To verify the hypothesis that cyanobacteria naturally biosynthesising polyphenolic compounds possess an active enzymatic system that enables them to transform these substances, such an ability of the biocatalytic systems of whole cells of these biota was assessed for the first time. One halophilic strain and seven freshwater strains of cyanobacteria representing four of the five taxonomic orders of Cyanophyta were examined to determine the following: (i) whether they contain polyphenols, including flavonoids; (ii) the resistance of their cultures when suppressed by the presence of exogenous hydroxychalcones—precursors of flavonoid biosynthesis and (iii) whether these photoautotrophs can transform hydroxylated chalcones. All examined strains were found to contain polyphenols and flavonoids, and the growth of their cultures was inhibited in the presence of 2′- hydroxychalcone, 2″-hydroxychalcone and 4″-hydroxychalcone. We also confirmed that the examined cyanobacteria trans- formed hydroxychalcones via hydrogenative bio-reduction and formed the corresponding hydroxydihydro derivatives with yields above 90% whenever the substrates were bioavailable for such a conversion. Moreover, we observed that the routes and efficiency of biohydrogenation (and hydroxylation) of chalcones were dependent on the location of the hydroxyl substituent. The final products obtained as the results of biotransformations were extracted from the media and identified by mass spectrom- 1 13 etry (LC-MS/MS) and nuclear magnetic resonance ( HNMR, C NMR, COSY, HSQC). Based on those results, we believe that the very efficient biohydrogenation of hydroxychalcones, which may easy be scaled up for biotechnological purposes, reflects the natural activity of the cyanobacterial defence system, because hydroxydihydrochalcones were less active inhibitors of the growth of cyanobacterial cultures than the corresponding substrates. . . . Keywords Hydroxylated chalcones Biocatalysis Cyanobacteria Regiospecific hydrogenation Introduction abolic pathways in photosynthesising organisms. Structurally, chalcones are composed of two aryl groups (A- and B-rings) Chalcones are a group of plant-derived compounds belonging linked by an open-chain three-carbon unit α,β-unsaturated car- to the flavonoid family that are synthesised through the bonyl system. Naturally occurring chalcones are broadly dis- phenylpropanoid pathway and play a vital role in different met- tributed in various plant species and possess a conspicuously yellow colour (Attar et al. 2011; Ballester et al. 2010). They are biogenetic precursors of all known flavonoids and exist most commonly in hydroxylated form. Chalcones are frequently pro- * Jacek Lipok duced in nature by hydroxylases in the biosynthetic pathways email@example.com of plants (Ni et al. 2004). In view of their relative structural Beata Żyszka-Haberecht simplicity and associated facility of synthesis, coupled with firstname.lastname@example.org their attractive biological activities (Avila et al. 2008; Anna Poliwoda Karaman et al. 2010), chalcones continue to enjoy considerable email@example.com attention from chemists researching new molecular scaffolds for the creation of novel therapeutics (Okoniewska et al. Department of Analytical and Ecological Chemistry, Faculty of Chemistry, University of Opole, Oleska 48, 45-052 Opole, Poland 2016; Ritter et al. 2014; Stompor et al. 2015). 7098 Appl Microbiol Biotechnol (2018) 102:7097–7111 Biocatalysis represents a successful and, in many cases, membrane system that makes them highly suitable hosts for outstanding substitute to standard chemical synthesis (Park expressing the P450 enzymes that participate in et al. 2009) or extraction of chalcones from plant material phenylpropanoids biosynthetic (Melis 1999;Xue andHe (Du et al. 2010). This strategy is particularly valuable for 2015). As prokaryotic microalgae, cyanobacteria are an uncon- microbial whole cell systems, which afford cheaper, scalable, ventional and attractive source of phenylpropanoids and create and more available means of efficiently acquiring derivatives great opportunities for discovering unique secondary metabolites of chalcones (de Carvalho and da Fonseca 2006). Thus far, with potential biological activities due to the facility of mass- transformations of hydroxylated chalcones by whole cell sys- scale cultivation. Moreover, several inherent features of tems have been performed only by heterotrophic microorgan- cyanobacteria, such as oxygenic photosynthesis, amenability to isms and the main reactions of these processes mimic the genetic engineering and capacity to survive in extreme habitats natural metabolic conversion pathways of these molecules in or under chemical stress, make them prominent cell factories for plant kingdom. Therefore, hydrogenation, dehydrogenation, targeted biotransformations (Xue and He 2015). We hypothesise hydroxylation, O-methylation, O-demethylation, glycosyla- then that organisms that produce phenylpropanoids, which are tion, deglycosylation, C ring cleavage of the benzo-γ-pyrone also precursors of flavonoids (Goiris et al. 2014; Singh et al. system, cyclization and carbonyl reduction were found as the 2017a), must possess active enzymatic pathways that enable main ways of transformation (Cao et al. 2015). Microbial them to transform these compounds according to cellular needs. conversion of compounds containing a three-carbon enone To verify this hypothesis, the ability of the biocatalytic moiety has attracted increasing attention, particularly with systems of whole cells of halophilic and freshwater species the recent rediscovery of ene-reductases from the old yellow to transform hydroxylated chalcones was studied. All experi- enzyme family. These flavoenzymes selectively catalyse the ments were performed in batch cultures on both analytical and hydrogenation of activated C=C bonds of α,β-unsaturated preparative scales under conditions mimicking their environ- carbonyl molecules in excellent yield (Fu et al. 2013). Ene- mental response. reductases have also been identified in prokaryotic microalgae, i.e. cyanobacteria (Fu et al. 2013), and used for light-induced, photocatalytic reduction of C=C bonds Materials and methods (Koninger et al. 2016) that conduces to the formation of dihydrochalcones. Because of their high sweet taste, these Chemicals compounds have potential applications in the food industry for the production of low-calorie, multicomponent, nontoxic The substrate 2′-hydroxychalcone was obtained following the and safety sweeteners of natural origin. Consequently, their method of Yadav et al. (Yadav et al. 2012). The substrate 