N-methyl AEP (M1AEP). Here, we describe a heterogeneous group of FAD-dependent oxidoreductases that efficiently oxidize M1AEP to directly generate PAA, thus expanding the versatility and usefulness of the hydrolytic AEP degradation pathways. Furthermore, some of these enzymes can also efficiently oxidize plain AEP. By doing so, they surrogate the role of PhnW in organisms that do not possess the transaminase and create novel versions of the AEP degradation pathways in which PAA is generated solely by oxidative deamination.Phosphonates—compounds containing a direct C–P bond—represent an important source of phosphorus in some environments. The most common natural phosphonate is 2-aminoethylphosphonate (AEP). Many bacteria can break AEP down through specialized “hydrolytic” pathways, which start with the conversion of AEP into phosphonoacetaldehyde (PAA), catalyzed by the transaminase PhnW. However, the substrate scope of these pathways is very narrow, as PhnW cannot process other common AEP-related phosphonates, notably   •2-aminoethylphosphonate (AEP) is the most common natural phosphonate•FAD oxidoreductases are often encoded in bacterial gene clusters for AEP catabolism•N-monomethyl-AEPThese enzymes serve to degrade the naturally occurring •Some of these enzymes also act on AEP, creating a novel AEP catabolic pathway Enzymology; Bacteriology; Microbial metabolism This is exemplified by the archetypal catabolic pathway, glycolysis, which is commonly described as a route for the degradation of glucose, yet it also catabolizes several other sugars which are funneled into glycolysis by various ancillary enzymes or short “tributary” pathways.,,,,,,,,,Herein, we looked for ancillary enzymes that might expand the versatility of the catabolic pathways for 2-aminoethylphosphonate , which is the most prevalent natural phosphonate.,In the most common case, the hydrolytic degradation pathway begins with a reaction catalyzed by the transaminase PhnW, which converts AEP into phosphonoacetaldehyde (PAA)phnWX (or phnWYA) combination.(R)-2-amino-1-hydroxyethylphosphonate (R-HAEP). The reaction catalyzed by PbfA generates PAA, which can be subsequently processed by PhnX (or by PhnY).Given the widespread diffusion of both the PhnWX and PhnWYA pathways, it is biologically plausible that accessory enzymes may have evolved to convey into these catabolic routes other naturally occurring, AEP-related phosphonates. As a case in point, we recently reported the identification and characterization of one enzyme that is encoded in over 13% of the bacterial gene clusters containing the phnWX type in search of additional enzyme-coding genes recurring within these clusters. As in the case of PbfA, we relied on the assumption that enzymes able to “funnel” different compounds into the AEP hydrolytic pathway would be genomically associated with phnW and phnX.Encouraged by the discovery of PbfA, we performed further genomic analyses, parsing firstly the bacterial gene clusters of the ,,R)-HAEP; in particular, they could act on N-methyl AEP (M1AEP), which is another frequently found natural product,,,,,,During this analysis, we were struck by the frequent presence, in these clusters, of genes apparently coding for oxidoreductases dependent on flavin adenine dinucleotide (FAD). In terms of sequence, these putative enzymes were rather heterogeneous and fell into at least three distinct subgroups, which we termed PbfB, PbfC, and PbfD. However, irrespective of their subgroup, all these proteins showed a substantial similarity to amine oxidoreductases, i.e., enzymes that oxidize various primary and secondary amines, leading to the formation of aldehydes or ketones.1AEP to PAA, which can be subsequently processed to generate inorganic phosphate. We characterized in detail representative enzymes from the PbfC and PbfD groups, highlighting substantial mechanistic differences between them. In particular, a PbfD enzyme from Mariniblastus fucicola was found to oxidize plain AEP and M1AEP with comparable efficiencies, suggesting that in vivo this enzyme can perform a double duty. By generating PAA not only from M1AEP but also from AEP, this oxidoreductase would in effect surrogate the function of the transaminase PhnW, whose gene is absent in the M. fucicola gene cluster.Indeed, we present evidence that enzymes from all three groups are able to convert MSalmonella entericaVibrio splendidusphnX or with the phnY/phnA duo (phnWX and phnWYA clusters). Conversely, we excluded from the analysis those cases in which phnW was located close to the pepM and ppd genes, which are known to participate in AEP biosynthesis.,,We visually inspected the genomic contexts of the genes implicated in the hydrolytic AEP degradation routes, beginning with the pyridoxal phosphate (PLP)-dependent aminotransferase PhnW, which converts AEP to PAA . BecausephnWX and the phnWYA type. Furthermore, upon expanding our analysis, we observed that genes for similar FAD-dependent enzymes were sometimes also associated with phnX alone . Exemplary cases of different cluster types that include these enzymes are shown in By examining the AEP degradative clusters in many organisms, we noticed that such clusters often included genes apparently coding for FAD-dependent oxidoreductases (Pfam: PF01266). These genes were found associated with clusters of both the Despite sharing the same conserved domain, the encoded protein sequences displayed a remarkable diversity, often retaining a very limited similarity within the FAD-dependent oxidoreductase domain . Phyloge,,2O2. However, for the mammalian sarcosine dehydrogenase (SarDH) the physiological electron acceptor is a flavoproteinHelicobacter pylori (DadA) the electron acceptor is presumably a quinoneWhile these three subgroups of FAD-dependent proteins were sequence-wise quite dissimilar, they were consistently related to enzymes that catalyze the oxidative deamination of primary or secondary amines. This type of reaction leads to the formation of intermediate imine species, which spontaneously hydrolyze with the eventual generation of carbonyl groups.Enzymes from the PbfB, PbfC, and PbfD subgroups were never co-present in the same cluster. This observation, together with the fact that the different FAD-dependent enzymes were invariably similar to amine oxidoreductases, suggested that they all might be performing essentially the same transformation in aminophosphonate degradation.But what could be the function of these oxidoreductases? As outlined in the N-methyl derivative of AEP (M1AEP), which is reportedly quite common in nature.,,,,,,Based on these considerations, our initial hypothesis was that PbfB, PbfC, and PbfD enzymes could all catalyze the oxidative deamination of some AEP derivative(s) alkylated on the amino group. In particular, they could act on the 1AEP would produce methylamine and PAA, which could be then conveniently hydrolyzed by PhnX. Given the structural similarity of M1AEP to plain AEP, we also expected that the FAD enzymes could oxidize AEP with some efficiency, producing PAA with the release of ammonia. This reaction would be redundant with respect to the reaction catalyzed by PhnW, which also generates PAA from AEP  and Mariniblastus fucicola (henceforth termed PbfD2) were purchased with codon optimization for expression in E. coli. The genes were inserted in the bacterial expression vector pET28a and the recombinant His-tagged proteins were expressed and purified from E. coli cells as described in the Synthetic genes encoding the PbfB protein from All four recombinant proteins appeared to be soluble to some extent; however, despite our efforts, PbfB could not be purified by metal-affinity chromatography from cell extracts, which greatly limited its characterization (see in the following section). On the other hand, we obtained the recombinant PbfC, PbfD1, and PbfD2 enzymes in pure form and proceeded to characterize their catalytic activity toward a number of potential substrates, including some non-phosphonate AEP analogs.,,,,We initially used 96-well plate assays to test the actual ability of the FAD enzymes to oxidize various amino compounds, employing the artificial dye DCPIP as the final electron acceptor and PMS as an electron mediator.2O2, which can be used by horseradish peroxidase to oxidize the chromogenic compound o-dianisidine, turning the solution to deep pink. Contrary to the DCPIP assay, this method entailed a single readout taken after a relatively long incubation, thus it could only qualitatively assess the capacity of the enzyme to reduce molecular oxygen.A second type of colorimetric assay served to evaluate the ability of the enzymes to oxidize potential substrates with electron transfer to oxygen. The assay measured the formation of H1AEP, as well as some activity toward the closely related compound N-ethyl-AEP . In particular, PbfC did not react appreciably with other compounds: moreover, activity of this enzyme was readily observed in the DCPIP assay, but barely detectable in the peroxidase-based assay, suggesting that PbfC inefficiently uses molecular oxygen as the electron acceptor.The results of these microtiter assays are reported in 1AEP in both the DCPIP-based assay and the peroxidase-o-dianisidine assay. These two enzymes also showed a clear reactivity toward AEP. Furthermore, PbfD1, but not PbfD2, appeared to appreciably oxidize N,N-dimethyl-2-aminoethylphosphonate (M2AEP).On the other hand, PbfD1 and PbfD2 enzymes manifested a strong oxidative activity toward M1AEP (such as sarcosine) or of AEP . The enzymes also failed to oxidize 2-hydroxyethyl phosphonate, an analog of AEP that contains a hydroxyl group in place of the amino group (data not shown).None of the enzymes