulic acid, respectively (Fig. 2, C–E and Fig. S4, A–D). Based on these findings, the reaction products were identified as dehydrozingerone and ferulic acid (Fig. 3).Figure 3 The reaction formulae for degradation of curcumin and its derivatives by CurH. As for asymmetric compounds, the structures flipped horizontally are shown in parentheses.

Using oxygen isotopes, we next investigated what kind of an oxygen atom was incorporated into the reaction products. When curcumin and the purified recombinant CurH were incubated in the reaction mixture containing 89% of H218O/11% of H216O, dehydrozingerone and ferulic acid were detected as m/z 191 [M-H]- and 195 [M-H]- products, respectively (Fig. 2, H and I). These findings indicated that these products were 18O-unlabeled dehydrozingerone and single 18O-labeled ferulic acid. In the reaction mixture under 18O2, on the other hand, the mass peaks of dehydrozingerone and ferulic acid were observed to be m/z 191 [M-H]- and 193 [M-H]-, respectively (Fig. 2, J and K), indicating that an 18O atom was not incorporated into either product. These findings demonstrated that one oxygen atom derived from water was incorporated into ferulic acid during the CurH reaction.

The stoichiometry of substrate consumption and product formation during the degradation of curcumin was next examined in the standard assay A mixture. The amounts of curcumin consumed, and dehydrozingerone and ferulic acid formed in the reaction mixture were determined, respectively. At several time points, the molar amount of the dehydrozingerone increase was consistent with that of the curcumin decrease (Fig. S5). On the other hand, that of ferulic acid formed was about 45% of that of curcumin consumed, the reason for which remains unknown. These findings indicated that dehydrozingerone was formed with the consumption of curcumin in a 1:1 stoichiometry. In order to calculate the CurH activity accurately, the amount of dehydrozingerone formed was measured in the following experiments, unless otherwise stated.

Substrate specificity

Instead of curcumin, we examined whether CurH could utilize each of the compounds listed in Fig. S6 as a substrate, and the compounds degraded by CurH and the corresponding products are summarized in Figure 3. In the case of bisdemethoxycurcumin, 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione, demethoxycurcumin, DHC, THC, tetrahydrobisdemethoxycurcumin, and tetrahydrodemethoxycurcumin, peaks predicted to be reaction products were observed in association with the consumption of each substrate on HPLC (Figs. S7, A and B, S8, A and B, S9, A and B, S10, A and B, S11, A and B, S12, A and B, and S13, A and B). These findings suggested that the corresponding products would be formed during the CurH reaction.

When bisdemethoxycurcumin was used as a substrate, two peaks of probable reaction products appeared. The retention times of these compounds were found to be consistent with those of authentic 4-hydroxybenzylideneacetone [4-(4-hydroxyphenyl)-3-buten-2-one] and p-hydroxycinnamic acid (p-coumaric acid), respectively (Fig. S7, B–D). LC/MS analyses revealed that the corresponding compounds exhibited m/z 161 [M-H]- and 163 [M-H]- values (Fig. S7, E and F), which were in a good agreement with the molecular masses of 4-hydroxybenzylideneacetone and p-hydroxycinnamic acid, respectively. These findings demonstrated that 4-hydroxybenzylideneacetone and p-hydroxycinnamic acid were generated as the reaction products from bisdemethoxycurcumin by CurH (Fig. 3).

The retention times of two probable compounds produced in the reaction mixture containing 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione as a substrate were identical to those of authentic 3,4-dimethoxybenzylideneacetone and 3,4-dimethoxycinnamic acid, respectively (Fig. S8, B–D). We observed good agreement between peaks of two compounds on HPLC chromatograms and peaks with m/z 207 [M-H]- and 207 [M + H]+ values, which were the molecular masses of 3,4-dimethoxybenzylideneacetone and 3,4-dimethoxycinnamic acid, respectively, on mass chromatograms (Fig. S8, B, E, and F). Our findings indicated that 3,4-dimethoxybenzylideneacetone and 3,4-dimethoxycinnamic acid were reaction products from 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione (Fig. 3).

Interestingly, four peaks predicted to be reaction products were obtained on HPLC in the case of demethoxycurcumin as a substrate. The retention times of the four compounds were proved to be identical to those of authentic dehydrozingerone, p-hydroxycinnamic acid (p-coumaric acid), 4-hydroxybenzylideneacetone and ferulic acid, respectively (Fig. S9, B–F). Among them, two probable products exhibited m/z 163 [M-H]- and 161 [M-H]- values, which were in complete agreement with the molecular masses of p-hydroxycinnamic acid (p-coumaric acid) and 4-hydroxybenzylideneacetone, respectively (Fig. S9, G and H). The peaks of the other two probable products on the HPLC chromatograms were close to mass chromatogram peaks with m/z 193 [M-H]- and 193 [M + H]+ values, being derived from ferulic acid and dehydrozingerone, respectively (Fig. S9, B, I, and J). These findings indicated that CurH acted on asymmetric demethoxycurcumin, and produced four compounds [i.e., dehydrozingerone, p-hydroxycinnamic acid (p-coumaric acid), 4-hydroxybenzylideneacetone, and ferulic acid] (Fig. 3).

As for another asymmetric DHC used as a substrate, the retention times of four reaction products were also found to be consistent with those of authentic dehydrozingerone, dihydroferulic acid [3-(4-hydroxy-3-methoxyphenyl)-propionic acid], zingerone and ferulic acid (Fig. S10, B–F). The retention times of the two peaks derived from two compounds among them on the HPLC chromatograms were noted to be same as those of mass chromatogram peaks with m/z 193 [M-H]- and 193 [M + H]+ values, which were in a good agreement with the molecular masses of ferulic acid and dehydrozingerone, respectively (Fig. S10, B, G, and H). A series of HPLC analyses revealed that dehydrozingerone, dihydroferulic acid, zingerone, and ferulic acid were reaction products from DHC (Fig. 3).

The retention times of two compounds formed in the reaction mixture containing THC as a substrate were identical to those of authentic zingerone and dihydroferulic acid, respectively (Fig. S11, B–D). The two mass peaks derived from the compounds were detected at m/z 195 [M + H]+ in the positive ion mode and 195 [M-H]- in the negative ion mode, respectively (Fig. S11, E and F). These findings demonstrated that zingerone and dihydroferulic acid were the consequent products from THC (Fig. 3).

When tetrahydrobisdemethoxycurcumin was used as a substrate, the retention times of two possible reaction products were consistent with those of authentic 4-(4-hydroxyphenyl)-butan-2-one and 3-(4-hydroxyphenyl) propionic acid, respectively (Fig. S12, B–D), indicating that the two compounds were yielded as reaction products from tetrahydrobisdemethoxycurcumin by CurH (Fig. 3).