2″- UV-protective, antioxidant and health-promoting properties hydroxychalcone (cat. no. 513067) was purchased from make dihydrochalcones of interest to the pharmaceutical and Sigma-Aldrich (Poznan, Poland). The synthesis of 4″- cosmetics industries (DuBois et al. 1977; Winnig et al. 2007). hydroxychalcone was carried out according to the literature Biocatalytic transformations using microorganisms are antic- method (Tran et al. 2009) under strongly basic conditions ipated to be more selective and environmentally friendly (KOH) using methanol as a solvent. All hydroxylated (Matsuda et al. 2009; Nakamura et al. 2003); however, the mi- chalcone stock solutions were prepared in dimethyl sulfoxide crobes do not always deliver the expected reactions. The major (DMSO) (15 mg/mL) and sterilised by filtration prior to their disadvantage of bioconversion is the prerequisite for an aqueous addition to the medium. All stock solutions were made on the ambience for most enzymatic reactions, which can be a limita- same day of use and kept in dark to prevent photodamage tion for the slightly soluble organic compounds acted as sub- (Zyszka et al. 2017a). All components of the medium were strates. Additionally, a real difficulty associated with the use of purchased from Avantor Performance Materials Poland S.A. biocatalysts is the successive supply of the cofactors NADH or (Gliwice, Poland). Other chemicals were of analytical grade NADPH, which deliver reducing power and chemical energy. In purity, purchased from commercial suppliers: Sigma-Aldrich, case of oxygenic photoautotrophic organisms, the solar light Fluka and Avantor Performance Materials Poland S.A. All the energy captured by photosynthetic systems is converted to elec- solutions for culture preparation and maintenance used in this trochemical energy to regenerate NADPH from NADP via study were prepared using water from Milli-Q water (Merck, photosynthetic electron-transfer reactions. NADPH is the soli- Millipore, Germany), whereas special Millipore Milli-Q A10 tary reducing agent for the reduction of exogenous artificial sub- water of high purity was used for analytical purposes. strates (Nakamura and Yamanaka 2002). Therefore, living cyanobacterial cells that possess the ability to regenerate their Cyanobacterial strains and culture conditions own respective cofactors are hopeful and eminently effective pretender for novel ‘photobiocatalysts’. Additionally, One halophilic strain of cyanobacteria, Spirulina platensis cyanobacterial species possess thylakoids, an intracellular [strain C1 (PCC9438)], and seven freshwater strains, Appl Microbiol Biotechnol (2018) 102:7097–7111 7099 including Anabaena sp. [strain CCALA 007], Anabaena laxa (50, 100, 200, 400, 600 and 1000 μg/ml) was added to a 10- [strain CCALA 805], Aphanizomenon klebahnii [CCALA mL volumetric flask and mixed with 0.5 mL of Folin- 009], Nodularia moravica [strain CCALA 797], Ciocalteu phenol reagent. Then, 1 mL of H O was added to Chroococcus minutus [strain CCALA 055], Merismopedia the mixture and shaken vigorously. After a 2-min incubation glauca [strain CCALA 099] and Synechocystis aquatilis at room temperature, 1.5 mL of Na CO (20%, w/v)solution 2 3 [strain CCALA 190], were used in our experiments. The axe- was added, and the volume was then made up to the mark with nic strain of Spirulina platensis was purchased from the distilled water. The samples were incubated for 2 h in the dark. Pasteur Culture Collection (PCC) (Institute Pasteur, Paris), The mixture was allowed to stand for 2 h with intermittent whereas all freshwater strains were obtained from the shaking. After incubation for 120 min at room temperature in Culture Collection of Autotrophic Organisms (CCALA) the darkness, the absorbance against distilled water was deter- (Institute of Botany of the Academy of Sciences, mined at 760 nm with a UV-Visible spectrophotometer. TPC Czech Republic). was expressed as milligram gallic acid equivalents/g dry- Subcultures of cyanobacteria were revitalised every extract weight (mg GAE/g DW). A reagent blank was pre- 3 weeks by transferring 10-mL aliquots to 50 mL of fresh pared using distilled water. suitable media (Forlani et al. 2011). To prepare the inoculates used to initiate the experimental cultures, all tested Total flavonoid assay cyanobacteria were pre-grown in standard media: MSp (ATCC 1679) medium (pH 9.5) for the halophilic strain and The total flavonoid content (TFC) was determined by the al- BG11 (ATCC 616) or Z8 medium (pH 7.0) for the freshwater uminium chloride colorimetric assay (Zhishen et al. 1999). An strains. The cultures of the tested cyanobacteria were grown at aliquot (1 mL) of extracts or standard solutions of quercetin 24 ± 1 °C with 16-h day (1000 lx light intensity) and 8-h night (6.25, 12.5, 25, 50, 80 and 100 μg/mL) was added to a 10-mL photoperiods, corresponding to the conditions of a long day, in volumetric flask containing 4 mL of distilled water. To the 250-mL Erlenmeyer flasks containing 60 mL of each culture flask was added 0.30 ml of 5% NaNO , and after 5 min, (Allen 1968;Rippka etal. 1979; Żyszka et al. 2017a). 0.3 mL of 10% AlCl was added. After 5 min, 2 mL of 1 M NaOH was added, and the volume was made up to 10 mL with Searching for natural phenolics in cyanobacterial cells distilled water. The solution was mixed and incubated for 15 min in the dark, and the absorbance was then measured Biomass samples of all tested cyanobacterial strains were har- against the blank at 510 nm. The results were expressed as mg vested at the end of logarithmic phase (21-day-old subcul- quercetin equivalents/g dry-extract weight (mg QE/g DW). tures) by centrifugation at 5000×g for 1 min. The pellet was washed twice with distilled water to remove adhering compo- Determination of the nature of phenolic compounds nents of the medium, and subsequently, the cyanobacterial present in cyanobacterial cells cells were immediately freeze-dried at − 50 °C and stored at − 20 °C prior to extraction. In order to confirm the presence of phenolic compounds in the For the efficient extraction of polyphenols and flavonoids, cyanobacterial cells and determine the nature of these sub- 100 mg of lyophilized cells of each strain was sonicated for stances, liquid chromatography coupled with spectrophoto- 30 min in 3 mL of pure methanol using an ultrasonic bath metric and mass spectrometry detection (UHPLC-UV-MS/ (40 kHz, 100 W; Branson, Danbury, Ct, USA) and for a fur- MS) was used for chemical screening of extracts of tested ther 30 min using a Hielscher UP200HT ultrasonic cyanobacterial cells. For this purpose, 500 mg of lyophilized homogeniser (26 kHz, 200 W, Hielscher Ultrasonics GmbH, cells of each strain was homogenised in a ceramic mortar and Teltow, Germany). The homogenates were centrifuged at sonicated for 15 min in pure methanol (5 mL) using an 5000×g for 1 min, and the supernatants were filtered using Ultrasonic sonicator (40 kHz, 100 W; Branson, Danbury, Whatman No. 1 filter paper, pooled, concentrated to 1 mL CT, USA). Then, the samples were centrifuged for 3 min at and sterilised by filtration using a 0.22-μm Millipore filter. 