reacted with non-phosphonate analogs of M1AEP, other details of the hypothesized reaction had to be confirmed.Although the microtiter assays indicated the ability of the FAD enzymes to oxidize M1AEP (but not AEP) could generate AEP and formaldehyde, instead of PAA and methylamine. To test for this possibility, we employed a spectrophotometric assay in which the three oxidoreductases were incubated with M1AEP for 30 min; afterward, NADH and alcohol dehydrogenase (ADH) were added to detect the oxidation of NADH in case formaldehyde had been produced. The observation that NADH was not appreciably consumed (data not shown) implied that formaldehyde had not been formed (PAA is not a substrate for ADH). Conversely, when the oxidoreductases were incubated for 30 min with M1AEP and PhnX, which hydrolyzes PAA to acetaldehyde, and then the reaction mixture was supplemented with ADH and NADH, the latter was quickly oxidized. This observation was fully consistent with the hypothesis that the PbfC and PbfD enzymes produce PAA.In principle, the oxidation of M1AEP) released ammonia, as revealed by a coupled assay with GDH (data not shown). Other controls established that phosphate was not released in the reaction of the oxidoreductases with either M1AEP or AEP, except when PhnX was also present .We also demonstrated that the reaction of PbfD1 and PbfD2 with AEP , as described earlier. The second assay was conducted in the absence of artificial electron acceptors and coupled the production of PAA with the consecutive reactions of PhnX  and of alcohol dehydrogenases .Consistent with the findings of the microtiter assays in To establish the catalytic performances of the PbfC and PbfD oxidoreductases toward naturally occurring substrates, we measured the activity of these enzymes using the continuous spectrophotometric assays that worked best for each of them .The pertinent titrations fitted well to the Michaelis-Menten equation G–4I and cat/KM values) of the three enzymes toward M1AEP were consistently in the range 2×103 - 5×104 M−1s−1, whereas KM values ranged from about 0.2 mM to about 5 mM. These performances, while not outstanding, are comparable to those reported for other FAD-dependent oxidoreductases acting on relatively small substrates. For example, glycine oxidase from B. licheniformis , which seemed mostly due to a weaker substrate binding (100-fold difference in KM). In comparison, PbfD2 showed only a 4-fold difference in kcat/KM values between M1AEP and plain AEP, while PbfD1 specificity was somewhat intermediate, with an 18-fold discrimination toward AEP.PbfC showed the highest apparent catalytic efficiency in the oxidation of MN-ethyl-AEP and N-propyl-AEP, which are not known to occur in nature. The results, shown in While the experiments earlier refer to naturally occurring aminophosphonates, we also performed assays using V. vulnificus in pure form. Therefore, to test its activity, we resorted to performing assays on the extracts of E. coli cells that expressed this enzyme. In this case, however, assays based on the formation of H2O2  and as a tool that bacteria employ for surviving in nutrient-limited ecological niches. At least three types of phosphonate-degrading pathways have been identified so far, schematically distinguished based on the mechanism by which the C-P bond is ultimately cleaved—i.e., through either a hydrolytic, radical, or oxidative reaction.,,The hydrolytic pathways for AEP degradation are a case in point. While AEP is indeed the most abundant natural phosphonate,N-alkylated derivatives of AEP and (R)-HAEP, by either newly developed or adapted literature methods. M1AEP and M2AEP were obtained by reaction of diethyl 2-bromoethylphosphonate with the corresponding mono- and dialkylated amines, followed by acidic deprotection. M3AEP became accessible from diethyl 2-(dimethylamino)-ethylphosphonate by alkylation with methyl iodide following an adapted literature procedure.R)-M3HAEP was obtained from (R)-HAEP by methylation, while the (R)-M2HAEP was accessible by an Eschweiler-Clark approach. The monomethylated derivative (R)-M1HAEP was accessible from (R)-diisopropyl -1-hydroxyethyl)phosphonateThis study deals with the recurrent presence of genes encoding putative (and only weakly interrelated) FAD enzymes within bacterial clusters for AEP degradation. With “recurrent,” we refer to genes that are repeatedly found in such clusters—but whose presence is neither constant nor limited to a specific cluster type. The optional but frequent occurrence of these genes is very consistent with them encoding ancillary enzymes, acting on compounds related to the main substrate of the pathway. Establishing the precise function of these accessory enzymes, however, is often not straightforward. For example, to convincingly establish the specificity of the oxidoreductases described in this work we had to test a variety of phosphonatesA. baumannii is annotated in GenBank as a member of the TIGR03364 family , generating PAA that can be then hydrolyzed by PhnX to obtain acetaldehyde and inorganic phosphate.We have now provided strong evidence (in the case of PbfB) or complete proof (for the PbfC and PbfD proteins) that these FAD enzymes catalyze the oxidative deamination of the 1AEP is frequently found in nature , so the most obvious possibilities would be a quinone . Our data hint that the PbfB-type enzymes can carry out the M1AEP oxidation in the absence of acceptor dyes , through a reaction somehow resembling that of dimethylglycine oxidase .2AEP was just 6-fold less efficient than the oxidation of M1AEP .More relevant from a biological viewpoint was the ability of PbfD2 from th M1AEP . Such a  cluster  and appa1AEP oxidoreductase at least three times, co-opting different ancestor enzymes to perform the task. PbfB sequences might have emerged last, as they are the less divergent , and Betaproteobacteria, but also frequently observed in bacteria belonging to the FCB group and occasionally in Terrabacteria. The PbfD group of enzymes might have been the most ancient to emerge: this is suggested both by the large divergence between the sequences of these enzymes  and by the relatively broad distribution among bacterial phyla. PbfD homologs are encoded in AEP degradation clusters in Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, and bacteria from the FCB and Planctomycetota, Verrucomicrobiota, and Chlamydiota (PVC) groups (mostly Planctomycetota). Notably, PbfD homologs not associated with phnWX/phnWYA clusters were also detected in many Terrabacteria (mostly cyanobacteria).The heterogeneity of the FAD-dependent enzymes analyzed in this study, both in terms of sequence and mechanistic details, outlines a remarkable instance of convergent evolution. It suggests that evolution has “invented” an ancillary Mivergent  and theibacteria . On the Rhodoferax sediminis was found to possess pbfB-phnWYA genes, while the closely related Rhodoferax koreense possesses pbfD-phnX instead (phnW and phnX for example), and it is logical that these would move between organisms as a part of complete clusters rather than separately.The varying presence of PbfB, PbfC, and PbfD enzymes in closely related organisms is consistent with a significant degree of horizontal gene transfer involving these pathways. For example,  instead . Additio instead  are not ,Vibrio spp.Azospirillum spp.Acinetobacter spp.,,1AEP degradation, although in vivo studies will be required to fully establish this point.Overall, the oxidoreductases described here occur in all the major clades of PhnA and PhnX-possessing bacteria observed in earlier database searches, i.e., predominantly proteobacteria, with some representatives of the PVC group1AEP carried out by PbfC and PbfD enzymes generates methylamine as a product  can sustain marine microbes as a source of nitrogen even when phosphate levels are high. product  and in aVibrio spp. that possess pbfB  that will need to be further processed by other metabolic enzymes.The distribution of the four pathways among different bacteria may be an accident of evolution. Nevertheless, it should also be noted that certain pathways may suit better the lifestyles and metabolic setups of individual bacterial species. For example, a pathway based on PhnWYA requires three enzymes and is unable to process Min vitro, on recombinant proteins expressed in E.coli. It could be useful to complement our results with in vivo approaches, encompassing for example the deletion of the pbfB, pbfC, or pbfD genes in organisms that possess them. Such an approach might, however, be limited by several confounding factors and requires an understanding of the conditions under which the AEP degradation gene cluster is expressed in that organism. Furthermore, while our data strongly indicate that PbfB is involved in the processing of M1AEP, details about its exact reaction remain uncertain because we did not succeed in purifying the protein. In particular, the limits of assays conducted on whole cell extracts do not allow us to conclude whether PbfB behaves as an oxidase (like the PbfD enzymes) or as a dehydrogenase (like PbfC).The functional characterization described herein was conducted alessio.peracchi@unipr.it).Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Alessio Peracchi - and (R) enantiomers of HAEP were also prepared by methods described in the same publication.Racemic HAEP was synthesized starting from vinylphosphonic acid, as described in the literatureN-alkylated analogs of AEP and (R)-HAEP was accomplished by either newly developed or adapted literature methods, employing the following general procedures.The synthesis of the different 1H: 700.40 MHz, 13C: 176.12 MHz), AV III 600 , AV NEO 400  or AV NEO NanoBay 400 . Chemical shifts were referenced to solvent  peaks of CDCl3: δ = 7.26 ppm (1H NMR), δ = 77.00 ppm (13C NMR); CD3OD: δ = 3.31 ppm (1H NMR), δ = 49.00 ppm (13C NMR); HDO: δ = 4.79 ppm (1H NMR). 