In the case of the other asymmetric compound, tetrahydrodemethoxycurcumin, used as a substrate, the retention times of four possible reaction products were in accord with those of authentic zingerone, 3-(4-hydroxyphenyl) propionic acid, 4-(4-hydroxyphenyl)-butan-2-one and dihydroferulic acid, respectively (Fig. S13, B–F). LC/MS analyses revealed that the retention time of one of the peaks of reaction products on HPLC chromatograms was the same as that of the peak at m/z 195 [M + H]+ for zingerone on mass chromatograms (Fig. S13, B and G). These findings suggested that CurH acted on tetrahydrodemethoxycurcumin, forming zingerone, 3-(4-hydroxyphenyl) propionic acid, 4-(4-hydroxyphenyl)-butan-2-one and dihydroferulic acid as reaction products (Fig. 3).

Meanwhile, the enzyme did not act on hexahydrocurcumin, octahydrocurcumin, 1,6-diphenylhexane-1,3,4,6-tetrone, 1,6-diphenyl-1,6-hexanedione (1,4-dibenzoylbutane), 1,5-diphenyl-1,5-pentanedione (1,3-dibenzoylpropane), 1,4-diphenyl-1,4-butanedione (1,2-dibenzoylethane), 1,3-diphenyl-1,3-propanedione (dibenzoylmethane), 1,3-bis(4-methoxyphenyl)-1,3-propanedione, 1-(2-hydroxyphenyl)-1,3-butanedione [2-(acetoacetyl)phenol], or 1,2-diphenyl-1,2-ethanedione (benzil) listed in Fig. S6.

The apparent Km and Vmax values for symmetric compounds including curcumin among the substrates which CurH accepted were determined (Fig. 4, A–D and Table 2). These values for 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione and asymmetric substrates could not be determined due to the poor substrate solubility and/or very low activity under the assay conditions.Figure 4 Determination of kinetic parameters for CurH. Michaelis-Menten kinetics of CurH versus different substrates. Concentration dependence of CurH activity toward curcumin (A), bisdemethoxycurcumin (B), tetrahydrocurcumin (C), and tetrahydrobisdemethoxycurcumin (D).

Table 2 Substrate specificity

Substrate	Km	Vmax	kcat/Km	
mM	units/mg	s−1 mM−1	
Curcumin	0.071 ± 0.011	2.20 ± 0.13	0.0221	
Bisdemethoxycurcumin	0.14 ± 0.03	4.38 ± 0.50	0.0220	
Tetrahydrocurcumin (THC)	0.56 ± 0.21	1.05 ± 0.15	0.0132	
Tetrahydrobisdemethoxycurcumin	0.48 ± 0.16	0.699 ± 0.085	0.0102	
The apparent Km and Vmax values for 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione and asymmetric substrates among the substrates on which CurH acted (Fig. 3) could not be determined due to the poor substrate solubility and/or very low activity. The substrates which CurH did not accept are given under “Substrate specificity” in “Results”.

Molecular mass determination, metal analysis, and spectroscopic analysis of CurH

To determine the molecular mass of CurH, we subjected the purified recombinant enzyme to gel filtration on a Superose 6 HR10/30 column. The molecular mass of the native enzyme was found to be 79.2 kDa (Fig. S14), indicating that the enzyme consists of two identical subunits.

Qualitative analyses of the following metals in the purified recombinant enzyme solution were performed with an inductively coupled radiofrequency plasma spectrophotometer: Li, Be, B, Na, Mg, Al, Si, P, K, Ca, Sc, Ti, V, Cr, Mn, Fe, Co, Ni, Cu, Zn, Ga, Ge, As, Se, Sr, Rb, Sr, Y, Zr, Nb, Mo, Ru, Rh, Pd, Ag, Cd, In, Sn, Sb, Te, Ba, La, Ce, Pr, Nd, Sm, Eu, Gd, Tb, Dy, Ho, Rr, Tm, Yb, Lu, Hf, Ta, W, Re, Os, Ir, Pt, Au, Hg, Tl, Pb, and Bi. As a result, none of the 67 metals was detected within the limits of the assay.

The absorption spectrum of the purified recombinant enzyme in 10 mM KPB (pH 6.0) showed maximum absorbance at 280 nm, but no other peak absorption or shoulder was observed (Fig. S15). These findings suggested that no cofactor would bind to CurH.

Effects of temperature and pH on the activity and stability of CurH

The effects of pH and temperature on the enzyme activity were examined. CurH exhibited maximum activity around pH 6.0, as shown in Figure 5A. The optimal reaction temperature appeared to be 40 °C when the reaction was carried out for 40 min (Fig. 5B).Figure 5 Effects of temperature and pH on the activity and stability of CurH.A, reactions were carried out in Britton–Robinson buffer (pH 3.0–12.0). B, reactions were carried out at various temperatures. C, CurH was incubated at various pH values at 4 °C for 30 min in Britton–Robinson buffer at a concentration of 20 mM, an aliquot of each solution was taken, and then the activity of CurH was measured under the standard assay conditions. D, CurH was preincubated at various temperatures for 30 min in 10 mM KPB (pH 6.0), and then the residual activity was assayed.

The stability of CurH was examined at various pHs. After the enzyme had been incubated at 0 °C for 15 min in Britton-Robinson buffer (pH 2.0–12.0; 0.1 M), an aliquot of the enzyme solution was taken, and then the enzyme activity was assayed under the standard conditions. The enzyme was most stable in the pH range of 4.0 to 11.0, 80% of its initial activity being retained even at pH 3.5 (Fig. 5C).

In contrast to determination of the temperature optimum of the reaction, which was performed for 30 min at different temperatures, protein stability was investigated after incubation for 15 min at various temperatures in 10 mM KPB (pH 6.0). An aliquot of the enzyme solution was taken, and then the enzyme activity was assayed under the standard conditions. CurH was stable after heat treatment of 45 °C for 30 min, but unstable after that of 50 °C (Fig. 5D).