3000×g, and the supernatant was separated and collected for The resultant extracts were stored at 4 °C in the dark for combination with the next fraction extracted in the same way 24 h before determination of total phenolic and flavonoid from the cell debris with 5 mL of fresh methanol. Both meth- content. anol extracts were pooled and concentrated to 5 mL under a stream of nitrogen. To avoid contamination of the mass spec- Total phenolic assay trometer and to reduce matrix effects, chlorophylls and carot- enoids were largely removed from the extracts by adsorbing The total phenolic content (TPC) was determined according to these components on an OASIS® MCX ion exchange column a modified Folin-Ciocalteu method (Singleton et al. 1999). An (6 mL, 200 of mg sorbent, Waters, Milford, MA, USA). After aliquot (100 μL) of extracts or standard solution of gallic acid conditioning the column with 5 mL of methanol, the 7100 Appl Microbiol Biotechnol (2018) 102:7097–7111 concentrated cell-free extract was loaded, and the analytes of of chlorophyll in each experimental or control culture repli- interest were selectively eluted using 5 mL of methanol cate, the growth curves were generated and the growth rates of (Goiris et al. 2014). the examined photoautotrophs were calculated. Finally, the The cell-free extracts were dried under a stream of nitrogen, ratio of the growth rates of the experimental cultures with redissolved in a mixture of 600 μL of methanol and 400 μLof respect to the appropriate controls were determined to illus- formate buffer (4 g of ammonium formate and 300 μLof trate the differences in the dynamics of cyanobacterial growth formic acid per litre of buffer) and centrifuged (10 min, dependent on the strain and the examined hydroxylated 13,000×g). The samples were then subjected to UHPLC- chalcone. The percentage values of the ratios of the growth UV-ESI-MS analysis employing a Dionex UltiMate 3000 rates of the appropriate strain were correlated with the appro- UHPLC system (Sunnyvale, CA, USA) coupled to a priate compound and presented as the heat map in the results. micrOTOF-QII mass spectrometer (Bruker Daltonics, Germany). Chromatographic separation was carried out on a Analytical scale biotransformations of hydroxylated Gemini-NX C18 column (150 mm × 4.6 mm i.d. 3μm, 110 Å) chalcones thermostated at 30° and protected by a Gemini NX pre- column (4 × 3 mm). The mobile phase consisted of acetoni- Screening (analytical) scale biotransformations of all hydroxyl- trile with 0.1% formic acid v/v (eluent A) and water containing ated chalcones were studied according to previously described 0.1% formic acid v/v (eluent B) delivered by a gradient at a methodology (Żyszka et al. 2017a). The cyanobacterial inocu- −1 flow rate 250 μLmin . The program for gradient elution was lum was added to 100-mL Erlenmeyer flasks containing 30 mL 40% A/60% B (0 min), 100% A/0%B (15 min), 100% A/0% of the respective culture medium supplemented with the appro- B (20 min), 40% A/60% B (25 min) and 40% A/60% B priate hydroxylated chalcone stock solution (0.13%, v/v) to ob- (30 min). The injection volume was 20 μl. The autosampler tain a final concentration of 20 mg/L of hydroxylated chalcone. temperature was set at 4 °C, and the presence of polyphenols The stability of the tested chalcones was positively verified in was monitored at 254 nm. appropriate substrate control samples consisting of the solution ESI mass spectra were recorded in positive and negative of the tested flavonoid in sterile cultivation medium, whereas the full scan mode. The electrospray ionisation source parameters culture controls were established as consisted of cyanobacterial were as follows: capillary voltage, 3.5 kV; nebuliser pressure, cells in medium cultivated without the hydroxylated chalcone. 1.2 bar; drying temperature, 200 °C; dry gas flow rate, All experiments, including the controls, were performed at least −1 8.0 L min . Nitrogen was used as both the nebulising and in triplicate and were incubated under adjusted conditions of collision gas. Spectra were recorded in positive and negative light and temperature for 14 days. After this time, the cells and modes. Full-scan spectra were acquired in the range of m/z culture media were separated by filtration followed by centrifu- 50–1000. gation, and each repetition was extracted three times with 10 mL Polyphenols were characterised and identified according to of ethyl acetate. Then, these extracts were combined, dried over accurate measurements of the mass parent ions and fragments anhydrous magnesium sulphate, and the solvent was removed during MS/MS analysis using Metabolite Detect 2.0 software using a vacuum evaporator. The remaining residue dissolved in (Bruker Daltonics, Germany). 200 μL of methanol was subjected to analysis by high- performance planar chromatography (HPTLC) and liquid Determination of the growth of microorganisms chromatography-mass spectrometry (LC-MS). The resulting in the presence of hydroxylated chalcones biotransformation yield was established as the average of the values calculated based on the quantification of the products In order to determine the highest concentration of hydroxyl- obtained with respect to the areas of relevant peaks. ated chalcones that did not induce the sudden death of cyanobacterial cells (Żyszka et al. 2017b), a set of screening Determination of the biotransformation course experiments performed in a concentration range from 5 up to 100 mg/L was arranged. Based on the results of these exper- The products of biotransformation were identified by HPTLC iments, a concentration of 20 mg/L was chosen. and UHPLC-UV-ESI-MS techniques similarly as it was re- The growth of the examined photoautotrophs was assessed ported for dihydrochalcone (Żyszka et al. 2017a). To examine by time-course measurements of total chlorophyll content in the course of biotransformation, the products of this process experimental cultures, as described in our previous paper were separated in HPTLC experiments, on TLC aluminium (Żyszka et al. 2017b). Briefly, representative samples of the plates using a CAMAG Linomat 5 applicator (CAMAG, cells were obtained by centrifugation, then