31P NMR spectra were referenced to external H3PO4 (85% aqueous solution), δ = 0.00 ppm. Chemical shifts (δ) are given in ppm and coupling constants (J) in Hz. Spectroscopic data for all synthesized compounds are provided in the Supporting Information. Mass spectra were recorded on a Bruker maXis UHR-TOF (instrument type: Qq-TOF), the samples were ionized by electrospray ionization (ESI). IR spectra were recorded on a Bruker Vertex 70 IR spectrometer in ATR mode. Optical rotations were measured at 20°C on a Perkin-Elmer 341 polarimeter in a 1 dm cell. [α]D values are given in 10-1 deg cm2 g-1. Melting points were determined on a Büchi Melting Point-system M-560 instrument or a Reichert-Jung Thermovar hot-stage microscope and are uncorrected. NMR spectra for all synthesized compounds, are provided in the NMR spectra were recorded on a Bruker BioSpin AV III HD 700 (254 glass plates (coating 0.25 mm thick) and spots were visualized by UV light (254 nm) and/or dipping the plate in a Cer-Ammonium-Molybdate (CAM) solution , a KMnO4 solution  or a ninhydrin solution [0.5% ninhydrin in ethanol (98%v/v)], followed by heating with a heat gun. Chromatographic purifications were either performed manually or carried out on a fully automated Biotage Isolera Prime flash purification system using KP-Sil separation cartridges 10-100 g , filled with Merck silica gel 60 (230-400 mesh).Thin layer chromatography (TLC) was carried out using Merck silica gel 60 FAll mentioned chemicals and solvents were bought from ABCR, Acros, Fluka, Sigma-Aldrich or TCI and were used without any further purification, unless stated differently.1) has been prepared by an adapted literature procedure.1, 1.152 g, 4.7 mmol, 84%) as a colorless liquid. 1H NMR  δ = 4.17-4.06 , 3.53 , 2.38 , 1.33 ; 13C NMR  δ = 62.02 , 30.79 , 23.83 , 16.41 ; 31P NMR (162.03 MHz) δ = 25.57 (s); HRMS calculated for [C6H14O3PBr+Na]+: 266.9762, found: 266.9759; IR (ATR) v = 2981, 2907, 1273, 1016, 955, 786, 509 cm-1.Related to 5-7) were obtained following the same two-step procedure from 1 as outlined below . For procedure A, either methyl amine (to obtain 2), ethyl amine (to obtain 3) or n-propyl amine (to obtain 4) was used as monoalkylamine component. Methylamine was used as aqueous solution (40% w/w in water), thus the total volume of added ethanol was reduced to 4.7 mL.All three monoalkylated phosphonic acids  in dry EtOH (10 mL), and the resulting mixture was stirred for 3 h at room temperature. The solvent was removed under reduced pressure to give diethyl 2-alkylamino-ethylphosphonates , in sufficient purity for the next step.The respective monoalkylamine (12 eq.) was added at 0°C to a solution of diethyl-(2-bromoethyl)phosphonate  δ = 4.14-4.03  2.85 , 2.42 , 1.96 , 1.94 , 1.31 ; 13C NMR  δ = 61.56 , 45.28 , 35.9  26.17 , 16.40 ; 31P NMR  δ = 30.49 (s); HRMS calculated for [C7H18NO3P+H]+: 190.1103, found: 190.1100; IR (ATR) v = 3312, 2980, 2794, 1230, 1021, 788, 538 cm-1.Related to 1H NMR  δ = 4.17-4.03 , 2.91 , 2.66 , 1.98 , 1.65 , 1.32 , 1.11 ; 13C NMR  δ = 61.59 , 43.73 , 43.15 , 26.46 , 16.44 , 15.13 ; 31P NMR (161.98 MHz) δ = 27.07 (s); HRMS calculated for [C8H20NO3P+H]+: 210.1254, found: 210.1267.Related to 1H NMR  δ = 4.15-4.03 , 2.89 , 2.56 , 1.97 , 1.64 , 1.49 , 1.31 , 0.91 ; 13C NMR  δ = 61.57 , 51.48 , 43.31 , 26.53 , 23.10 , 16.46 , 11.75 ; 31P NMR  δ = 27.53 (s); HRMS calculated for [C9H22NO3P+H]+: 224.1410, found: 224.1418.Related to 2-4) was dissolved in hydrochloric acid  and stirred at 120°C for 6 h. The crude product was concentrated in vacuo and purified by ion exchange chromatography (Dowex 50W × 8 / H+-form) using H2O as eluent. The product containing fractions (ninhydrin positive) were pooled and the solvent was removed under reduced pressure to yield  δ = 3.24 , 2.75 , 2.03-1.96 ; 13C NMR  δ = 44.85 , 32.46 , 24.90 ; 31P NMR  δ = 18.23 (s); HRMS calculated for the dimer [C6H20N2O6P2+H]+: 279.0869, found: 279.0854; anal. calc. for C3H10NO3P × 0.1 H2O: C 25.57%, H 7.30%, N 9.94%, O 35.20%, P 21.98%, found: C 25.82%, H 7.44%, N 9.91%, O 34.15%, P 21.79%; IR (ATR) v = 3102, 2694, 2385, 1681, 1586, 1133, 918, 443 cm-1.Related to 1H NMR  δ = 3.27-3.21 , 3.13 , 2.03-1.94 , 1.30 ; 13C NMR  δ = 42.82 , 42.56 , 25.09 , 10.50 ; 31P NMR  δ = 18.16 (s); HRMS calculated for [C4H12NO3P+Na]+: 176.0447, found: 176.0444; anal. calc. for C4H12NO3P: C 31.38%, H 7.90%, N 9.15%, O 31.35%, P 20.23%, found: C 31.15%, H 8.11%, N 9.21%, O 31.53%, P 19.03%; IR (ATR) v = 2993, 2857, 2733, 2433, 2338, 1711, 1139, 1020, 916 cm-1.Related to 1H NMR  δ = 3.28-3.28 , 3.04 , 2.03-1.95 , 1.49 , 0.99 ; 13C NMR  δ = 48.92 , 43.28 , 25.00 , 19.16 , 10.09 ; 31P NMR  δ = 18.26 (s); HRMS calculated for [C5H14NO3P+Na]+: 190.0604, found: 190.0601; anal. calc. for C5H14NO3P: C 35.93%, H 8.44%, N 8.38%, O 28.72%, P 18.53%, found: C 35.62%, H 8.61%, N 8.42%, O 29.09%, P 17.30%; IR (ATR) v = 2967, 2761, 2599, 2460, 2325, 1710, 1496, 1146, 1014 cm-1.Related to 8 was obtained from 1 and dimethyl amine (12 eq.) following procedure A. 1H NMR  δ = 4.13-4.04  2.73-2.65 , 2.32 , 2.04-1.97 , 1.30 ; 13C NMR  δ = 61.70 , 52.57 , 44.44  23.86 , 16.39 ; 31P NMR  δ = 29.45 (s); HRMS calculated for [C8H20NO3P+H]+: 210.1259, found: 210.1250; IR (ATR) v = 3471, 2979, 2820, 2768, 1234, 1022, 952 cm-1.Related to 9 was obtained from 8 following procedure B. m.p. 144-147°C; 1H NMR  δ = 3.38-3.31 , 2.92 , 2.09-1.99 ; 13C NMR  δ = 53.73 , 42.38 , 23.72 ; 31P NMR  δ = 17.64 (s); HRMS calculated for the dimer [C8H24N2O6P2+H]+: 307.1182, found: 307.1182; anal. calc. for C4H12NO3P × 0.25 H2O: C 35.93%, H 8.44%, N 8.38%, O 28.72%, P 18.53%, found: C 35.62%, H 8.61%, N 8.42%, O 29.09%, P 17.30%; IR (ATR) v = 3224, 2642, 2360, 1499, 1148, 1042, 903, 473 cm-1.Related to 8 was dissolved in ethyl acetate (10 mL), washed with an aqueous solution of sodium hydroxide , the organic phase was dried (MgSO4), and concentrated in vacuo. The resulting residue was used for the synthesis of diethyl 2--ethylphosphonate iodide  following a literature procedure1H NMR  δ = 4.26-4.17  3.65-3.59 , 3.17 , 2.58-2.49 , 1.35 ; 13C NMR  δ = 64.13 , 60.22 , 52.57 , 15.52 ; 31P NMR  δ = 27.24 ; HRMS calculated for [C9H23INO3P-I]+: 224.1410, found: 224.1410; anal. calc. for C9H23INO3P × 0.1 H2O: C 30.63%, H 6.63%, N 3.97%, O 14.05%, P 8.78%, found: C 30.47%, H 6.64%, N 4.16%, O 13.82%, P 8.44%; IR (ATR) v = 2980, 1739, 1480, 1240, 1020, 969, 821, 558 cm-1.Related to N,N,N-trimethylammonium-ethylphosphonate iodide  was dissolved in H2O (1 mL), conc. HCl  was added and the mixture was stirred at 110°C for 12 h. The solvent was removed under reduced pressure and the crude product was purified by ion exchange chromatography (Dowex Monosphere-550A / OH–-form). For this purpose, the product was applied to the ion exchange resin, then the column was washed with H2O (3 column volumes), then the product was eluted with an aqueous HCOOH solution (5% v/v). Finally the solvent was removed in vacuo to give 2-N,N,N-trimethylammonio-ethylphosphonic acid . The solid residue can be crystallized from EtOH to give colorless crystals. m.p. 248-250°C; 1H NMR  δ = 3.55-3.50 , 3.15 , 2.17-2.09 ; 13C NMR  δ = 62.77 , 52.50  22.73 ; 31P NMR  δ = 16.83 ; HRMS calculated for the dimer [C10H28N2O6P2+H]+: 335.1495, found: 335.1493; anal. calc. for C5H14NO3P × H2O: C 32.43%, H 8.71%, N 7.56%, O 34.56%, P 16.73%, found: C 32.15%, H 8.79%, N 7.65%, O 34.68%, P 16.08%; IR (ATR) v = 2980, 1739, 1480, 1240, 1020, 969, 821, 558 cm-1.Related to R)-1-Hydroxy-2-aminoethylphosphonic acidR)-13; 141 mg, 1.00 mmol, 1 eq.] was dissolved in formic acid , formaldehyde  was added and the mixture was refluxed for 18 h. The solvent was removed under reduced pressure and the crude residue was purified by ion exchange chromatography (Dowex 50W × 8 / H+-form) with H2O as eluent. The product containing fractions (ninhydrin positive) were pooled and the solvent was removed in vacuo to yield (R)- 1-hydroxy-2-(dimethylammonio)-ethylphosphonic acid  as colorless solid which can be recrystallized from H2O/EtOH. m.p. 257°C (decomp.); 1H NMR  δ = 4.16-4.08 , 3.44-3.35 , 3.00 , 3.00 ; 13C NMR  δ = 63.10 , 59.06 , 44.60 , 41.01 ; 31P NMR  δ = 14.31 (s); HRMS calculated for the dimer [C8H24N2O8P2+H]+: 339.1081, found: 339.1080; anal. calc. for C4H12NO4P × H2O: C 25.67%, H 7.54%, N 7.49%, O 42.75%, P 16.55%, found: C 25.51%, H 7.70%, N 7.40%, O 42.35%, P 16.03%; IR (ATR) v = 3165, 3006, 2619, 1458, 1085, 922, 560, 540 cm-1; c 1.09, H2O).Related to R)-13  was dissolved in MeOH (45 mL) and K2CO3  was added. After 10 minutes, methyl iodide  was added and stirring was continued for 48 h. The solvent was removed in vacuo and the residue was dissolved in H2O (3.4 mL). Subsequently, the pH was adjusted to 1 with conc. H2SO4 and the resulting mixture was stirred for 1 h. The solvent was removed under reduced pressure, the residue was dissolved in water and washed Et2O (3 × 4 mL). The aqueous phase was concentrated to dryness, dissolved in a minimal amount of water and applied to an ion exchange chromatography resin (Dowex Monosphere-550A / OH–-form). An aqueous solution of formic acid (5% v/v) was used as eluent. The product containing fractions were pooled and concentrated. The obtained amorphous solid can be recrystallized from EtOH/H2O to give (R)-1-hydroxy-2--ethylphosphonic acid  as colorless needles. m.p. 269-272°C; 1H NMR : δ = 4.34 , 3.65 , 3.58 , 3.24 ; 13C NMR : δ = 68.06 , 64.17 , 53.97 , 53.95 , 53.92  ppm; 31P NMR : δ = 14.05 ; HRMS calculated for [C5H14NO4P+H]+: 184.0739, found: 