Inhibitors of CurH enzyme activity

Various compounds were investigated as to their inhibitory effects on the enzyme activity (Table 3). The enzyme was very sensitive to CdCl2, FeCl3, HgCl2, and CuCl2, the inhibition being 100%. On the contrary, the activity increased by about three times in the presence of Na2MoO4. CurH was completely inhibited by one thiol-specific reagent, N-ethylmaleimide, while iodoacetate and 5,5′-dithio-bis-2-nitrobenzoate caused only partial inhibition. The addition of carbonyl-specific reagents and a serine-modifying reagent (diisopropyl fluorophosphates) did not cause inhibition of the enzymatic activity. The enzyme was not sensitive to chelating agents such as α,α′-dipyridyl, o-phenanthroline, EDTA, and diethyldithiocarbamate. Whereas, the addition of oxidizing reagents such as H2O2 and ammonium persulfate showed partial inhibitory effects on the enzymatic activity (35% and 10%, respectively), and that of some reducing reagents, i.e., 2-mercaptoethanol and DTT, showed enhancing effects (282% and 273%, respectively). Hydroxylamine and NaN3, both of which were regarded as major inhibitors for hemoproteins, enhanced the activity of CurH.Table 3 Effects of various compounds on the activity of CurH

Inhibitor	Relative activity	
%	
None	100	
Na2MoO4	287	
LiCl, NaCl, MgCl2, CaCl2, BaCl2, MnCl2, ZnCl2, CoCl2, RbCl, CsCl, SrCl2, PbCl2, NiCl2	91.6–109	
AlCl3	62.5	
FeSO4	24.9	
CdCl2, FeCl3, HgCl2, CuCl2	0	
Iodoacetate	88.4	
5, 5′-Dithio-bis-2-nitrobenzoate	27.3	
N-Ethylmaleimide	0	
Hydroxylamine	218	
Phenylhydrazine	109	
Semicarbazide	86.2	
Aminoguanidine	101	
α, α′-Dipyridyl	133	
o-Phenanthroline	137	
EDTA	88.8	
Diethyldithiocarbamate	111	
NaN3	202	
KCN	136	
Dithiothreitol	273	
2-Mercaptoethanol	282	
Na2S2O4	70.6	
H2O2	65.2	
Ammonium persulfate	90.0	
Diisopropyl fluorophosphate	114	
Each compound was added to the standard reaction mixture without the substrate, and then assaying of the enzyme (standard assay B) was performed after adding the substrate. The final concentrations of the tested compounds were 1 mM, unless otherwise stated.

Homology analysis and mutational analysis of CurH

The deduced amino acid sequences of curH showed similarity with those of acetyl-CoA C-acetyltransferases from Clostridium acetobutylicum ATCC (American Type Culture Collection) 824 [44% identity; accession number 4WYR_A (chain A of homodimer)], E. coli [44% identity; accession number 4WYS_A (chain A of homotetramer)] (22), Zoogloea ramigera [37% identity; accession number 1DLU_A (chain A of homotetramer)] (23), and Pseudomonas fragi 3-ketoacyl-CoA thiolase [39% identity; accession number 1WDK_C], which formed a β-oxidation multienzyme complex with a fatty oxidation complex α subunit consisting of 2-enoyl-CoA hydratase and 3-hydroxyacyl CoA dehydrogenase (24) (Fig. 6). In acetyl-CoA C-acetyltransferases of Z. ramigera, cysteine and histidine residues (i.e., Cys89, His348, and Cys378) have been reported for the first time to participate in the active site (23, 25). In other enzymes, the corresponding three amino acids have been indicated as catalytic residues based on their similar sequence location in the Z. ramigera enzyme. Cys90, His360, and Cys390, corresponding to the above three active residues, were conserved in CurH, respectively (Fig. 6).Figure 6 Alignment of amino acid sequences of curcuminoid hydrolase (CurH) and the acetyl-CoA C-acetyltransferase family.Arrows indicate the amino acid residues in red letters (i.e., C90, H360, and C390 in CurH) that were mutated in this work. Regions in green and red rectangles are described under “Discussion”. CurH, curcuminoid hydrolase from Rhodococcus sp. strain No. 34; 4WYR_A, acetyl-CoA C-acetyltransferase from Clostridium acetobutylicum ATCC824; 4WYS_A, acetyl-CoA C-acetyltransferases from Escherichia coli; 1DLU_A, acetyl-CoA C-acetyltransferase from Zoogloea ramigera; and 1WDK_C, 3-ketoacyl-CoA thiolase from Pseudomonas fragi. Identical residues are denoted by asterisks. ATCC, American Type Culture Collection.

To understand the structural basis of the unprecedented catalysis by CurH, we predicted the structure using the AlphaFold2 (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb) (26). The overall dimeric structure of CurH obtained with AlphaFold2 was almost the same as those of 4WYR, 4WYS, 1DLU, and 1WDK, respectively (Fig. 7). Moreover, the positions of the Cys90, His360, and Cys390 residues in the CurH structure were found to be closely identical to those of the three active residues in 1DLU_A (Fig. 8A). Thus, we constructed three CurH mutant enzymes (i.e., C90A, H360A, and C390A) by site-directed mutagenesis. According to the same procedure as used for the recombinant WT enzyme, each of the mutant enzymes was expressed in E. coli, purified to homogeneity (Fig. 9A), and then characterized. We compared each purified mutant enzyme with the WT CurH by CD spectrum analysis (Fig. 9B). The results indicated that essentially no major change in the overall conformation of the enzyme protein was induced by the mutations. The three mutant enzymes (C90A, H360A, and C390A) did not exhibit detectable curcumin-degrading activity at all.Figure 7 Structure model of CurH.A, the homodimeric structure of CurH predicted by AlphaFold2. Chains A and B are colored cyan and pale cyan, respectively. B, overlay of the structure of CurH and the crystal structure of acetyl-CoA C-acetyltransferase from Clostridium acetobutylicum (4WYR; homodimer). Chains A and B of 4WYR are colored pink and magenta, respectively. C, overlay of the structure of CurH and the crystal structure of acetyl-CoA C-acetyltransferase from Escherichia coli (4WYS; homotetramer). Chains A and C of 4WYS are colored pink, and chains B and D of 4WYS are colored magenta. D, overlay of the structure of CurH and crystal structure of the acetyl-CoA C-acetyltransferase from Zoogloea ramigera (1DLU; homotetramer). Chains A and C of 1DLU are colored pink, and chains B and D of 1DLU are colored magenta. E, overlay of the structure of CurH and crystal structure of the fatty acid β-oxidation multienzyme complex from Pseudomonas fragi (1WDK; α2β2 heterotetramer). Chains C and D (for 3-ketoacyl-CoA thiolase) of 1WDK are colored pink and magenta, respectively. Chains A and B (for fatty oxidation complex α subunit) of 1WDK are colored orange and yellow, respectively.

Figure 8 Structure model of CurH. Overlay of the AlphaFold2-predicted structure for CurH (monomer) in cyan and the crystal structure of acetyl-CoA C-acetyltransferase from Zoogloea ramigera (1DLU_A; monomer) in pink. A, the regions marked out are the possible active sites. Active amino acid residues in CurH and the acetyl-CoA C-acetyltransferase are shown at the top and bottom, respectively. The distances in Ångstrom units between atoms in CurH and the acetyl-CoA C-acetyltransferase are indicated in cyan and pink, respectively. B, residues 120 to 171 in CurH and residues 119 to 156 in 1DLU_A are colored green and red, respectively. The possible active sites are indicated in the pale orange rectangle region.