re-suspended in Muttenz, Switzerland). To verify the presence of expected methanol in order to extract chlorophyll. The content of this derivatives of flavonoids, the plates were sprayed with appro- photosynthetic dye was determined spectrophotometrically, priate cerium phosphomolybdate reagent and were gently with the use of Arnon’s formula. Basing on the average levels heated until coloured spots appeared. The retention Appl Microbiol Biotechnol (2018) 102:7097–7111 7101 coefficients (R ) and colours of the spots were recorded using whereas the lowest concentration was observed in the extract CAMAG VisionCats software. of A. laxa. With respect to flavonoid content, S. platensis was The changes in the chemical composition of the biotransfor- ranked only third (6.80 ± 0.34 mg/g QE), with higher contents mation media towards the transformation of the tested chalcones in N. moravica and S. aquatilis, which contained 7.90 ± 0.40 were studied by UHPLC-UV-ESI-MS using the same analytical and 6.94 ± 0.35 mg/g QE, respectively. The lowest TFC strategy as for the determination of the chemical nature of the among the tested strains was found in cells of M. glauca. phenolics contained in the cells of the studied cyanobacteria. The UHPLC-MS/MS analysis allowed us to determine the chemical nature of the phenolic compounds extracted from Preparative-scale biotransformations of hydroxylated the cyanobacterial biomass. Using the set of standards to for- chalcones tify appropriate samples, we confirmed the natural presence of phenolic acids, including protocatechuic, gallic, ferulic and Eight repetitions of the same experiment were performed in sinapic acid, as well as the presence of flavonoids, including 1000-mL Erlenmeyer flasks containing 250 mL of the cultures apigenin, genistein, naringenin, acacetin, biochanin A, quer- to scale-up the biotransformation process. After 14 days of cetin and dihydroquercetin, in all the studied microalgal incubation, the mixtures were extracted with ethyl acetate strains. Therefore, we surmised that the examined photoauto- (each repetition, 3 × 80 mL), dried over anhydrous MgSO trophic microorganisms also possess a pool of adequate en- and the solvent was removed. The products of transformation zymes required for the biosynthesis and transformation of were separated by preparative thin layer chromatography these compounds with prominent antioxidant properties. (PTLC) using a CAMAG system and silica gel glass plates without a fluorescent indicator and were extracted with ethyl The sensitivity of cyanobacterial species acetate, which was then evaporated under a stream of nitro- to hydroxylated chalcones gen. Then, the structures of the isolated products were deter- mined and confirmed by mass spectrometry (MS) and nuclear To verify the assumption about the influence of exogenous 1 13 magnetic resonance techniques: H NMR and CNMR,in- hydroxylated chalcones, the growth of these microorganisms cluding COSYand HSQC correlation experiments. The NMR was examined in terms of chlorophyll content, which is a spectra were measured in DMSO-d6 using a Bruker determinant of the metabolic activity of phototrophs, at the UltraShield 400 MHz spectrometer with tetramethylsilane time of harvest in exponential phase. (TMS) as an internal reference. The influences of structurally similar hydroxylated chalcones differing only in the location of one hydroxyl group at the aromatic ring, i.e. 2′-hydroxychalcone, 2″- Results hydroxychalcone, and 4″-hydroxychalcone, on the growth of the eight studied cyanobacteria are illustrated in Fig. 2. Phenolic composition of tested cyanobacterial species There was marked variation in the content of total chloro- phyll in the cyanobacterial strains (Fig. 2a). The maximum We tested the hypothesis that cyanobacterial cells naturally content of chlorophyll was observed in A. laxa, whereas the contain phenolic compounds and therefore possess the enzy- minimum content of chlorophyll was recorded in M. glauca. matic apparatus enabling them to transform these compounds. The halophilic strain, S. platensis, was generally more resis- Cell-free extracts of eight strains of cyanobacteria belonging tant to the tested hydroxylated chalcones than the freshwater to different genera, including one halophilic strain (S. cyanobacteria, although the level of sensitivity observed was platensis) and seven freshwater strains (A. laxa, Anabaena dependent on the chemical. 2″-Hydroxychalcone inhibited the sp., A. klebahnii, N. moravica, C. minutus, M. glauca and S. growth of the cyanobacterial species most effectively (Fig. aquatilis), were evaluated for the total content of phenolics 2d). The growth of all freshwater species stopped completely (TPC) and flavonoids (TFC). The total amounts of phenolic after 4 days of culture, whereas the growth rate of S. platensis compounds, including flavonoids, were determined spectro- was reduced by 72% during the 14 days of the experiment photometrically, and the results showed that the phenolic com- compared with the appropriate control culture. By contrast, 2′- position differed significantly among the strains (Fig. 1). The hydroxychalcone (in fact, the mixture of 2′-hydroxychalcone TPC varied from 10.23 ± 0.51 to 49.87 ± 2.49 mg/g GAE, and and flavanone) affected cyanobacterial growth in a species- the total flavonoid content ranged between 1.87 ± 0.09 and specific manner (Fig. 2c). This compound entirely inhibited 7.90 ± 0.40 mg/g QE of the extracts. The cyanobacterium S. the growth of the freshwater cyanobacteria M. glauca, N. platensis, which is well-known for its nutraceutical features, moravica, S. aquatilis and A. klebahnii but only limited the was the richest in phenolic compounds among the examined growth rate of S. platensis and A. laxa by 62 and 69%, respec- strains. Relatively high values of TPC were also observed in tively. The most resistant cyanobacteria against 2′- A. klebahnii, Anabaena sp., S. aquatilis and N. moravica, hydroxychalcone were Ch. minutus and Anabaena sp., whose 7102 Appl Microbiol Biotechnol (2018) 102:7097–7111 Fig. 1 Total phenol and flavonoid content of the tested cyanobacterial cell-free extracts growth was suppressed by 42 and 22%, respectively. 