184.0730; anal. calc. for C5H14NO4P × 2.5 H2O: C 26.32%, H 8.39%, N 6.14%, O 45.57%, P 13.57%, found: C 26.37%, H 8.45%, N 6.06%, O 45.20%, P 13.44%; IR (ATR): v = 3160, 2337, 1599, 1478, 1248, 1084, 967, 914 cm-1; c 0.91, D2O).Related to R)-diisopropyl -1-hydroxyethyl)phosphonateR)-12, 2.03 mmol, 721 mg, 1eq.] in ethanol (10 mL) and the mixture was stirred at 80°C for 2 h. The reaction mixture was filtered, concentrated in vacuo and the resulting residue was purified by column chromatography  giving (R)-diisopropyl 2-amino-1-hydroxyethyl-phosphonate  as a colorless oil. 1H NMR  δ = 4.78-4.69 , 3.80 , 2.95 , 2.84 ; 1.36-1.33 ; 13C NMR  δ = 73.15 , 72.98 , 69.74 , 43.79 , 24.38 , 24.36 , 24.23 ; 31P NMR  δ = 21.84 (s); HRMS calculated for [C8H20NO4P+H]+: 226.1208, found: 226.1203; IR (ATR) v = 3286, 2979, 2934, 1467, 1375, 1212, 975, 567 cm-1.Related to R)-14  was dissolved in dry THF (5 mL), then triethylamine  and tosyl chloride  were added dropwise in a row at 0°C. After stirring for 20 h at room temperature, CH2Cl2 (5 mL) was added and the mixture was washed with brine. The layers were separated and the aqueous phase was extracted with CH2Cl2 (3 × 5 mL). The combined organic layers were dried (MgSO4), filtered and concentrated in vacuo. The crude product was purified by MPLC , to give (R)-diisopropyl 1-hydroxy-2-(tosylamido)-ethylphosphonate  as a colorless, crystalline solid; m.p. 118-119°C. 1H NMR  δ = 7.75 , 7.31 , 5.43  4.75-4.66  3.85 , 3.71 , 3.35 , 3,14 , 2.42 , 1.62 , 1.31 , 1.31  , 1.30  , 1.26 ; 13C NMR  δ = 143.57 , 136.73 , 129.76 , 127.15 , 72.10 , 72.06 , 66.71 , 44.47  24.07 , 23.88 , 23.81 , 21.5 ; 31P NMR  δ = 19.75 (s); HRMS calculated for [C15H26NO6PS+Na]+: 402,1116, found: 402,1108; anal. calc. for C15H26NO6PS: C 47.49%, H 6.91%, N 8.45%, O 25.30%, P 8.16%, S 3.69%, found: C 47.38%, H 6.87%, N 8.28%, P 7.95%, S 3.68%; IR (ATR) v = 3470, 3433, 3092, 2983, 2904, 1738, 981 cm-1; c 0.60, acetone).Related to R)-15  in dry DMF (1 mL). After stirring at room temperature for 18 h, the reaction mixture was quenched by addition of H2O (2mL). The organic phase was separated, and the aqueous phase was extracted with Et2O (3 × 3 mL). The combined organic layers were dried (MgSO4), filtered and concentrated in vacuo. The residue was purified by MPLC , giving (R)-diisopropyl 1-hydroxy-2-(N-methyl-N-tosyl-amido)-ethylphosphonate  as a colorless solid; m.p. 110-111°C; 1H NMR  δ = 7.68 , 7.33 , 4.77  4.06 , 3.38-3.27 , 2.88 , 2.79 , 2.44 , 1.37-1.34 ; 13C NMR  δ = 143.71 , 134.45 , 129.82 , 127.43 , 71.95 , 71.74 , 67.25 , 52.00 , 36.73 , 24.13 , 24.00 , 23.97 , 21.54 ; 31P NMR  δ = 19.34 (s); IR (ATR) v = 3267, 2985, 2922, 2853, 1278, 945, 525 cm-1; c 1.04, acetone). Ee-determination was accomplished by 31P NMR spectroscopy in the presence of a chiral solvating agent ((+)-(R)-tert-butylphenylphosphinothioic acid),3): δ = 97.17 , 19.49 , 19.67  ≙ ee ≥ 99%.Related to R)-16  was allowed to react in a manner similar to that described in general procedure B to give (R)-1-hydroxy-2-(methylammonio)ethylphosphonic acid . m.p.. 261-264°C; 1H NMR  δ = 4.02 , 3.36 , 3.24 , 2.79 ; 13C NMR  δ = 64.39 , 50.72 , 32.77 ; 31P NMR  δ = 14.59 (s); HRMS calculated for the dimer [C6H21N2O8P2+H]+: 311.0768, found: 311.0766; anal. calc. for C3H10NO4P: C 23.23%, H 6.50%, N 9.03%, O 41.26%, P 19.97%, found: C 23.02%, H 6.52%, N 8.82%, O 41.20%, P 20.05%; IR (ATR) v = 3194, 2994, 2702, 2521, 2460, 1632, 956, 482 cm-1; c 0.84, H2O).Related to Vibrio vulnificus , Azospirillum sp. B510  and Acinetobacter baumannii .An initial visual inspection of bacterial AEP degradation gene clusters was performed using tools publicly available at the Department of Energy Integrated Microbial Genomes (IMG) databasei.e. AEP degradation gene clusters) were considered for further analysis.Homologs of the three oxidoreductases were identified using a BLASTp searchThe phylogenetic relationships among PbfB, PbfC and PbfD, as well as between these enzymes and other functionally characterized oxidoreductases sharing a PF01266 domain, were investigated as follows.Based on genomic searches, we assembled an initial set encompassing all the sequences containing a PF01266 domain that we identified within AEP degradation gene clusters (260 sequences). This initial set was reduced to 64 representative proteins, primarily by removing all sequences showing a pairwise identity higher than 90% with CD-HIT.N-methyl aminobutyrate oxidases (MabO); (iv) two glycine oxidases (ThiO); (v) two γ-glutamylputrescine oxidases (PuuB); (vi) two N-methyl-L-tryptophan oxidases (SolA); (vii) two mitochondrial sarcosine dehydrogenases (SarDH); (viii) two mitochondrial dimethylglycine dehydrogenases (DMGDH). The accession IDs of these additional sequences are provided in A second, larger dataset (dataset B) comprising a total of 80 sequences was subsequently prepared by adding a custom selection of oxidoreductases of known function, which shared with PbfA, PbfB and PbfC the presence of the PF01266 domain. In addition to the 64 sequences from dataset A, dataset B contained: (i) two monomeric sarcosine oxidases (SoxA); (ii) two D-amino acid dehydrogenases (DadA); (iii) two γ-ModelFinderThe synthetic codon-optimized genes of the FAD-dependent enzymes PbfB (GenBank: WP_049798008), PbfC (GenBank: WP_012976454), PbfD1 (GenBank: WP_079548425) and PbfD2 (GenBank: WP_075082418) were purchased from Proteogenix  or from BaseGene BV  and cloned into the NdeI/NotI restriction sites of a pET28a-N-His-tag vector.Azospirillum was overexpressed in E. coli Tuner (DE3) cells. A 10-mL bacterial preculture grown overnight at 37°C was used to inoculate 1L of Luria-Bertani broth (LB) containing kanamycin (50 μg/mL) until OD600 reached 0.7-0.8. The temperature was then lowered to 20°C and protein expression was induced by adding 0.25 mM isopropyl-β-D-1-thiogalactopyranoside (IPTG). 20 hours after induction, the cells were harvested by centrifugation. The cell pellets were washed once with phosphate-buffered saline solution (PBS), resuspended, centrifuged again, and finally stored at −20°C.Related to buffer A , supplemented with 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM benzamidine, 10% glycerol and 10 μM FAD. The cell suspension was sonicated and centrifuged .For protein purification, pellets were resuspended in After sonication, the cell lysate was cleared by centrifugation . The supernatant was recovered and continuously looped (flow rate: 1 mL/min) on a 5 mL HisTrap™ Fast Flow column (Cytiva) for 2 hours at 4°C through a peristaltic pump. Afterwards, the loaded column was mounted onto an AKTA Pure System FPLC apparatus and proteins were eluted with a gradient of buffer A supplemented with 208 mM imidazole. Fractions containing PbfC were pooled and extensively dialyzed against buffer A, supplemented with 10% glycerol and 10 μM FAD, then concentrated by an Amicon® Ultra-2 3K device (Merck) to 3 mL. The protein was further loaded into a 5 mL loop of AKTA Pure System FPLC and purified by size exclusion chromatography (SEC) with a HiPrep™ 16/60 Sephacryl S-300 HR (Cytiva) column, equilibrated with the same buffer as above (buffer A plus 10% glycerol and 10 μM FAD). The protein was finally concentrated by Amicon centrifugation and subdivided into aliquots (about 0.5 mL each) to be stored at −80°C. Protein purity was assessed higher than 95% by SDS-PAGE and the final yield was ≈3 mg of purified PbfC per liter of bacterial culture. A SDS-PAGE of purified PbfC (side by side with purified PbfD1 and PbfD2) is shown in Acinetobacter baumannii was expressed in E. coli Tuner (DE3) cells (EMD Biosciences), which were grown in self-inducing medium (LB containing glucose 0.5 g/L and lactose 2 g/L). A 10-mL bacterial preculture grown overnight at 37°C was used to inoculate 1L of self-inducing medium containing kanamycin (50 μg/mL). The culture was kept under agitation at 20°C for up to 24 h, then the cells were harvested by centrifuging at 7,200 × g for 10 min at 4°C.Related to buffer B  and centrifuged again. The supernatant was discarded, while the pellets were stored at −20°C.The pellet was resuspended in For protein purification, the pelleted cells were thawed on ice and resuspended in buffer B supplemented with 1 mM PMSF, 1 mM benzamidine, 10% glycerol and 10 μM FAD. Lysozyme  was added to the cell suspension, which was then incubated at 4°C under agitation for 30 minutes and finally sonicated. At the end of sonication, the lysate was centrifuged at 26,200 × g for 40 minutes (at 4°C) and the supernatant was used for the next purification step.PbfD1 purification exploited a 5 mL HisTrap™ Fast Flow column (Cytiva). The cleared lysate was continuously looped (flow rate: 2 mL/min) on the column for 2 hours at 4°C, after which the column was transferred to an AKTA Pure System FPLC apparatus.The protein was eluted with a gradient of buffer B supplemented with 250 mM imidazole. PbfD1 fractions were pooled and extensively dialyzed against a storage buffer . The protein was subdivided into aliquots (about 0.5 mL each) to be stored at −80°C. Protein purity was assessed higher than 95% by SDS-PAGE and the final yield was ≈100 mg of purified PbfD1 per liter of bacterial culture.E. coli Chaperone Competent pGro7/BL21 cells (Takara) and exploiting D-sorbitol as a carbon source in the growth medium. This reportedly promotes the activation of an osmotic stress response helpful to increase protein solubility.Related to A 10-mL bacterial preculture grown at 37°C overnight was used to inoculate 1 L of self-inducing medium (LB containing glucose 0.5 g/L and lactose 2 g/L) supplemented with kanamycin (50 μg/mL), chloramphenicol (20 μg/mL), arabinose  and D-sorbitol (5 mg/mL). The culture was kept under