Figure 9 Characterization of purified WT CurH and its mutants.A, SDS-PAGE of the WT CurH and its mutants. Protein bands were detected by staining with Coomassie brilliant blue. Lane M, marker proteins; lane 1, WT CurH; lane 2, C90A; lane 3, H360A; and lane 4, C390A. (B) CD spectra for the WT CurH and its mutants. The WT CurH (black), and C90A (red), H360A (green), and C390A (blue) mutants were examined.

Discussion

Although the impressive properties (such as antioxidant, antitumor, antiinflammatory, neuroprotective, and antidiabetic activities) of curcumin have been found via both laboratory and clinical research, no medical use for curcumin has been confirmed. The reasons for this are that curcumin is unstable and not bioavailable. Curcuminoid biosynthesis involving diketide-CoA synthases and curcumin synthases 1 to 3 (CURS1, CURS2, and CURS3) in C. longa has already been disclosed (27, 28, 29). As for curcumin metabolism, two phases in several mammalian tissues have been reported so far (15, 16, 17, 18, 19). Phase I metabolism comprises the reduction of the four double bonds of the heptadiene-3,5-dione structure: curcumin → DHC → THC → hexahydrocurcumin → octahydrocurcumin (14, 15, 16, 17, 18, 19). During phase II, curcumin and its reduced metabolites are conjugated with a monoglucuronide, a monosulfate and a mixed sulfate/glucuronide: conjugated curcumin → conjugated DHC → conjugated THC → conjugated hexahydrocurcumin → conjugated octahydrocurcumin (14, 15, 16, 17, 18, 19). However, there had been neither purification of the enzymes involved in the metabolic pathway for curcumin nor identification of their genes in living organisms. Thus, the detailed mechanism of curcumin metabolism has not been identified at all.

We were interested in determining whether intestinal microorganisms could metabolize curcumin, because curcumin has been taken orally as a food agent since ancient times. In the previous study, we isolated microorganisms exhibiting curcumin-converting activity from human feces. For the first time, we succeeded in purifying the enzyme (CurA) that uses curcumin as a substrate (20). The enzyme has a narrow substrate spectrum, preferentially acting on curcumin. Because CurA was demonstrated to catalyze the two-step reduction of curcumin to THC, we named this enzyme “NADPH-dependent curcumin/DHC reductase”. To the best of our knowledge, CurA has been reported as the sole purified and characterized enzyme responsible for curcumin transformation. Other enzymes that are thus far unknown to be involved in curcumin metabolism remain to be identified. While curcumin and curcuminoid have been reported to be produced 1.87% and 3.2%, respectively, in rhizomes of fresh C. longa (30), these compounds have never been deposited in the soil; the existence of unknown enzymes for curcumin metabolism could be expected in the natural rhizosphere. However, there has been no report on the biodegradation of curcumin in the soil at either the protein or gene level. Therefore, we addressed the question of whether soil microorganisms that can metabolize curcumin exist or not.

To isolate curcumin-converting microorganisms from human feces in the previous study (20), we used a synthetic medium comprising 0.05% (w/v) KH2PO4, 0.05% K2HPO4, 0.1% NaCl, 0.05% MgSO4·7H2O, 0.2% yeast extract, 0.05% curcumin, and 5% Tween 20. Although yeast extract was added as a nitrogen source to this medium, it could be used as a carbon source, too, by intestinal E. coli. Tween, which was added to enhance curcumin solubility in this medium, is also utilized as a carbon source by some microorganisms. For example, members of the genus Malassezia have the ability to assimilate various Tweens (31). In an attempt to obtain curcumin-degrading bacteria with another metabolic pathway, here, we used another screening medium containing curcumin as the sole carbon source, as described under “Experimental procedures”.

In this study, we purified the second enzyme that uses curcumin as a substrate. Based on the molecular masses of probable products on LC/MS and comparison of HPLC profiles (retention times and UV-visible absorption spectra) of the probable products and authentic compounds, we identified dehydrozingerone and ferulic acid as the two reaction products. During these investigations, we discovered a unique curcumin-metabolic pathway (curcumin → dehydrozingerone + ferulic acid) involving the thus-far unknown curcumin-degrading enzyme (CurH) (Fig. 3). The molecular formulae of dehydrozingerone and ferulic acid yielded from curcumin (C21H20O6) were C11H12O3 and C10H10O4, respectively. These findings suggested that two hydrogen atoms and one oxygen atom were incorporated into the two products through the enzymatic reaction. In addition, our oxygen isotope analysis revealed that one oxygen atom derived from the water molecule was incorporated into ferulic acid (Fig. 2I). These findings demonstrated that the products were generated via hydrolytic cleavage of a carbon-carbon bond in the β-diketone moiety. When CurH acted on an asymmetric substrate, it gave four reaction products. In other words, CurH has been found to hydrolyze curcumin or curcumin analogs through this thus-far unknown biotransformation pathway. Whereas, we previously demonstrated that curcumin was metabolized through a reduction reaction by CurA in the human microbiome (20), we here clarified that curcumin would be degraded via a hydrolytic reaction in natural environments. In addition, CurH was proved to act on 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione, demethoxycurcumin (known as curcumin II among natural curcuminoids), bisdemethoxycurcumin (known as curcumin III among natural curcuminoids) besides curcumin (known as curcumin I among natural curcuminoids), and the reductive metabolites (DHC, THC, tetrahydrodemethoxycurcumin, and tetrahydrobisdemethoxycurcumin), all of which possess a β-diketone moiety (Fig. 3). Hexahydrocurcumin and octahydrocurcumin, which structurally resemble curcumin but possess no β-diketone moiety, were inert as substrates. CurH did not act on 1,4-diphenyl-1,4-butanedione (1,2-dibenzoylethane), 1,5-diphenyl-1,5-pentanedione (1,3-dibenzoylpropane), or 1,6-diphenyl-1,6-hexanedione (1,3-dibenzoylbutane) (Fig. S6), which possess γ-, δ-, ε-diketone moieties, respectively. As a result of kinetic analyses (Table 2), the affinities for curcuminoids with a 1,6-heptadiene-3,5-dione backbone (e.g., curcumin and bisdemethoxycurcumin) were found to be lower than those for tetrahydrocurcuminoids with a heptane-3,5-dione backbone (e.g., THC and tetrahydrobisdemethoxycurcumin). According to the catalytic efficiencies (kcat/Km), potential substrates for CurH could be natural curcuminoids. Unlike CurA, CurH was proved to be independent of any organic cofactors (e.g., NADPH, flavin, and heme) in the enzymatic reaction. Due to the catalytic reaction and the substrate specificity, we named the purified enzyme (CurH) a “curcuminoid hydrolase”. Notably, the Km value (0.071 mM) for curcumin was low enough for CurH to serve as a substrate under biological conditions, although the Vmax value was low. When Rhodococcus sp. strain No. 34 was grown in the screening medium containing curcumin as the sole carbon source, the curcuminoid hydrolase could play a physiological role for curcumin metabolism. Further studies are required to understand the biological role of CurH.