4″- tested compounds, 2′-hydroxychalcone (1), 2″- Hydroxychalcone had the lowest negative influence on the hydroxychalcone (2) and 4″-hydroxychalcone (3), which all examined strains, as the cultures of S. aquatilis and contain the three-carbon enone moiety 1,3-diphenyl-2- Anabaena sp. were suppressed by only 15 and 11%, respec- propen-1-one, in biocatalytic transformations conducted by tively. Although the growth of A. klebahnii was completely the suppressed cyanobacteria was studied. inhibited, the growth of other strains was reduced by approx- imately 40 to 53% (Fig. 2b). The effects of the hydroxylated (Bio)conversions of 2′-hydroxychalcone (1) chalcones on the cyanobacterial strains was apparent in less than 4 days from the moment of supplementation, indicating Two products of transformation of 2′-hydroxychalcone were ob- efficient transport of these compounds into the cells (Żyszka et tained in quantifiable amounts. According to concentration, the al. 2017b). The heat map presented in Fig. 3 summarises the first was flavanone (4), whereas 2′-hydroxydihydrochalcone (5) growth inhibition patterns of these compounds as was second (Fig. 4). characterised by the ratios of the growth rates of the experi- The nature of the observed conversions was very interest- mental cultures to the growth rates of the appropriate controls, ing. 2′-Hydroxychalcone was not fully accessible to the expressed as percentage values. cyanobacterial catalysts during cultivation because this com- pound had already undergone isomerisation to flavanone via Biocatalytic transformations of hydroxylated intramolecular rearrangement at the time of introduction to chalcones by cyanobacteria each of the used culture media. This process occurs under fully environmentally friendly conditions (microbial medium To understand the interactions between the hydroxylated at room temperature) and strongly favours the formation of chalcones and cyanobacteria more deeply, the fate of the flavanone, since this substance was detected at a ratio of 20:1 Fig. 2 Influence of hydroxylated chalcones, b 4″-hydroxychalcone, c 2′-hydroxychalcone, d 2″-hydroxychalcone, on the growth of cyanobacterial species. a Appropriate control cultures without hydroxylated chalcones Appl Microbiol Biotechnol (2018) 102:7097–7111 7103 Fig. 3 Heat map depicting the intensity of cyanobacterial growth inhibition under the influence of hydroxylated chalcones. High- intensity (red) cells indicate a high inhibitory effect, and low- intensity (yellow) cells indicate a low inhibitory effect over the initial 2′-hydroxychalcone. This conversion seems to cyanobacterial biotransformation of 2′-hydroxychalcone be natural for 2′-hydroxychalcone because the presence of (Janeczko et al. 2013). This conclusion is supported by the flavanone was even confirmed in the purchased standard. exemplary LC-MS profiles of two samples in Fig. 5a, b: 2′- Independent of this chemical rearrangement, all tested (hal- hydroxychalcone in Z8 medium (substrate control), 2′- ophilic and freshwater) cyanobacterial species were able to hydroxychalcone in the experimental culture of A. laxa after biotransform 2′-hydroxychalcone during the 14 days of incu- 14 days of bio-catalytic interactions and the adequate MS/MS bation to form the same product, albeit with moderate efficien- spectra of the analysed compounds. cy. The MS/MS spectra of the product, the mass of the mo- The results of the scaled-up transformation experiments with lecular ion, its fragmentation pathway and the specific differ- 2′-hydroxychalcone fully confirmed those obtained at the ana- ences compared to the spectrum of 2′-hydroxychalcone clear- lytical scale, both with respect to the chemical nature of the ly suggested the bio-reduction of the propenyl chain and thus products and the dynamics and effectiveness of conversion. the formation of 2′-hydroxydihydrochalcone. The structures Moreover, the experiments performed on the preparative scale of the MS/MS fragment ions in positive ion mode for 2′- yielded sufficient amounts (15–30 mg) of the products for sep- hydroxychalcone and 2′-hydroxydihydrochalcone are pre- aration by PTLC and for fully credible confirmation of their sented in Fig. 5a, b, along with the proposed pattern of structures by MS and NMR. The results of NMR experiments 1 13 fragmentation. ( H, C, HSQC and COSY) performed following preparative The retention time, the high agreement with the pattern of (PTLC) separation of the product undoubtedly confirmed the specific fragmentation and the compatibility with spectro- structures of flavanone and 2′-hydroxydihydrochalcone. scopic data published in the literature confirmed that 2′- The efficiency of the biocatalytic conversions performed hydroxydihydrochalcone was obtained as the product of by each of the examined strains of cyanobacteria was Fig. 4 Conversion of 2′- hydroxychalcone to flavanone (4) and 2′-hydroxydihydrochalcone (5) 7104 Appl Microbiol Biotechnol (2018) 102:7097–7111 Fig. 5 LC-MS profiles of 2′-hydroxychalcone in Z8 medium (substrate hydroxydihydrochalcone (m/z 227.09) and the proposed general routes control) (a)and 2′-hydroxychalcone in culture with A. laxa (experimental of fragmentation of 2′-hydroxychalcone and the corresponding 2′- culture) (b) obtained after 14 days of incubation. Adequate MS/MS hydroxydihydrochalcone based on the fragment ions observed in the spectra of 2′-hydroxychalcone and flavanone (m/z 225.09) and 2′- MS/MS spectra in positive ion mode are also shown established based on quantitative determination using liquid media used. Therefore, its transformation was the result of chromatography coupled with mass spectrometry LC-MS biocatalytic processes during the 14 days of cyanobacterial (Table 1). culture. Notably, these conversions, independent of the strain, The moderate yield of biotransformation of 2′- were completely chemoselective, producing only the corre- hydroxychalcone into 2′-hydroxydihydrochalcone, which sponding 2″-hydroxydihydrochalcone, as shown in Fig. 6. did not exceed 23% for the most effective biocatalyst, A. laxa, Importantly, no other products were detected by TLC and and was less than 5% for S. platensis, seems to be the result of LC-MS separation. specific competition between intramolecular rearrangement The LC-MS profiles of the extracted fractions showed that and the biocatalytic process. The latter was not favourable the fraction containing the product after incubation was more because, as an enzymatic reaction, it had to be preceded by homogenous than the corresponding fraction extracted from the transport of the substrate into cyanobacterial cells. Thus, the substrate control (Fig. 7). We suppose that a type of puri- although less effective, the reduction of the double C=C bond fication process was responsible for this result; that is, the of the three-carbon enone linker of the aromatic rings in cyanobacterial cells were able to clean-up the biotransforma- chalcone molecules tends to be the preferred route of biotrans- tion medium and somehow absorb some impurities introduced formation of these substances by