agitation at 20°C for up to 24 h, then the cells were harvested by centrifugation at 7,200 × g for 10 min (4°C). After removing the supernatant, the pellet was resuspended in buffer B, then centrifuged again, discarding the supernatant. The pellets were then stored at −20°C waiting to proceed with lysis and purification.For protein purification, pellets were thawed and resuspended in buffer B, supplemented with 1 mM PMSF, 1 mM benzamidine, 10% glycerol, 0.05% Triton® X-100 and 10 μM FAD. After adding lysozyme 1 mg/mL, the cell suspension was incubated in a cold room under agitation for 30 minutes.2 and thawed quickly at 37°C in a thermostated water bath. The freezing-thawing procedure was repeated two more times, after which the suspension was sonicated briefly. At the end of sonication, the lysate was centrifuged at the maximum speed ; the pellet was discarded and the supernatant was retained for further processing.The suspension was frozen in liquid NPbfD2 purification exploited also in this case a 5 mL HisTrap™ Fast Flow column (Cytiva). The lysate was continuously looped (flow rate: 2 mL/min) on the column for 2 hours at 4°C, after which the column was transferred to the FPLC apparatus.2, 5 mM ATP) to displace the chaperonins bound to nickel resin; subsequently, PbfD2 was eluted with a gradient of buffer B supplemented with 250 mM imidazole. PbfD2 fractions were pooled and concentrated by an Amicon® Ultra-2 3K device (Merck), then extensively dialyzed against a buffer containing 50 mM TEA pH 7.5, 200 mM NaCl and 10% glycerol. The protein was subdivided into aliquots (about 0.5 mL each) to be stored at −80°C. Protein purity was assessed higher than 80% by SDS-PAGE and the final yield was ≈2.5 mg of purified PbfD2 per liter of bacterial culture.The column was first washed with 30 mL of chaperone-removal buffer  or with a thermostated V-750 (Jasco Inc.) UV-Vis spectrophotometer and corrected for buffer contribution.450nm = 11300 M−1cm−1 as described.max) were: ε462nm=10,650 M-1 cm-1 for PbfC (Azospirillum sp. B510); ε447=13,340 M-1 cm-1 for PbfD1 (A. baumannii) and ε456=11,950 M-1 cm-1 for PbfD2 (M. fucicola). In subsequent experiments, enzyme concentrations were calculated based on these coefficients.Denaturation of the proteins with sodium dodecyl sulfate, aimed at identifying the type of cofactor  and at calculating the extinction coefficients of protein-bound flavin, was carried out as described by Aliverti et al.The oxidoreductase activity of the recombinant enzymes was first assessed through two assays conducted in 96-well microtiter format. These colorimetric assays served to rapidly screen the ability of the enzymes to oxidize a number of potential substrates (both phosphonates and non-phosphonates), transferring the electrons to either an ‘artificial’ electron acceptor or to molecular oxygen.In the first assay, the redox dye 2,6-dichlorophenol indophenol (DCPIP) was the final electron acceptor, whereas phenazine methylsulfate (PMS) served as an electron mediator.o-dianisidine.o-dianisidine . After another 5 min, 3.5 M sulfuric acid was added to each well  and the plate was photographed.A single-point plate assay was used to screen for the ability of the recombinant enzymes to use molecular oxygen as the electron acceptor. The plate assay detected the release of hydrogen peroxide, which was revealed by exploiting horseradish peroxidase and the chromogenic substrate E. coli cells in phosphate-free buffer (50 mM sodium 4-(2-hydroxyethyl)-1-piperazineethanesulfonate, pH 7.5).Phosphate release was also determined in a microtiter assay, by employing the BIOMOL® Green kit according to the manufacturer’s instructions. Enzymes that had been stored in phosphate or pyrophosphate buffer were dialyzed against a different buffer  before using them in these assays. Likewise, cell extracts to be subjected to this assay were obtained by lysing the E. coli cells expressing the recombinant enzyme), 4 μM PhnX, 2 mM MgCl2 and the substrate. After a one-hour incubation at room temperature, 18 μL of the reaction mixture (or 36 μL in the case of cell extracts) was transferred into 200 μL of BIOMOL® Green reagent to block the enzyme activity. Color development was assessed after another 30 min, by measuring the absorbance at 600 nm using a Cary 50 UV-Vis spectrophotometer (Varian).A typical reaction contained 50 mM TEA-HCl buffer pH 8.0, the oxidoreductase under examination , we utilized GDH to convert α-ketoglutarate to L-glutamate by using ammonia and oxidizing NADH, whose disappearance can be monitored spectroscopically at 340 nm.For this assay, the oxidoreductase under examination  was incubated in a solution containing 50 mM TEA-HCl buffer pH 8.0, 2 mM AEP and 1 mM α-ketoglutarate. After 30 min, the mixture was supplemented with NADH  and finally with GDH . The disappearance of NADH absorption at 340 nm was monitored by a UV-Vis Cary 50 spectrophotometer. In control experiments where either the oxidoreductase or AEP was omitted, no significant NADH oxidation was observed.1H NMR spectra were recorded with a Bruker Avance-III 400 spectrometer equipped with a CRPN2-DR-BB/1H&19F-5mm-EZ PRODIGY 400 CRYOPROBE in non-spinning mode at room temperature using a gradient-based solvent suppression pulse sequence (1D Excitation Sculpting using 180º water-selective pulses “zgesgp”) for the water signal suppression .The reaction mixture  contained 30 mM sodium phosphate buffer pH 8.0, 2 mM MgCl2, 5 mM substrate, 0.5 μM enzyme, 10% D2O and 1 mM of deuterated trimethylsilyl propionic acid, used as internal chemical shift reference (δ=0.00 ppm). NMR spectra were processed and analyzed with MestReNova v12.0.4.1AEP and M2AEP (for PbfD1 only) with oxygen as the electron acceptor was monitored by coupling the production of PAA to that of acetaldehyde catalyzed by PhnX and to the oxidation of NADH by alcohol dehydrogenase (ADH).,The oxidation of AEP, M2, PhnX (4 μM) and ADH (6 U), in addition to the enzyme under examination and the substrate . The disappearance of NADH was monitored spectrophotometrically  on a Cary 50 UV-Vis spectrophotometer.The reactions were conducted at 25°C in a final volume of 120 μl, containing 50 mM TEA-HCl buffer pH 8.0, 0.25 mM NADH, 2.5 mM MgClTo obtain the kinetic parameters of the oxidases, reactions at different substrate concentrations were repeated in triplicate and the data were nonlinearly least-squares fitted to the Michaelis-Menten equation using Sigma Plot.1AEP catalyzed by PbfC was typically measured through a continuous kinetic assay based on DCPIP and PMS, similar to the assay reported by Augustin and coworkers.-1 cm-1) had been previously established under the same assay conditions. Each reaction was repeated in triplicate and the kinetic parameters were computed by nonlinear least-squares fitting of the data to the Michaelis-Menten equation using Sigma Plot.Since PbfC was very inefficient at utilizing molecular oxygen as the electron acceptor, the oxidative deamination of AEP and MMaximum likelihood phylogenetic trees shown in Kinetic parameters for the recombinant enzymes towards given substrates were obtained by nonlinear least-squares fitting of the experimental data  to the Michaelis-Menten equation using SigmaPlot 14.0 . Data were collected in triplicate and the resulting kinetic parameters are reported in


Fungi are prolific producers of bioactive small molecules of pharmaceutical or agricultural interest. The secondary metabolism of higher fungi (Dikarya) has been well-investigated which led to > 39,000 described compounds. However, natural product researchers scarcely drew attention to early-diverging fungi (Mucoro- and Zoopagomycota) as they are considered to rarely produce secondary metabolites. Indeed, only 15 compounds have as yet been isolated from the entire phylum of the Zoopagomycota.Phytophthora sp. Lindolin biosynthesis was reconstituted in vitro and relies on the activity of two enzymes of dissimilar evolutionary origin: Whilst the IAA–CoA ligase LinA has evolved from fungal 4-coumaryl-CoA synthetases, the subsequently acting IAA-CoA:anthranilate N-indole-3-acetyltransferase LinB is a unique enzyme across all kingdoms of life.Here, we showcase eight species of the order Kickxellales (phylum Zoopagomycota) as potent producers of the indole-3-acetic acid (IAA)-derived compounds lindolins A and B. The compounds are produced both under laboratory conditions and in the natural soil habitat suggesting a specialized ecological function. Indeed, lindolin A is a selective agent against plant-pathogenic oomycetes such as This is the first report on bioactive secondary metabolites in the subphylum Kickxellomycotina and the first evidence for a non-clustered, two-step biosynthetic route of secondary metabolites in early-diverging fungi. Thus, the generally accepted “gene cluster hypothesis” for natural products needs to be reconsidered for early diverging fungi.The online version contains supplementary material available at 10.1186/s40694-023-00166-x. Mucor, Rhizopus, Phycomyces and Mortierella encompassing plant symbionts and potent producers of industrially relevant polyunsaturated fatty acids and β-carotene-derived pigments − , respectively, dominated in the culture filtrate extracts , as an amide of indole-3-acetic acid (IAA) and anthranilic acid  was assigned as a derivative of 4 with a 5ʹ-hydroxy group at the anthranilic moiety  and minor amounts of the 3-related compounds indole lactic acid  and indole ethanol  in the extracts as determined by GC–MS/MS using a trimethylsilyl derivatization procedure of commercially available or synthesized reference standards   showed the highest similarity (76% pairwise identity) and lowest bit-score   . Hereby,e A Fig. B. We