Interestingly, homology searches of a protein database demonstrated that CurH exhibited considerable similarity to members of the acetyl-CoA C-acetyltransferases/thiolase family such as acetyl-CoA C-acetyltransferase or 3-ketoacyl-CoA thiolase (Fig. 6). The latter enzymes have been reported to contain crucial two cysteine residues and one histidine residue in their active sites (22, 23, 24, 25). The Cys90, His360, and Cys390 residues, which correspond to these three essential residues, were found to be conserved in CurH (Fig. 6). Structure prediction of CurH with AlphaFold2 suggested that the Cys90, His360, and Cys390 residues in CurH were located close to one another. Furthermore, superposition of the predicted structure for CurH with the crystal structure for the acetyl-CoA C-acetyltransferase from Z. ramigera (1DLU_A) revealed that the positions of the three residues in CurH strongly coincided with those of the active residues (i.e.., Cys89, His348, and Cys378) in 1DLU_A (Fig. 8A). These findings suggested that the three residues in CurH serve as active amino acids. Mutational analysis of CurH clearly demonstrated that the Cys90, His360, and Cys390 residues were all crucial for curcuminoid hydrolase activity. Based on these findings and the oxygen isotope analyses, we here propose a possible reaction mechanism for CurH (Fig. 10). The substrate (curcumin) enters the active site of CurH [step (i)]. The catalytic cycle is initiated by His360-dependent deprotonation of the catalytic Cys90 to provide the anionic thiolate, followed by the attack by the activated anionic thiolate on the carbonyl C3 atom of curcumin. Concomitantly, the proton of the thiol group in Cys390 is transferred to the C4 atom of curcumin [step (ii)], leading to the formation of a covalent S-feruloyl-enzyme intermediate and release of dehydrozingerone (as a product) [step (iii)]. Next, the nucleophilic catalysis of Cys390 to a water molecule [step (iv)] results in the formation of a hydroxide anion, which attacks the acyl-enzyme intermediate [step (iv)]. The protonated His360 should donate a proton to the catalytic Cys90 in a tetrahedral intermediate [step (v)]. Finally, ferulic acid (as the other product) is liberated, and the initial state of the enzyme is regenerated for another catalytic cycle [step (vi)]. Involvement of the cysteine residue(s) located at the catalytic site in the hydrolytic degradation mechanism for curcuminoid was consistent with the experimental observation of inhibition of the enzyme activity caused by the addition of SH reagents to CurH (Table 3).Figure 10 A possible reaction mechanism for CurH. Steps (i)-(vi) are described under “Discussion”.

As mentioned above, CurH shows sequence similarity to acetyl-CoA C-acetyltransferases/thiolases, such as acetyl-CoA acetyltransferase and 3-ketoacyl-CoA thiolase (Fig. 6). In general, acetyl-CoA C-acetyltransferases are ubiquitous enzymes that have key roles in many vital biochemical pathways (25, 32). These enzymes can be divided into two broad categories: biosynthetic enzymes (EC 2.3.1.9) and degradative enzymes (EC 2.3.1.16). The former, also known as acetoacetyl-CoA thiolases, catalyze the biological Claisen condensation of two acetyl-CoA molecules to yield acetoacetyl-CoA; these enzymes are involved in the carbon-carbon bond-forming condensation step at the start of isoprenoid biogenetic pathways such as steroidogenesis. The latter functions in the β-oxidation pathway for fatty acids with four other enzymes (Fig. S16A). Fatty acid is converted to acyl-CoA by acyl-CoA synthetase (fatty acid-CoA ligase) in the presence of CoA and ATP. The degradation of acyl-CoA compounds proceeds in a cyclic manner, resulting in the release of one unit of acetyl-CoA in each cycle. Acyl-CoA dehydrogenase catalyzes the oxidation of acyl-CoA to enoyl-CoA. Enoyl-CoA hydratase performs the hydration of enoyl-CoA to 3-hydroxyacyl-CoA, and 3-hydroxyacyl-CoA dehydrogenase oxidizes this intermediate to 3-ketoacyl-CoA. Acetyl-CoA C-acetyltransferase, also known as 3-ketoacyl-CoA thiolase, catalyzes the last step in the cycle, where acetyl-CoA and a shortened acyl-CoA (reduced by two carbon atoms) are yielded via thiolytic degradation of 3-ketoacyl-CoA. In our experiments, the draft genome of strain No. 34 was found to contain at least 33 genes encoding acetyl-CoA C-acetyltransferase homologs, and 21 loci containing one or a few acetyl-CoA C-acetyltransferase homolog genes. In loci 2, 4, 14, 15 and 18 (Fig. S16B), the gene encoding an acetyl-CoA C-acetyltransferase homolog and the genes encoding four other enzymes (acyl-CoA synthetase, acyl-CoA dehydrogenase, enoyl-CoA hydratase, and 3-hydroxyacyl-CoA dehydrogenase homologs) are located together, indicating that the gene products for the acetyl-CoA C-acetyltransferase homolog in these loci are mainly responsible for the release of acetyl-CoA from 3-ketoacyl-CoA in the β-oxidation pathway for fatty acids in strain No. 34. In locus 1, in which the curH gene is located, on the other hand, genes coding for acyl-CoA synthetase, acyl-CoA dehydrogenase, and 3-hydroxyacyl-CoA dehydrogenase homologs are not present, and only the enoyl-CoA hydratase homolog gene is located in the opposite direction of the curH gene (Fig. S16B). Moreover, the enoyl-CoA hydratase homolog gene and the curH gene must be divergently transcribed, and the two putative transcriptional units could be distinctly different from each other. Together with the very low Km value (0.071 mM) of CurH for curcumin described above, the gene organization in locus 1 indicates that CurH plays a key role in hydrolytic degradation of curcumin or its analogs physiologically rather than thiolytic degradation of 3-ketoacyl-CoA for fatty acid β-oxidation, although CurH shows sequence similarity to acetyl-CoA C-acetyltransferase. In the overlayed structures of the acetyl-CoA C-acetyltransferase from Z. ramigera (1DLU_A), the two structures including each active site were entirely identical to each other (Fig. 8A). Intriguingly, the region of residues 120 to 171 in CurH exhibited little amino acid sequence similarity to that of residues 119 to 156 in the Z. ramigera enzyme (Fig. 6). The structure consisting of residues 120 to 171 was located away from the active amino acids in CurH, while that consisting of residues 119 to 156 was located closer to the active amino acids in 1DLU_A (Fig. 8B). The wide cavity around the active site of CurH would make access possible for bulky substrates with two aryl side chains like curcumin. Our findings provide thus-far unknown insights into the function of members of the acetyl-CoA C-acetyltransferases/thiolase family.