cyanobacteria. together with the substrate. This ability of the tested cyanobacteria may simplify biotechnological processes asso- Biotransformations of 2″-hydroxychalcone (2) ciated with these microalgae. Similar to substrate 1, to confirm the efficiency of conver- In contrast to 2′-hydroxychalcone (1), the B-ring substituted sion and to isolate the product in an amount permitting spec- 2″-hydroxychalcone (2) was stable in each of the microbial troscopic analysis, a 14-day biotransformation of substrate 2 Appl Microbiol Biotechnol (2018) 102:7097–7111 7105 Table 1 The efficiency of the Cyanobacterial strain Substrates biocatalytic transformation of the tested hydroxychalcones by 2′- 2″- 4″- cyanobacteria in experiments hydroxychalcone hydroxychalcone hydroxychalcone carried out at preparative scale 1 2 3 S. platensis 5 (4.3 ± 0.6) 6 (20.9 ± 2.7) 7 (99.3 ± 0.1) Product A. laxa 5 (22.9 ± 0.2) 6 (98.7 ± 0.4) 7 (99.3 ± 0.1) (conversion) [%] Anabaena sp. 5 (16.3± 1.0) 6 (97.5 ± 0.3) 7 (31.2 ± 3.1) 8 (28.4 ± 1.5) 9 (16.2 ± 0.6) A. klebahnii 5 (5.4 ± 0.6) 6 (97.8 ± 0.3) 7 (41.6 ± 4.3) 8 (20.0 ± 3.1) 9 (22.6 ± 0.7) N. moravica 5 (8.7 ± 0.9) 6 (97.5 ± 1.1) 7 (82.2 ± 0.3) 8 (4.4 ± 0.3) 9 (6.8 ± 0.0) Ch. minutus 5 (15.6 ± 0.7) 6 (94.0 ± 0.3) 7 (98.5 ± 0.5) M. glauca 5 (6.7 ± 1.0) 6 (75.6 ± 6.8) 7 (46.5 ± 5.6) S. aquatilis 5 (15.8 ± 0.4) 6 (98.6 ± 0.5) 7 (99.1 ± 0.0) was conducted on a larger scale. The comparison of the H hydrogenated only the C=C bond in the three-carbon enone NMR, C NMR and MS spectra of the product with those of subunit and formed the corresponding dihydrochalcone (Fig. the substrate fully confirmed the lack of the double C=C bond 8). By contrast, the strains Anabaena sp., A. klebahnii and N. of the enone moiety and the presence of the unchanged car- moravica also produced 4″-hydroxy-1,3-diphenylpropan-1-ol bonyl group C=O, thus verifying biocatalytic reduction of this (8), and 4″,x″-dihydroxydihydrochalcone (9) (Fig. 8). bond as the only observed route of transformation by the ex- Notably, the latter three strains belong to the same taxonomic amined cyanobacteria. order, Nostocales, whereas the other examined cyanobacteria The bioconversion of 2″-hydroxychalcone proceeded with represent different orders of Cyanophyta. highly satisfactory efficiency. Most of the tested cyanobacterial The LC-MS profiles of the fractions extracted from the cul- strains, including A. laxa, Anabaena sp., A. klebahnii, N. tures of the strains that converted the substrate only to its moravica, Ch. minutus,and S. aquatilis, converted this sub- dihydro-derivative supported this mode of conversion as well strate into its dihydro-derivative in nearly 100% yield (Table as the ability of the cyanobacteria to purify the transformation 1). Only M. glauca and S. platensis formed the product with media to enhance the concentration of this product. Similar to the lower efficiencies of approximately 76 and 21%, respectively. results for the cultures supplemented with 2″-hydroxychalcone, the activity of these cyanobacteria supports the use of these mi- Bioconversions of 4″-hydroxychalcone (3) croorganisms as biocatalysts. In contrast to the results above, the LC-MS profiles of the Cyanobacterial transformations of 4″-hydroxychalcone (3) re- fractions extracted from experimental cultures of the represen- sulted in the increased abundance of three metabolites: 4″- tatives of Nostocales that transformed the substrate into three hydroxydihydrochalcone (7), 4″-hydroxy-1,3-diphenylpropan- more abundant products revealed that these strains trans- 1-ol (8) and 4″,x-dihydroxydihydrochalcone (9) (Fig. 8). formed 4″-hydroxychalcone differently. Nevertheless, even Only when 4″-hydroxychalcone was used as the substrate in these cases, the identified products were hydrogenated or were the processes of bioconversion strain dependent. S. additionally hydroxylated derivatives of the substrate. The platensis, A. laxa, Ch. minutus, M. glauca and S. aquatilis LC-MS profile of the culture of A. klebahnii (Fig. 9)is an Fig. 6 Biotransformation of 2″- hydroxychalcone catalysed by all tested cyanobacterial strains 7106 Appl Microbiol Biotechnol (2018) 102:7097–7111 Fig. 7 LC-MS profiles prepared after 14 days of incubation of 2″-hydroxychalcone in Z8 medium (substrate control) (a) and 2″-hydroxychalcone in the experimental culture of A. laxa (b) and adequate MS/MS spectra of 2″-hydroxychalcone (m/z 225.09) and 2″-hydroxydihydrochalcone (m/z 227.09) example that fully reflects this mode of biotransformation. structures of all products were determined by MS, HNMR The results for the bioconversion of 4″-hydroxychalcone and C NMR. Moreover, the position of the additional hy- again confirm our finding of the specific inclination of droxyl group identified as the second hydroxyl substituent in cyanobacteria to bio-hydrogenate hydroxychalcones. the A/B ring of 4″,x″-dihydroxydihydrochalcone was The spectroscopic data obtained from the analytical cul- established more precisely. The HSQC and COSY experi- tures were fully confirmed after isolation of the products of ments strongly suggest that this substituent is located at the bioconversion from the scaled-up (preparative) cultures. The 3″ or 5″ position, i.e. ‘ortho’ to the initial 4″ hydroxyl group. Fig. 8 Cyanobacterial transformations of 4″- hydroxychalcone (3) led to the formation of 4″- hydroxydihydrochalcone (7), 4″- hydroxy-1,3-diphenylpropan-1-ol (8) and 4″,x- dihydroxydihydrochalcone (9) Appl Microbiol Biotechnol (2018) 102:7097–7111 7107 Fig. 9 LC-MS profile of the fraction of interest extracted from the culture of A. klebahnii with 4″-hydroxychalcone after 14 days of incubation and adequate MS/MS spectra of 4″-hydroxychalcone (m/z 225.09), 4″-hydroxydihydrochalcone (m/z 227.09) and 4″,x-dihydroxydihydrochalcone (m/z 243.09) Regarding the efficiency of the bioconversion of 4″- remained the dominant product, with yields of 31, 42 and hydroxychalcone as a strain-dependent process, it should be 82%, respectively. The less-abundant product, 4″-hydroxy- emphasised that four (S. platensis, A. laxa, Ch. minutus and S. 1,3-diphenylpropan-1-ol, had yields of 28, 20 and 4%, respec- aquatilis) of the five cyanobacterial strains transformed the tively, whereas dihydroxydihydrochalcone was produced in substrate in the most typical way (in this study)—by 16, 23 and 7% yield, respectively. regiospecific hydrogenation of the enone subunit to form 4″- hydroxydihydrochalcone with > 99% yield (Table 1). By con- Spectral data of isolated metabolites trast, M. glauca, which was a less effective biocatalyst for both 2′-and 2″-hydroxychalcones, transformed 4″- hydro Flavanone (4) xychalcone with 