sealinA was heterologously expressed in Escherichia coli, and the enzyme was purified as a C-terminal His6-tagged soluble fusion protein. We investigated the substrate specificity of LinA by measuring the released AMP by monitoring the depletion of NADH with a well-established enzyme-coupled assay  for 7–14 days at 25 °C (Coemansia spp.) or 30 °C (Linderina sp. and Martensiomyces sp.). The mycoparasite Dimargaris bacillispora CBS218.59 was grown on its fungal host Cokeromyces recurvatus on V8 agar  at 25 °C for 14 days. To induce lindolin biosynthesis, 100 mL meat medium , MEP medium, or PDB [26.5 g L−1 potato dextrose broth (Carl Roth)] were inoculated with three agar blocks (9 mm2) of the respective strains and incubated at 160 rpm at 25 °C or 30 °C for 3 to 21 days. Cultures were additionally grown on meat medium, MEP and PDB agar plates containing 20 g L−1 agar for 7–28 days at 25 °C or 30 °C. To show l-tryptophan-dependent production of 3–5, L. pennispora was cultivated on Aspergillus minimal medium (AMM) [ d-glucose but with 1% (w/v) casamino acids (Difco) and with or without the supplementation of 5 mM l-tryptophan, for three days. L. pennispora, M. pterosporus and C. furcata were cultivated emersly on 20 g autoclaved potting soil  supplemented with or without 1% (w/w) d-glucose for ten days. Oomycete strains (P. megasperma and P. macrosporum) were verified by ITS sequencing [F. graminearum was cultivated on MEP agar at 30 °C for 4 days. Escherichia coli XL1 blue was used for plasmid propagation and E. coli SoluBL21 (for LinA) and BL21 (for LinB) were used for protein production  or 50 µg mL−1 kanamycin (Roth), if required.All fungal EDF strains were initially verified by ITS sequencing  (Additioum (AMM)  without g, 10 min). 5 µL thereof were used for GC–MS and UHPLC measurements . The mycelium was rinsed with water, lyophilized to dryness, weighed to determine the fungal dry biomass and extracted with 5 mL methanol per 100 mg dry fungal biomass. The culture filtrates were adjusted to pH 6 by addition of 3 M HCl or 3 M NaOH prior to extraction with 100 mL ethyl acetate. Cultivated agar plates (25 mL) were sliced in 5 × 5 mm pieces and extracted overnight with 25 mL ethyl acetate. 20 g of soil samples were extracted with 40 mL of ethyl acetate. In all extraction procedures, uninoculated liquid medium, agar or soil served as negative control. The organic phase was collected and evaporated to dryness using a rotary evaporator. The residue was dissolved in 2 mL methanol, ultrasonicated for 1 min, and centrifuged to remove contaminating particles , IOL (2) and IAA (3) were determined by GC–MS on a Trace 1310 gas chromatograph (Thermo Fisher Scientific) coupled with a TSQ 9000 electron impact (EI)-triple quadrupole mass spectrometer using method 1  and split injection (split ratio 1:10). Total ion current (TIC) values were recorded in the mass range of 45–500 amu, with a scan time of 0.2 s. and a MS delay of 4 min. As 1 and 3 were too hydrophilic for optimal GC runs, silylation of 1–3 was conducted with N-methyl-N-(trimethylsilyl) trifluoroacetamide (MSTFA) prior to GC analysis: Methanolic fungal extracts were dried in vacuo and resuspended in 150 µL MSTFA. Samples were mixed for 15 s, ultrasonicated for 1 min and centrifuged. A volume of 1 µL of the supernatant was injected. The National Institute of Standards and Technology (NIST) Mass Spectra Search Program version 2.4 was used for comparison of the EI-MS spectra.The compounds ILA  casamino acids and 5 mM l-tryptophan with orbital shaking at 160 rpm at 30 °C for 10 days. Metabolites were extracted from the supernatant as described above. The residue was dissolved in 130 mL dichloromethane and reduced to 40 mL with a rotary evaporator. Purification of 4 and 5 was conducted using a Büchi C-810 Flash Chromatograph and an Agilent 1200 Infinity HPLC system equipped with a diode array detector (DAD). The extract was subjected to a normal phase column  on the flash chromatograph using method 3  was evaporated to dryness and dissolved in 3 mL methanol. In four individual runs, 750 µL thereof were flash-chromatographed using a reverse phase column  running method 4  was evaporated to dryness and dissolved in 3.5 mL methanol. Purification was accomplished on a reverse phase column  using method 6 . NMR spectra were recorded on a Bruker Avance III 600 MHz spectrometer at 300 K. DMSO-d6 served as solvent and internal standard (δH = 2.50 ppm and δC = 39.52 ppm).Fraction III (containing 2) was synthesized by reduction of IAA (3) with lithium aluminium hydride in THF according to Du et al. [3) with coenzyme A trilithium salt (Merck) according to Pourmasoumi et al. [Indole-3-ethanol  and B (5) (7.8–500 µM) for 4–6 days measuring the colony diameter. Pure solvent (methanol) or serial dilutions of hygromycin B (10–400 µM) served as negative and positive controls, respectively. Antiproliferative or cytotoxic activities were determined for human umbilical vein endothelial cells (HUVEC) or cervical cancer cells (HeLa), respectively, as described [4 and 5 was calculated on radish seedlings (Kiepenkerl) according to a published protocol [4 and 5 (growth-inhibitory effect). Solvent (methanol) and 3 (1 µM) were negative and positive controls, respectively. A competitive assay using 1 µM 3 and 1 µM 4 or 5 was carried out similarly (auxin-inhibitory effect). Root and hypocotyl growth were measured daily for a period of 5 days.Antimicrobial activities were initially determined at a concentration of 1 mg mLprotocol : Bacilluprotocol , 113. Inprotocol  suppleme−1. Genomic DNA was isolated as described [20 oligonucelotides. Expression analysis was performed at the AnalytikJena qTower3 using the qPCR Mix EvaGreen (Bio&SELL) and oligonucleotides with a minimum primer efficiency of 95% , the TEF transcription factor  and the glyceraldehyde-3-phosphate dehydrogenase  served as internal standards. Gene expression levels were determined as described by Pfaffl [Fungal mycelium from 36 h cultures in AMM, PDB and meat medium was lysed with glass beads (1–5 mm) in a FastPrep Homogenizer (MP Bio) for 2 min at 4.5 m sescribed . RNA wasy Pfaffl .linA and linB were amplified from L. pennispora cDNA using Phusion High-Fidelity DNA polymerase (NEB) and the oligonucleotides listed in Additional file NcoI/HindIII and NheI/EcoRI  and BL21 (linB), cultivated in 400 mL LB medium supplemented with 50 µg mL−1 kanamycin at 37 °C until an optical density of 0.6 was reached. Then, expression was initiated by adding 1 mM IPTG. After another 16 h of incubation at 16 °C, cells were harvested by centrifugation  and disrupted in 5 mL lysis buffer  by ultrasonication. The enzymes were purified using the Protino Ni2+ NTA Agarose (Macharey & Nagel) according to the manufacturer’s protocol by a step gradient (10–250 mM imidazole). The proteins were subsequently concentrated and re-buffered in assay buffer  using the Amicon Ultra-15 Central Filter System (30 kDa cut-off). The protein concentration was determined by the Pierce BCA protein Assay (Thermo Fisher Scientific) using albumin as reference standard. The purity was additionally verified by SDS polyacrylamide gel electrophoresis , 1 U mL−1 pyruvate kinase (Merck), 3 U mL−1 lactate dehydrogenase (Merck) and 1 µM LinA. The reaction was initiated with 1 mM of substrates  or 5-hydroxyanthranilate (HAA) (Merck). Water served as negative control. After 2 h of incubation at 25 °C, the reaction was stopped by shock freezing in liquid nitrogen and lyophilization. The residue was dissolved in an equal volume acetonitrile: water (50: 50), briefly centrifuged  and immediately subjected to UHPLC analysis using method 2  contained 200 mM Tris (pH 7.8), 20 mM MgCl4 and 5 in vitro, the following assay was set up in a 500 µL scale: 200 mM Tris (pH 7.8), 20 mM MgCl2, 250 µM CoA, 2.5 mM ATP, 1 µM LinA, 1 µM LinB, and 1 mM AA and/or HAA. As controls, individual assay components were omitted. The reaction was initiated by addition of 1 mM IAA (3). The assay was incubated for 120 min at 25 °C. Lindolin detection was carried out by UHPLC-MS measurements as described above.To produce linA and linB gene loci (± 20 kb) extracted from the genomes of eight verified lindolin-producing Kickxellales species. The minimum alignment sequence identity was set to 30%.Comparative gene cluster analysis was performed with CAGECAT , an onliL. pennispora LinA comprises 553 aa. Moreover, biochemically verified aryl-CoA ligases were additionally included [LinA-related enzymes were identified using the BLAST search tool  screening the non-redundant protein sequences (nr) of NCBI for each fungal order/phylum and other domains of life separately. For each order or phylum, ten (if available) most-likely hits (according to bit score) were depicted for further analysis. Sequences with less than < 400 or > 700 amino acids were omitted as included , 53, 57.included   screening the non-redundant protein sequences (nr) of NCBI for each fungal order/phylum and other domains of life separately. For each order or phylum, 100 (if available) most-likely hits (according to bit score) were depicted for further analysis. Moreover, biochemically verified aryl-transferases were additionally included , 58, 120Additional file 1: Table S1. Microbial strains.Additional file 2: Figure S1. Atom numbering and selected COSY (bold lines) and HMBC (red arrows) in 4.Additional file 3: Figure S2.1H NMR spectrum of 4 in DMSO-d6.Additional file 4: Figure S3.13C NMR spectrum of 4 in DMSO-d6.Additional file 5: Figure S4. DEPT-135 NMR spectrum of 4 in DMSO-d6.Additional file 6: Figure S5.1H-1H COSY spectrum of 4 in DMSO-d6.Additional file 7: Figure S6.1H-13C HSQC spectrum of 4 in DMSO-d6.Additional file 8: Figure S7.1H-13C