CurH belongs to the β-diketone hydrolases, which are classified as EC 3.7.1- {“Hydrolases [3] acting on C-C bonds [7] in ketonic substances [1]”} by the “Enzyme Commission on Nomenclature” (https://iubmb.qmul.ac.uk/enzyme/EC3/7/1/), because our experiments demonstrated that CurH cleaved the carbon-carbon bond between two carbonyl groups to yield two products: one bearing a methyl ketone terminus and the other bearing a carboxylic acid terminus (Fig. 3). Among the 23 enzymes listed in the EC 3.7.1- group, ten kinds of β-diketone hydrolases are registered, and four types of possible catalytic mechanisms for β-diketone hydrolases are proposed (33). Interestingly, however, CurH exhibits no amino acid sequence similarity to β-diketone hydrolases so far reported. The catalytic mechanism of CurH that we here uncovered is also significantly different from those of the four types of β-diketone hydrolases. These findings indicate that CurH is structurally unrelated to other β-diketone hydrolases, while the chemical equations for all the enzymes are the same as one another. Thus, we discovered an unexpected member of the β-diketone hydrolases named CurH.

Discovery of so-far unknown biocatalysts obtained from screening can often provide many breakthroughs. Enzymes and ribozymes, which are composed of proteins and RNAs, respectively, are known as biocatalysts in living organisms in general. During the investigations on the screening of useful enzymes, we discovered that living organisms produced certain natural low-molecular mass organic compounds (e.g., actinorhodin, 2,6-dimethoxyquinone, antiarol, and juglone), and used them as organocatalysts (34). In chemical manufacturing, the use of various enzymes has been widely accepted owing to the mild reaction conditions used in their application and the selectivity displayed by these enzymes in the bioconversion of a variety of substrates. In fact, nitrile hydratases have been used for manufacturing processes of acrylamide, one of commodity chemicals, at the industrial level not only in Japan but also in the United States and France (5, 35). Most studies on organic synthetic transformation have focused on the use of hydrolases, such as esterases, lipases, proteases, and epoxide hydrolases, which cleave a carbon-heteroatom bond (36). The hydrolases, which cleave a carbon-carbon bond, have also attracted considerable interest as biocatalysts for environmentally benign processes (37). Here, we discovered and characterized CurH probably involved in metabolism of curcuminoids in nature. Some of the reaction products yielded by CurH in the present study exhibit biological effects. For instance, zingerone is able to scavenge and degrade free radicals and reactive oxygen species in the body and inhibits enzymes involved in the generation of these reactive oxygen species (38). Ferulic acid shows antiproliferative and apoptotic effects on human breast cancer or hepatoma cells (39). Zingerone is prepared from vanillin and acetone followed by catalytic hydrogenation (https://patents.google.com/patent/US2381210A/en) or isolated from ginger roots, and ferulic acid is prepared from rice bran pitch, a blackish-brown waste oil generated in the process of the rice bran oil production (40). Our discovery of CurH would pave the way toward the alternative and/or large-scale preparation of compounds enzymatically obtained from curcuminoids or tetrahydrocurcuminoids. β-Diketone hydrolases that were previously reported use the following substrates (33, 37): β-diketo acids (e.g., oxaloacetate and acyl pyruvate), β-diketones with unbranched long alkyl groups (e.g., 4,6-nonanedione and 3,5-heptanedione), cycloalkyl β-diketones (cylcohexan-1,3-dione), and bicyclic β-diketones (6-oxocamphor). On the other hand, CurH accepted β-diketones with aryl-alkenyl groups [curcumin, 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione, demethoxycurcumin, and bisdemethoxycurcumin] as well as β-diketones with aryl-alkyl groups (THC, tetrahydrodemethoxycurcumin, and tetrahydrobisdemethoxycurcumin) as substrates. Until now, a wide variety of curcumin analogs have been synthesized by modifications of the functionality of curcumin, e.g., the aryl side chain, double bond, diketo, or active methylene (41, 42, 43). If CurH recognizes those synthetic analogs as substrates and forms the corresponding hydrolyzed reaction products, bioactive compounds with thus-far unidentified functions may certainly be found among the resultant reaction products. The unusual type of β-diketone hydrolase, CurH, raises the possibility as a valuable biocatalyst to generate many bioactive substances.

Experimental procedures

Isolation of curcumin-degrading bacteria

Curcumin-degrading bacteria were isolated from soil samples by the following enrichment culture method. Step 1 was as follows: a spoonful of a soil sample was added to a test tube containing 10 ml of a screening medium (pH 7.5) consisting of 0.1% (w/v) (NH4)2SO4, 0.05% (w/v) K2HPO4, 0.05% (w/v) KH2PO4, 0.2% (w/v) NaCl, 0.05% (w/v) MgSO4·7H2O, 0.001% (w/v) FeSO4·7H2O, 10% (v/v) tap water, and 0.05% (w/v) curcumin (final concentration), followed by incubation at 28 °C for 1 month. Step 2 was as follows: 1% (v/v) of the cultivated medium was added to the same fresh medium, followed by incubation at 28 °C for 1 month. Step 2 was repeated twice. After the enrichment, the culture broth was spread on agar plates, and colonies that grew on these plates on incubation at 28 °C were isolated.

Phylogenetic analysis

The 16S rDNA sequences were aligned by using the Clustal W program (44). The evolutionary tree for the data set was inferred by the neighbor-joining method (45). The stability of relationships was assessed by performing bootstrap analyses of the neighbor-joining data based on 1000 resamplings.

Biochemical and physiological analysis of a curcumin-degrading microorganism

API Coryne was used to determine the assimilation of some carbon sources and the presence of important enzymes according to the instructions of the manufacturer (bioMérieux).

Purification of the curcumin-degrading enzyme (CurH) from Rhodococcus sp. no. 34

All purification steps were carried out at 0 to 4 °C. KPB (pH 6.0) was used throughout the purification. Centrifugation was performed for 20 min at 16,200g.

Step 1: Preparation of cell-free extracts

Washed cells from 30 L of 2 × YT medium (46) containing 0.000005% (w/v) curcumin were suspended in 6 L of 10 mM KPB (pH 6.0), and then disrupted by sonication at 200 W for 30 min with an Insonator model 201 M (Kubota). The cell debris was removed by centrifugation.

Step 2: DEAE-Sephacel column chromatography

The resultant supernatant solution was applied to a DEAE-Sephacel column (300 ml; GE HealthCare UK Ltd) equilibrated with 10 mM KPB (pH 6.0). Protein was eluted by increasing KCl linearly from 0 to 0.6 M in the same buffer. The active fractions were collected, and then ammonium sulfate was added to give 40% saturation. After centrifugation of the suspension, the supernatant was used for the following step.