47% yield. This study is the first to report 1 6 that the halophilic cyanobacterium S. platensis is a highly H NMR (400 MHz) (DMSO-d ) δ (ppm)—7.81 (dd, 1H, H- effective biocatalyst. In addition, the activities of the 5), 7.39–7.62 (m, 6H, H-7, H-2′,H-3′,H-4′,H-5′,H-6′), 7.10 cyanobacterial enzymatic systems of Anabaena sp., A. (m, 2H, H-6, H-8), 5.68 (dd, 1H, H-2), 3.26 (dd, 1H, H-3), 13 6 klebahnii and N. moravica transformed 4″-hydroxychalcone, 2.84 (dd, 1H, H-3); C NMR (100 MHz) (DMSO-d ) δ leading to the formation of three products that were identical (ppm)—191.63 (C-4), 161.11 (C-8a), 138.95 (C-1), 136.34 for all three strains. These substances were sufficiently abun- (C-7), 128.59 (C-3′,C-5′), 128.57 (C-4′), 126.67 (C-5), dant to be identified. Importantly, 4″-hydoxydihydrochalcone 126.36 (C-2′,C-6′), 121.51 (C-6), 120.68 (C-4a), 118.09 (C- 7108 Appl Microbiol Biotechnol (2018) 102:7097–7111 8), 78.85 (C-2), 43.54 (C-3); ESI-MS m/z 225.0865 [M + H] 4″-hydroxy-1,3-diphenylpropan-1-ol (8) (calcd. for C H O + H, 225.0910); the retention time 15 12 2 19.84 min. As an example, transformation of 4″-hydroxychalcone (3) (80 mg) in the Anabaena sp. culture yielded 21 mg of com- 1 6 2′-hydroxydihydrochalcone/2′-hydroxy-1,3-- pound 8; H NMR (400 MHz) (DMSO-d ) δ (ppm)—9.14 (s, diphenylpropan-1-one (5) 1H, OH), 7.30 (d, 4H, H-2″,H-6″,H-2′,H-6′), 7.21 (dt, 1H, H-4′), 6.94 (dd, 2H, C-3′,C-5′), 6.64 (dd, 2H, C-3″,C-5″), As an example, 14-day transformation of 2′-hydroxychalcone 4.48 (dt, 1H, CH-OH), 2.44 (m, 2H, H-2), 1.81 (m, 2H, H-3), 13 6 (1) (80 mg) in the A. laxa culture yielded 17 mg of compound 1.23 (s, 1H, OH); C NMR (100 MHz) (DMSO-d ) δ 1 6 5; H NMR (400 MHz) (DMSO-d ) δ (ppm)—11.87 (s, 1H, (ppm)—155.21 (C-4″), 146.40 (C-1″), 132.21 (C-1′), 129.06 OH), 7.95 (dd, 1H, H-6′), 7.51 (ddd, 1H, H-4′), 7.29 (t, 2H, H- (C-3′,C-5′), 128.00 (C-2″,C-6″), 126.64 (C-4′), 125.81 (C-2′, 3″,H-5″), 7.28 (t, 2H, H-2″,H-6″), 7.18 (t, 1H, H-4″), 6.96 C-6′), 115.05 (C-3″,C-5″), 71.98 (C-1), 41.47 (C-3), 30.69 (dd, 1H, H-3′), 6.93 (td, 1H, H-5′), 3.36 (t, 2H, H-3), 2.95 (t, (C-2); ESI-MS m/z 229.1103 [M + H] (calcd. for C H O + 15 16 2 13 6 2H, H-2); C NMR (100 MHz) (DMSO-d ) δ (ppm)— H, 229.1223); the retention times 7–9.68 min. 205.21 (C-1), 160.61 (C-2′), 141.03 (C-1″), 136.04 (C-4′), 130.79 (C-6′), 128.44 (C-3″,C-5″), 128.34 (C-2″,C-6″), 4″,x″-dihydroxydihydrochalcone/4″,x″-dihydroxy-1,3-- 125.98 (C-4″), 120.38 (C-1′), 119.26 (C-5′), 117.65 (C-3′), diphenylpropan-1-one (9) (x = 3″ or 5″) 40.07 (C-3), 29.41 (C-2); ESI-MS m/z 227.0948 [M + H] (calcd. for C H O + H, 227.1066); the retention times 1– As an example, transformation of 4″-hydroxychalcone (3) 15 14 2 22.26, 4–21.91 min. (80 mg) in the Anabaena sp. culture yielded 14 mg of com- 1 6 pound 9; H NMR (400 MHz) (DMSO-d ) δ (ppm)—9.23 (d, 2″-hydroxydihydrochalcone/2″-hydroxy-1,3-- 1H, OH (3″ or 5″)), 9.08 (d, 1H, OH (4″)), 7.96 (dd, 2H, H-2′, diphenylpropan-1-one (6) H-6′), 7.62 (ddd, 1H, H-4′), 7.52 (t, 2H, H-3′,H-5′), 7.30 (t, 1H, H-2″ or H-6″), 7.08 (ddd, 1H, H-2″ or H-6″), 6.68 (dd, As an example, transformation of 2″-hydroxychalcone (2) 1H, H-3″ or H-5″), 3.27 (t, 2H, H-2), 2.86 (t, 2H, H-3); C (40 mg) in the A. klebahnii culture yielded 38 mg of com- NMR (100 MHz) (DMSO-d ) δ (ppm)—200.72 (C-1), 1 6 pound 6; H NMR (400 MHz) (DMSO-d ) δ (ppm)—9.39 158.38 (C-3″ or C-5″), 155.65 (C-4″), 136.52 (C-1″), 133.35 (s, 1H, OH), 7.97 (dd, 2H, H-2′,H-6′), 7.63 (ddd, 1H, H-4′), (C-4′), 131.18 (C-1′), 129.25 (C-3′,C-5′), 128.76 (C-2″or C- 7.52 (t, 2H, H-3′,H-5′), 7.12 (dd, 1H, H-6″), 7.01 (ddd, 1H, H- 6″), 128.58 (C-2″or C-6″), 127.93 (C-2′,C-6′), 115.34 (C-3″or 4″), 6.78 (dd, 1H, H-3″), 6.70 (td, 1H, H-5″), 3.27 (t, 2H, H-3), C-5″), 39.548 (C-3), 28.72 (C-2); ESI-MS m/z 243.0903 [M + 13 6 + 2.86 (t, 2H, H-2); C NMR (100 MHz) (DMSO-d ) δ H] (calcd. for C H O + H, 243.1015); the retention times 15 14 3 (ppm)—199.64 (C-1), 155.21 (C-2″), 136.61 (C-1″), 133.13 8–12.89 min. (C-4′), 129.89 (C-6″), 128.75 (C-3′,C-5′), 127.93 (C-2′,C-6′), 127.14 (C-1′), 127.08 (C-4″), 118.91 (C-5″), 114.85 (C-3″), 38.10 (C-3), 24.84 (C-2); ESI-MS m/z 227.0948 [M + H] Discussion (calcd. For C H O + H, 227.1066); the retention times 2– 15 14 2 17.31, 5–17.80 min. The production of a wide range of secondary metabolites is a common adaptation of cyanobacteria to compete successfully 4″-hydroxydihydrochalcone/4″-hydroxy-1,3-- in different ecosystems. Because of their phototrophic lifestyle diphenylpropan-1-one (7) and constant exposure to high oxygen and radical stresses, these biota have a high capability for producing plentiful effi- As an example, transformation of 4″-hydroxychalcone (3) cient protective chemicals against oxidative and radical (40 mg) in the Ch. minutus culture yielded 37 mg of com- stressors and exhibit adaptive responses to oxidative stresses 1 6 pound 7; H NMR (400 MHz) (DMSO-d ) δ (ppm)—9.14 (s, by stimulating their intrinsic antioxidant defence systems 1H, OH), 7.95 (dd, 2H, H-2′,H-6′), 7.62 (ddd, 1H, H-4′), 7.51 based on mediator compounds, including polyphenolic sub- (t, 2H, H-3′,H-5′), 7.05 (dd, 2H, H-2″,H-6″), 6.65 (dd, 2H, H- stances (Babić et al. 2016). Previously published data indicate 3″, H-5″), 3.30 (t, 2H, H-2), 2.82 (t, 2H, H-3); C NMR that the phenolic acids and flavonoids in cyanobacterial cells (100 MHz) (DMSO-d ) δ (ppm)—199.46 (C-1), 155.46 (C- may be responsible for their antioxidant properties (Singh et 4″), 136.50 (C-1″), 133.32 (C-4′), 131.15 (C-1′), 129.28 (C-3′, al. 2017a), and thus a higher content of phenolics in C-5′), 128.76 (C-2″,C-6″), 127.96 (C-2′,C-6′), 115.19 (C-3″, cyanobacterial species may be presumed as an adaptation C-5″), 39.50 (C-3), 28.76 (C-2); ESI-MS m/z 227.0948 [M + strategy of these organisms against abiotic stresses in their H] (calcd. for C H O + H, 227.1066); the retention times specific habitats. Therefore, the properties of these organisms 15 14 2 3–16.22, 6–16.35 min. may also include functional values such as free-radical Appl Microbiol Biotechnol (2018) 102:7097–7111 7109 quenching, metal chelation and ROS-scavenging activity 10% Pd-C as a catalyst and EtOAc as a solvent. Although (Singh et al. 2017a). Our evaluations are consistent with liter- dihydrochalcones are obtained in good yields under these condi- ature reports showing that metabolites such as flavonoids are tions (Vijaya Bhaskar Reddy et al. 2017), the efficiency of these involved in the defence system of cyanobacteria against ad- processes is far lower than hereby reported 99%, highlighting the verse conditions (Singh et al. 2017b). value of the presented cyanobacterial catalytic systems. This bio- Independent of the intrinsic content