HMBC spectrum of 4 in DMSO-d6.Additional file 9: Figure S8.1H-1H TOCSY spectrum of 4 in DMSO-d6.Additional file 10: Figure S9. ESI-MS/MS spectrum of 4.Additional file 11: Table S2. NMR data of 4 in DMSO-d6.Additional file 12: Figure S10. Atom numbering and selected COSY (bold lines) and HMBC (red arrows) in 5.Additional file 13: Figure S11.1H NMR spectrum of 5 in DMSO-d6.Additional file 14: Figure S12.13C NMR spectrum of 5 in DMSO-d6.Additional file 15: Figure S13. DEPT-135 NMR spectrum of 5 in DMSO-d6.Additional file 16: Figure S14.1H-1H COSY spectrum of 5 in DMSO-d6.Additional file 17: Figure S15.1H-13C HSQC spectrum of 5 in DMSO-d6.Additional file 18: Figure S16.1H-13C HMBC spectrum of 5 in DMSO-d6.Additional file 19: Figure S17.1H-1H TOCSY spectrum of 5 in DMSO-d6.Additional file 20: Figure S18. ESI-MS/MS spectrum of 5.Additional file 21: Table S3. NMR data of 5 in DMSO-d6.Additional file 22: Figure S19. GC-MS/MS spectrum of ILA (1).Additional file 23: Figure S20. GC-MS/MS spectrum of IOL (2).Additional file 24: Figure S21. GC-MS/MS spectrum of IAA (3).Additional file 25: Figure S22. Production of lindolin A and B in Kickxellales and related species.Additional file 26: Figure S23. Antimicrobial activities of 4 and 5.Additional file 27: Figure S24. Antiproliferative and cytotoxic activities of 4 and 5 against mammalian cells.Additional file 28: Figure S25. Plant growth activating and IAA-interfering properties of IAA (3), lindolin A (4) and lindolin B (5) on radish seedlings.Additional file 29: Figure S26. Initial screening for antifungal and anti-oomycete properties of lindolin A (4).Additional file 30: Figure S27. Determination of the minimal inhibitory concentration (MIC50) of lindolin A (4) on oomycetes.Additional file 31: Table S4. BLAST search for homologous genes required for IAA production in Kickxellales.Additional file 32: Figure S28. Quantification of IAA (3), lindolin A (4) and lindolin (5) in cultures of L. pennispora.Additional file 33: Figure S29. SDS polyacrylamid gel electrophoresis (SDS-PAGE) of purified His6-tagged LinA and LinB.Additional file 34: Figure S30. Determination of the optimal reaction conditions for LinA.Additional file 35: Figure S31. Activity of LinB during competitive substrate supply.Additional file 36: Figure S32. Determination of the optimal reaction conditions for LinB.Additional file 37: Figure S33. Clinker comparison between linA gene loci in Kickxellales.Additional file 38: Figure S34. Clinker comparison between linB gene loci in Kickxellales.Additional file 39: Table S8. BlastP analysis of LinA-like proteins in species of various kingdoms of life.Additional file 40: Table S9. BlastP analysis of LinB-like proteins in species of various kingdoms of life.Additional file 41: Figure S35. Alignment lengths of LinB-like proteins in SSN.Additional file 42: Figure S36. Percent identity depending on the alignment score of LinB-like proteins in SSN.Additional file 43: Figure S37. Histogram of sequence lengths of LinB-like proteins used for SSN analysis.Additional file 44: Table S5. Chromatographic Methods.Additional file 45: Table S6. Oligonucleotides used in this study.Additional file 46: Table S7. Plasmids used in this study.


In addition, Cu12a-π can be transformed into cluster-based nonporous adaptive crystals (Cu12b-NACs) via ligand-exchange following a dissociation-reassembly mechanism. Moreover, Cu12a-π can efficiently remove 97.2% of iodine from saturated iodine aqueous solutions with a high uptake capacity of 2.96 g·g−1. These prospective results positioned at cluster-based porous supramolecular framework and enlighten follow-up researchers to design and synthesize such materials with better performance.The assembly of cluster-based π-stacked porous supramolecular frameworks presents daunting challenges, including the design of suitable cluster building units, control of the sufficient C-H···π interactions, trade-off between structural dynamics and stability as well as understanding the resulting collective properties. Herein, we report a cluster-based C-H···π interaction-stacked porous supramolecular framework, namely, Cu12a-π, consisting of Cu The assembly of cluster-based π-stacked porous supramolecular frameworks presents daunting challenges. Here, the authors report a dynamic and transformable supramolecular framework based on Cu12 cluster and C-H···π-stacking with permanent porosity Recently, a series of crystalline porous materials (CPMs) were assembled through intermolecular weak non-covalent interactions, which involve hydrogen bond10, π···π or C-H···π stacking16, electrostatic interactions, van der Waals forces18, etc. Wherein, π-stacked organic frameworks (πOFs), one of the newly emerging CPMs, have greatly broadened the boundaries of porous science, but few examples associated to such CPMs have been reported. Moreover, most of the reported πOFs are stabilized not merely by pure π···π stacking interactions, whereas other forces are also involved to a greater or lesser extent, e.g. C-H···π stacking interactions12. As a result, it is hard to figure out how much the C-H···π stacking interactions contribute the stability of CPMs. Therefore, the construction of supramolecular networks with permanent pores stabilized by C-H···π stacking interactions is meaningful to investigate the performance of such a non-covalent interaction in CPMs.Inspired by fascinating molecular pores that nature has evolved, scientists have made great progress in constructing porous materials21. The rational construction of such CPMs may require the following two main factors: 1) the robust building unit; 2) the medium strength of the bridge linker, where the frame structure can balance the stability and flexibility. However, the most reported πOFs or HOFs (hydrogen-bonded organic frameworks) are constructed by organic building units, which may limit the stability on the process of dynamically structural transformation under drastic physical stimulation23. In addition, the further post-transformation of such OFs must be done through the redesign of the building unit. Therefore, the development of suitable building unit to rationally construct the C-H···π interaction-stacked porous supramolecular framework (PSF) with both dynamic structure and post-transformation is an important but currently challenging subject.In general, the CPMs stabilized by C-H···π interactions may show the similar dynamic behavior like third-generation MOFs due to the characteristic of non-covalent interactions, and can reversibly transform their structures in response to chemical guests via physical stimulationCu12a-π, based on non-covalent C-H···π interactions between cluster-based building units that are protected by 9-(prop-2-yn-1-yl)−9H-carbazole (Cbz-PrAH) and p-tert-butylthiacalix[4]arene (H4TC4A). Of these, the Cbz-PrAH contains extended aromatic ring, which is conducive to the formation of intra- and intermolecular non-covalent C-H···π interactions24; H4TC4A is a kind of macrocyclic ligand with intrinsic cavity structure29. As displayed in Fig. Cu12a), which can transform to guest-free PSF (Cu12a-π) through a dynamical single-crystal-to-single-crystal (SC-SC) fashion by removing guest molecules under heating and vacuum conditions. The monomer Cu12 nanocluster as a zero-dimensional (0D) cluster-based building unit, can be further supramolecularly assembled into C-H···π interactions-stacked PSF. Even more impressively, the post-transformation of Cu12a-π followed a dissociation-reassembly mechanism, giving the cluster-based nonporous adaptive crystals (NACs) Cu12b-NACs with updates of both structure and properties.To address these challenges, we designed a three-dimensional (3D) PSF, namely Cu12a, we selected copper(I) precursor, Cbz-PrACu  to react with H4TC4A via solvothermal method, giving the red rhombic crystals of Cu12a in a quite high yield of 91%  nanoclusters protected by Cbz-PrA‒ and TC4A4‒ ligands. The infrared (IR) spectroscopy confirmed the existence of Cbz-PrA‒ in Cu12a. Besides, elements of Cu12a were confirmed by energy-dispersive x-ray (EDX) spectroscopy. More synthetic details and basic characterizations are listed in the Supplementary Figures and Tables 4(TC4A)2]·(CH3COOCH2CH3). The structural anatomy of the monomer was shown in Fig. 4 units rotated by 45° in sequence, the equatorial copper atoms are bulged from the centres of four vertical faces of the cube about 0.446 Å. The Cu12 kernel exhibits a C4 symmetry with a C4 axis passing through the centres of two horizontal planes, and is consolidated by short Cu···Cu contacts (average distance of 2.76 Å), which are shorter than the sum of Bondi’s van der Waals radii of two Cu atoms (2.80 Å)30, implying an appreciable cuprophilic interactions. Furthermore, the types of capping ligands in the monomer of Cu12a are somewhat reminiscent of the Cu13a and Cu13b, where Cbz-PrA− and TC4A4− capped on the periphery of the copper kernel in our previous work31.Single-crystal X-ray diffraction (SCXRD) analyses revealed that the monomer of − ligands like nails hold only one coordination mode .The difference is that four Cbz-PrA36. Inspired by this, the guest EA molecules in Cu12a may endow it with similar behavior. Thus, the crystalline sample of Cu12a has been heated at 338 K upon vacuum for two days. Surprisingly, the activated sample of Cu12a, termed Cu12a-π, can still be analyzed via SCXRD, in which the guest EA molecules had been successfully removed to form permanent voids  layers of Cu12a and Cu12a-π both adopt AA stacking mode, but the latter occurred with a distinct displacement between each layers along the ab plane, resulting in a slight reduction in the porosity (from 11.5% to 9.1%). Impressively, upon further soaking