Step 3: HiPrep Butyl column chromatography

The enzyme solution from step 2 was applied to a HiPrep Butyl column (20 ml; GE HealthCare UK Ltd) equilibrated with 10 mM buffer containing 2 M ammonium sulfate. Protein was eluted by decreasing ammonium sulfate linearly from 2 to 0 M in the same buffer. The active fractions were pooled and then dialyzed against 10 mM buffer.

Step 4: Resource Q column chromatography

The dialyzed solution was applied to a Resource Q column (6 ml; GE HealthCare UK Ltd) equilibrated with 10 mM KPB (pH 6.0). Protein was eluted by increasing KCl linearly from 0 to 0.4 M in the same buffer. The active fractions were collected, and then ammonium sulfate was added to give 40% saturation. After centrifugation of the suspension, the supernatant was used for the next step.

Step 5: TOYOPEARL PPG column chromatography

The enzyme solution from step 4 was applied to a TOYOPEARL PPG 600 column (1 ml; Tosoh Co., Ltd) equilibrated with 10 mM buffer containing 2 M ammonium sulfate. Protein was eluted by decreasing ammonium sulfate linearly from 2 to 0 M in the same buffer. The active fractions were pooled and then dialyzed against 10 mM buffer.

Step 6: Mimetic Green 3 A6XL column chromatography

The enzyme solution from step 5 was applied to a PIKSI M Test Kit 0013 Mimetic Green 3 A6XL column (1 × 1 cm) (Affinity Chromatography Ltd) equilibrated with 10 mM KPB (pH 6.0). Protein was eluted from the column by increasing KCl linearly from 0 to 2 M in the same buffer, and then the active fractions were collected.

Step 7: Type Q column chromatography

The enzyme solution from step 6 was applied to an IEX Type Q column (4.6 × 50 mm; Nacalai Tesque Co, Inc) equilibrated with 10 mM KPB (pH 6.0). Protein was eluted from the column by increasing KCl linearly from 0 to 1 M in the same buffer. The homogeneity of the purified protein was confirmed by SDS-PAGE.

Enzyme assays

All of the reactions were performed under linear conditions with an appropriate amount of protein and a suitable reaction time, unless otherwise noted. In all the reactions, curcumin was protected from light exposure.

Standard assay A mixture comprised 10 mM KPB (pH 6.0), 0.1 mM curcumin, 5% (v/v) ethanol, 3 mM hydroxypropyl β-cyclodextrin and an appropriate amount of enzyme in a total volume of 100 μl. Ethanol and hydroxypropyl β-cyclodextrin were added to enhance the substrate solubility (47). The reaction was started by the addition of the enzyme and carried out at 28 °C for 30 min. The reaction was stopped by the addition of 100 μl acetonitrile/methanol/water/acetic acid (41:23:36:1, all by volume) to the reaction mixture, and a supernatant was obtained by centrifugation (23,300g, 10 min). The amount of dehydrozingerone in the reaction mixture was determined by HPLC with a Shimadzu LC-6A system equipped with a Cosmosil πnap column (reversed-phase, 4.6 × 150 mm; Nacalai Tesque). The following mobile phase was used at a flow rate of 0.6 ml/min and 40 °C: acetonitrile/methanol/water/acetic acid (41:23:36:1, all by volume). The absorbance was measured at 254 nm for dehydrozingerone. This standard assay was used for all enzyme activity assay experiments except for the substrate specificity and inhibitor analysis.

Inhibitor analysis was carried out under the standard assay B conditions. The standard assay B mixture comprised 100 mM Tris (pH 6.0), 0.1 mM curcumin, 5% (v/v) ethanol, 3 mM hydroxypropyl β-cyclodextrin, 1 mM each metal or inhibitor and an appropriate amount of enzyme in a total volume of 100 μl. In the case of α, α′-dipyridyl or o-phenanthroline among inhibitors, 1% methanol was added to the standard assay B mixture to enhance the solubility. When 5,5′-dithio-bis-2-nitrobenzoic acid was used as an inhibitor, the amount of dehydrozingerone in the reaction mixture was determined by LC-tandem mass spectrometry [a Shimadzu LC-20Avp system, equipped with a Cosmosil πnap column, coupled with an LCMS-8030 apparatus operating in the multiple reaction-monitoring mode]. The tandem mass spectrometry analysis data were acquired with a mass spectrometer with electrospray ionization in the positive and negative modes.

Degradation of each curcumin analog by CurH was examined under the standard assay C conditions. The standard assay C mixture comprised 10 mM KPB (pH 6.0), 0.15 mM substrate, 5% (v/v) ethanol, 3 mM hydroxypropyl β-cyclodextrin, and an appropriate amount of enzyme in a total volume of 100 μl. When bisdemethoxycurcumin or tetrahydrobisdemethoxycurcumin among curcumin analogs was used as a substrate, the HPLC method described for standard assay A was used. The absorbance was measured at 320 nm. As for THC as a substrate, 60% methanol was used as a mobile phase, and the absorbance was measured at 280 nm. When 1,7-bis-(3,4-dimethoxyphenyl)-1,6-heptadiene-3,5-dione, demethoxycurcumin, DHC, or tetrahydrodemethoxycurcumin was used as a substrate, the HPLC conditions were as follows: mobile phase A, 0.05% (v/v) formic acid; and mobile phase B, methanol. Samples were eluted at 1 ml/min using a linear gradient of 5 to 100% of mobile phase B over the course of 15 min after 5 min from the start (5% of mobile phase B). The final composition of 100% mobile phase B was held for 5 min from 20 to 25 min. Chromatographic separation was performed at 40 °C, and the absorbance was measured at 280 nm.

One unit of the CurH activity was defined as the amount of the enzyme that catalyzed the production of 1 nmol dehydrozingerone/min under the standard assay A and B conditions, respectively. In the case of bisdemethoxycurcumin, THC, or tetrahydrobisdemethoxycurcumin instead of curcumin as a substrate, one unit of the activity was defined as the amount of the enzyme that catalyzed the production of 1 nmol 4-hydroxybenzylideneacetone, zingerone, or 4-(4-hydroxyphenyl)-butan-2-one/min, respectively. Specific activity was expressed as units/mg of protein. The kcat values were calculated using a Mr of 42,244 for CurH. Kinetic parameters for demetoxycurcumin, DHC, and tetrahydrodemetoxycurcumin as a substrate were not determined because these substrates are asymmetric compounds. The protein concentrations were determined with a Nacalai Tesque protein assay kit using bovine serum albumin as the standard by the method of Bradford (48).