of polyphenolics, the technological approach does not require chemical catalysts, presence of these chemicals outside cyanobacterial cells may which may be highly specific and therefore expensive (Luan et significantly influence their growth and metabolism, as dem- al. 2014). For comparison, biotransformation of 2′- onstrated for naringenin (Żyszka et al. 2017b). This inhibition hydroxychalcone by yeast strains and filamentous fungi cultures is higher when the substance is structurally related to hydrox- afford the corresponding dihydrochalcone with 2–98% substrate ylated chalcones, which are biosynthetic precursors of all fla- conversion (Janeczko et al. 2013). And transformation of another vonoids. We found that the tested hydroxychalcones inhibited hydroxylated chalcone, chalconaringenin, in the culture of the growth of all eight examined strains representing four of Rhodococcus sp. led to dihydrochalconaringenin (phloretin) in five cyanobacterial taxonomic orders, and this inhibition 84% yield (Stompor et al. 2013). depended more on the structure of the hydroxychalcone than Remarkably lower efficiency of biotransformation of 2′- on the microbial strain. Our results are consistent with those hydroxychalcone together with its weaker inhibitory action obtained by Nakai and co-authors, who observed that the compared to 2″-hydroxychalcone drew our special attention. growth of the cyanobacterium Microcystis aeruginosa was Basing on the results of our study, we concluded that the inhibited in the presence of polyphenols such as ellagic and reason was the lack of bioavailability of 2′-hydroxychalcone, gallic acids and catechin (Nakai et al. 2005). The presence of which tended to intramolecular isomerisation into flavanone. flavonoids, such as 4′,5-dihydroxyflavone, apigenin, and The effective chemical formation of flavanone from the initial luteolin, also suppressed the growth of M. aeruginosa signif- chalcone proceeded under mild and totally environmentally icantly, and this effect increased with the concentration of friendly conditions (microbial media at room temperature) these compounds (Huang et al. 2015). Importantly, in the case and thus may be considered a fully ‘green’ process. This rear- of our study, the position of the substitution of the hydroxyl rangement, however, was not the only side process with un- group is likely responsible for this effect, suggesting that this expected benefits. All examined strains turned out to have a aspect is crucial for understanding the mechanism of interac- tendency to purify, probably by utilisable absorption, the bio- tions between chalcones and cyanobacterial species. transformation media of small amounts of impurities that ac- Studying mentioned interactions, we observed that each of the companied the substrates. In such circumstances, a simplified tested hydroxychalcones was bioconverted, mainly into the cor- extraction of pure product together with the mild conditions of the process significantly supports the use of cyanobacterial responding hydrogenated derivative. Moreover, the processes of (bio)hydrogenation of the C=C bond in the three-carbon enone biocatalysis in the hydrogenation of hydroxychalcones. subunit of the hydroxylated chalcones were regiospecific and The results presented have important implications when mainly proceeded by cyanobacterial transformation. This finding compared to those obtained during biotransformations of fully corresponds with the results of our previous study on the chalcone published previously (Żyszka et al. 2017a). The bio-reduction of chalcones (Żyszka et al. 2017a) and has impor- same strains of cyanobacteria transformed unsubstituted tant ecological implications because hydroxydihydrochalcones chalcone to its dihydroderivatives; however, only A. laxa have approximately 15–20% lower inhibitory effects on the and S. aquatilis performed this transformation with high effi- growth of cyanobacteria (data not presented). Therefore, this ciencies similar to those of the corresponding conversion of route of transformation may be considered a natural component 4″-hydroxychalcone. In the case of this compound, the routes of the cyanobacterial defence system since these organisms suc- and efficiency of biohydrogenation and hydroxylation were cessfully faced dozens of extreme environmental stresses during dependent on the presence and location of the hydroxyl sub- evolution (Żyszka et al. 2017b). This hypothesis is well- stituent. This information strengthens our conclusions on the supported by the results of the present study because the most structural dependence of the biotransformations of chalcones active growth inhibitor, 2″-hydroxychalcone, was converted with and their derivatives by cyanobacteria. In this way, these re- the highest yields. Thus, we confirmed that the inhibition of the sults support the usefulness of these substances as molecular growth of cyanobacterial cells supplemented with hydroxylated probes for more detailed studies of the enzymes responsible chalcones does not affect their activity as biocatalysts. This issue for these transformations. The existence of such enzymes in seems to be especially important, since whenever the substrates cyanobacterial cells and their catalytic mechanism has only were available for cyanobacterial catalysis, bioconversion was been postulated (Fu et al. 2013). highly efficient, with yields often above 99%. This finding is notable compared to the chemical hydrogenation of chalcones, Funding information This work was supported by Polish National which requires 24–48 h in an atmosphere of H Science Centre (NCN) grant number 2016/21/N/NZ9/02310. with the use of 2 7110 Appl Microbiol Biotechnol (2018) 102:7097–7111 different evolutionary lineages. J Phycol 50:483–492. https://doi. Compliance with ethical standards org/10.1111/jpy.12180 Huang H, Xiao X, Ghadouani A, Wu J, Nie Z, Peng C, Xu X, Shi J (2015) Conflict of interest The authors declare that they have no conflict of Effects of natural flavonoids on photosynthetic activity and cell interest. integrity in Microcystis aeruginosa. Toxins 7:66–80. https://doi. org/10.3390/toxins7010066 Ethical approval This article does not contain any studies with human Janeczko T, Gładkowski W, Kostrzewa-Susłow E (2013) Microbial trans- participants or animals performed by any of the authors. formation of chalcones to produce food sweetener derivatives. J Mol Catal B Enzym 98:55–61. https://doi.org/10.1016/j.molcatb.2013. 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Applied Microbiology and Biotechnology – Springer Journals
Published: Jun 4, 2018
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