Draft genome sequence of Rhodococcus sp. no. 34

Rhodococcus sp. No. 34 was cultured at 28 °C for 48 h in 50 ml of 2 × YT media. Cells were harvested by centrifugation, suspended in 10 ml of H2O, frozen in liquid nitrogen, and then crushed using a pestle and mortar. Chromosomal DNA from the strain was prepared by the method of Marmur (49). In order to remove proteins efficiently, a phenol solution (50) was used. Prepared DNA was treated with RNase, and then purified by ultracentrifugation (46). Draft genome sequencing of strain No. 34 was performed using an Illumina Hiseq platform (Hokkaido System Science Co, Ltd). The draft genome sequence was annotated with MiGAP (http://www.migap.org).

Expression and purification of the recombinant curcumin-degrading enzyme (CurH)

The coding sequence of the enzyme was amplified by PCR with genomic DNA extracted from strain No. 34 as a template. The following two oligonucleotide primers were used: a sense primer, 5′-TAAGAAGGAGATATACATATGCCCGAGGCAGTAATCGTCTCAGC -3′ containing an NdeI recognition site (underlined); and an antisense primer, 5′-TTGTCGACGGAGCTCGAATTCTCAGCTGAGGCGCTCGATCACCATTG-3′ containing an EcoRI recognition site (underlined). The amplified DNA was inserted into pET-24a(+) digested with NdeI and EcoRI using an In-Fusion HD Cloning kit (Takara Bio Inc), and checked by DNA sequencing. The resultant plasmid was designated as pET-curH; in this construct, curH was under the control of the T7 promoter.

E. coli BL21-CodonPlus(DE3)-RIL was transformed with pET-curH, and the recombinant cells were used for the overproduction and purification of the recombinant curcumin-degrading enzyme. The transformed cells were incubated with reciprocal shaking at 37 °C in 10 ml of 2 × YT medium containing 50 μg/ml kanamycin and 30 μg/ml chloramphenicol. After overnight cultivation, the entire culture was inoculated into 1 L of the same medium, followed by incubation with shaking at 28 °C for 2 h. IPTG was then added to a final concentration of 0.1 mM to induce the T7 promoter, and further cultivation was carried out at 28 °C for 12 h.

All purification steps were carried out at 0 to 4 °C. KPB (pH 6.0) was used throughout the purification. Centrifugation was performed for 20 min at 16,200g.

Step 1: Preparation of cell-free extracts

Washed cells from 0.5 l of culture broth were suspended in 25 ml of 10 mM KPB (pH 6.0) and then disrupted by sonication at 200 W for 30 min with an Insonator model 201 M. The cell debris was removed by centrifugation.

Step 2: HiPrep Butyl column chromatography

Ammonium sulfate was added to the cell-free extracts to give 50% saturation. After centrifugation of the suspension, the supernatant was applied to a HiPrep Butyl column (20 ml) equilibrated with 10 mM buffer containing 2 M ammonium sulfate. Protein was eluted by decreasing ammonium sulfate linearly from 2 to 0 M in the same buffer. The active fractions were pooled and then dialyzed against 10 mM buffer.

Step 3: Type Q column chromatography

The dialyzed solution was applied to an IEX Type Q column (4.6 × 50 mm) equilibrated with 10 mM KPB (pH 6.0). Protein was eluted by increasing KCl linearly from 0 to 1 M in the same buffer. The homogeneity of the purified recombinant protein was confirmed by SDS-PAGE.

Molecular mass determination

The purified recombinant enzyme sample was applied to a Superdex 200 10/300 GL column (GE HealthCare), which was attached to an ÄKTA purifier (GE HealthCare), and then eluted with 10 mM KPB (pH 6.0) containing 0.2 M KCl at the flow rate of 0.5 ml/min. The absorbance of the effluent was recorded at 280 nm. The molecular mass of the enzyme was calculated from the mobilities of the standard proteins, i.e., glutamate dehydrogenase (290 kDa), lactate dehydrogenase (142 kDa), enolase (67 kDa), myokinase (32 kDa), and cytochrome c (12.4 kDa).

Metal analysis

All glassware was soaked in 1 M HCl overnight and then exhaustively rinsed with distilled water before use. Prior to analysis, the enzyme was dialyzed against 10 mM KPB (pH 6.0). The enzyme sample (1.4 mg/ml) was analyzed with an inductively coupled radiofrequency plasma spectrophotometer, Shimadzu ICPS-8000 (27.120 MHz).

CD analysis

CD measurements were carried out with a Jasco spectropolarimeter, model J-720W (Japan Spectroscopic Company), equipped with a thermal incubation system at 20 °C, with a 0.1-cm light path cell. CD measurements of the purified recombinant enzyme and each mutant were carried out with a 0.1-cm light path cell at the protein concentration of 0.1 mg/ml in 10 mM KPB (pH 6.0) in the far-UV region (200–260 nm). The ellipticity in the CD spectra was normalized as to the protein concentration.

Electrophoresis

SDS-PAGE was performed in a 12% polyacrylamide slab gel according to Laemmli (51). The gel was stained with Coomassie brilliant blue R-250. The relative molecular mass of the enzyme subunit was determined from the relative mobilities of marker proteins, phosphorylase b (97 kDa), bovine serum albumin (66 kDa), ovalbumin (45 kDa), carbonic anhydrase (30 kDa), soybean trypsin inhibitor (20.1 kDa), and α-lactalbumin (14.4 kDa).

Protein structural prediction and visualization

The structural prediction of the full-length CurH analyzed in this study was obtained from the AlphaFold2_mmseq2 database (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb) (26). All structural images were made with PyMOL (Schrodinger, USA; https://pymol.org/2/).

Data availability

The nucleotide sequence data for the 16S rRNA gene and the curH gene reported in this article appear in the DDBJ/GenBank database under accession numbers LC777838 and LC777839, respectively.

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Supporting information

Supporting Figures S1–S16 and Table S1

Acknowledgments

This work was supported in part by a Grant-in-Aid for Scientific Research from MEXT. This work is partly supported by Center for Quantum and Information Life Sciences, University of Tsukuba. The authors thank Professor Kentaro Shiraki (University of Tsukuba) for the use of the CD spectrometer. This paper is dedicated to Professor Arnold L. Demain, who was a great scientist constantly in the forefront of research on secondary metabolism and industrial biotechnology but sadly passed away in April 2020.

Author contributions

Y. H., K. I., A. H., T. K., and M. K. formal analysis; Y. H., K. I., A. H., and K. K. investigation; Y. H. and K. I. writing–original draft; Y. H. and M. K. writing–review and editing; Y. H. and M. K. conceptualization.

Funding and additional information

Y. H. was in part supported by the 10.13039/501100002241 Japan Science and Technology Agency , ERATO (JPMJER1502 ). A. H. was supported by Grant-in-Aid for JSPS Fellows.
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