Cu12a-π in EA solvent for three days, the lattice again recovered to P21/c accompanied with the re-adsorption of the EA molecules. This indicates that the Cu12a and Cu12a-π as host matrixes can achieve reversible SC-SC transformation by removal and re-adsorption of the guest molecule, thus demonstrating the crystalline structural flexibility of Cu12a. Thermogravimetric analysis (TGA)  for at least 72 h at room temperature, their PXRD patterns were highly consistent well with the theoretical patterns of Cu12a, which demonstrated the framework highly dynamic stability to chemical solvents  Fig.  indicate8 K Fig. . Besidesure Fig. , which ints Fig. . This ex12 cluster shows significant cavity structures above the poles of copper kernel, which is conductive to the close arrangement of cavity structures in one plane. And the higher rim of the cavity in Cu12 cluster shows a maximum pore diameter of ~8.9 Å. Therefore, Cu12 cluster can be regarded as a porous 0D cluster-based building unit, associated with two kinds of C-H···π interactions (C(sp3)-H···π(Cbz)=3.1 Å and C(sp2)-H···π(Cbz)=3.1 Å) between a carbazole ring and other carbazole ring, as well as between an allylene and a carbazole ring  stacking via almost negligible interlayer H···H interactions (2.26 and 2.93 Å) of tertiary butyl to form a final 3D supramolecular framework -H···π(Cbz)=3.2 or 3.4 Å and C(sp2)-H···π(Cbz)=2.8 Å) due to the incorporation of guest EA solvents.In Fig. ing Fig. , togetheork Fig. . Of thesCu12a-π, it is clearly seen that the small cavities provided by the thiacalix[4]arene and permanent voids between each layers effectively endow the Cu12a-π with certain porosity  surface area of 31.93 (45.29) m2·g−1, and a wide pore size distribution in the range of 0.8–1.6 nm calculated by Horvath-Kawazoe (HK) model  at normal pressure, further confirming the real existence of permanent porosity in Cu12a-π.On the basis of packing characteristic of 41. Inspired by this, the Cu12 nanocluster acting as a building unit may feature ligand-exchange to remodel the PSFs. Therefore, we tried to transform Cu12a-π by dissolving crystalline samples in an isopropanol/tetrahydrofuran (v:v = 3:3) mixture, to which an excess of H4PTC4A  was added, and then the above solution continued to react under solvothermal condition to generate red block crystals of Cu12b. Except for the p-phenyl-thiacalix[4]arene CT in Cu12b leading to the variation in the absorption intensity of the two nanoclusters at ~300 nm (Please see details in the Supplementary Methods); (ii) the position of the maximum absorption peaks of the two ligands (H4TC4A and H4PTC4A) . As shown in Fig. Cu12a and Cu12b in mixed CH3OH and CHCl3 exhibit a series of +1 species  at the m/z range of 3010-3090 and 3170-3345, respectively, which correspond to their intact cluster bound with different cations or solvents. The most dominant peak 1b at m/z 3036.9040 and peak 2b at m/z 3196.6478 can be attributed to [Cu12(TC4A)2(Cbz-PrA)4Na]+  and [Cu12(PTC4A)2(Cbz-PrA)4Na]+ , respectively. However, as the reaction of H4PTC4A and Cu12a proceeded in CHCl3, the parent peaks of Cu12a (1a−1e) progressively faded away within 20 min  appeared at 30 min, and can be assigned to [Cu12(PTC4A)(TC4A)(Cbz-PrA)4H]+. This species 3a is the mono-substituted product of Cu12a, in which one of the TC4A4- ligands is replaced by PTC4A4-. After 80 min, we observed a species 2j, [Cu12(PTC4A)2(Cbz-PrA)4(CH3OH)(H2O)2K]+, which can be identified as the bis-substituted product of Cu12a. This result indicates that Cu12b is more thermodynamically stable than Cu12a, which also was confirmed by DFT calculations . More formula assignments for the above species in ESI-MS were performed as listed in Supplementary Table To verify the assumption on the transformation mechanism, we monitored the ligand-exchange process in the structural transformation from min Fig. , which i12 clusters was initially presumed to be a simple ligand-exchange process based on the single crystal structure and optical monitoring. However, further investigation using ESI-MS tracking revealed that this transformation was not a straightforward ligand-substitution, but rather a kinetically driven dissociation of the initial cluster, followed by a reassembly to form a thermodynamically stable new cluster. Thus, we have established the dissociation-reassembly (DR) mechanism to describe above transformation process, highlighting the dynamic nature of the dissociation and subsequent reassembly steps. The DR mechanism provides a conceptual framework to understand the intricate dynamics involved in the transformation of Cu12a to the more stable Cu12b50.The transformation between the two Cu4‒, we further checked the packing formed by Cu12 cluster in Cu12b. There is one THF molecule residing in the cavity of each PTC4A4‒ in Cu12b, which reduces the void produced by PTC4A4‒ ligand itself, so the Cu12b is a nonporous structure. The activated experiment of Cu12b was further carried out, and the activated sample namely Cu12b-NACs had been successfully removed the guest THF solvents , which are relatively nonporous but with medium surface areas56. In light of this, the analysis of N2 adsorption isotherm of Cu12b-NACs gives an experimental Brunauer-Emmett-Teller (Langmuir) surface area of 14.88 (12.63) m2·g-1, which is less than half of that of Cu12a-π, and the pore-size distribution exhibits broad and ambiguous peak  image of I2@Cu12a-π showed the crystal-like morphology  and Cu12a-π, which has been reported previously for iodine adsorption materials66. Moreover, the I3d X-ray photoelectron spectrum (XPS) of I2@Cu12a-π was measured, and two broad peaks at ~620 and ~632 eV indicate the presence of the I3‒ and I5‒ anions, respectively  with cluster-based building units held together through intralayer non-covalent C-H··π interactions. The Cu12a-π is successfully constructed through the SC-SC transformation by the removal of guest molecules from pristine Cu12a, showing unexpected structural dynamics and superior thermal stability than most reported π-stacked frameworks. Moreover, the changing in upper rim of thiacalix[4]arene, Cu12a-π can be transformed into the nonporous adaptive crystal Cu12b-NACs following a DR mechanism, realizing the structural and performance updates. Based on the permanent porosity, Cu12a-π can absorb iodine in aqueous solution reaching a high capacity of 2.96 g·g‒1. Overall, all unique structural features as well as the iodine adsorption performance of such cluster-based PSF are indicative of a bright future of this type of porous materials.In summary, we have successfully developed a dynamic and transformable 3D PSF  were mixed in 6.00 mL methanol/ethyl acetate (v:v = 3:3). The resulting suspension was sealed and heated at 80 °C for 2000 min. After cooling to room temperature, red crystals of Cu12a were formed . Selected IR peaks (cm−1): 2950 (s), 2906 (w), 2860 (w), 1732 (s), 1584 (m), 1450 (s), 1364 (m), 1315 (m), 1257 (s), 1208 (m), 1037 (m), 926 (w), 885 (w), 840 (s), 751 (s), 711 (s), 625 (w), 531 (m).Cbz-PrACu  and HCu12a  were dissolved in mixed solvents of 6.00 mL isopropanol/tetrahydrofuran (v:v = 3:3), then H4PTC4A  was added successively. The resulting suspension was sealed and heated at 80 °C for 2000 min. After cooling to room temperature, red crystals of Cu12b were formed . Selected IR peaks (cm−1): 2955 (s), 2901 (w), 2865 (w), 1777 (w), 1589 (m), 1454 (s), 1364 (m), 1311 (s), 1255 (s), 1202 (m), 916 (w), 876 (w), 831 (s), 741 (s), 714 (s), 616 (w), 526 (m).The crystals of Supplementary InformationPeer Review FileDescription of Additional Supplementary FilesSupplementary Data 1-3Source Data


The Pt on β-MnO2 inhibited CO oxidation below 210 °C but promoted it above 210 °C. A Pt/β-MnO2 catalyst contains more Pt4+ and less Pt2+. The results of operando DRIFTS-MS show that surface-terminal-type oxygen (M=O) plays an important role in CO oxidation. When the temperature was below 210 °C, Mn=O consumption on Pt/β-MnO2 was less than β-MnO2 due to Pt4+ inhibition on CO oxidation. When the temperature was above 210 °C, Pt4+ was reduced to Pt2+, and Mn=O consumption due to CO oxidation was greater than β-MnO2. The interaction of Pt and β-MnO2 is proposed.Pt-based catalysts exhibit unique catalytic properties in many chemical reactions. In particular, metal–support interactions (MSI) greatly improve catalytic activity. However, the current MSI mechanism between platinum (Pt) and the support is not clear enough. In this paper, the interaction of 1 wt% Pt nanoparticles (NPs) on  The low-intensity peak with a low wavenumber is attributed to the deformation mode of the metal–oxygen chain (Mn-O-Mn) in the MnO2 octahedral lattice out are the CO concentrations in gas streams from the inlet and outlet of the reactor, respectively.F is the total gas flow rate in mol s−1, catm is the amount of catalyst in g, Ptm is the weight ratio of Pt in the catalyst, and PtM is the molar atomic weight of Pt (195.08).The CO oxidation activity of two catalysts  was measured in a fixed-bed flow reactor fed with 1% CO/20% O g−1 h−1 . CO and The operando DRIFTS-MS was used to monitor surface species and gaseous products during CO oxidation using the method reported elsewhere . The catβ-MnO2 using deposition–precipitation was used to explore the effect of the metal–support interaction of Pt and β-MnO2 in CO oxidation. The main conclusions are as follows:In this study, 1 wt% Pt NPs loaded on (1)β-MnO2, it was found that Pt NPs (1–2 nm) were anchored on β-MnO2 and formed Pt-O-Mn interfaces. CO oxidation was inhibited at a temperature below 210 °C and promoted above 210 °C;After loading Pt on (2)+2 dominates CO oxidation. The ratios of Pt+2 before the reaction and after the reaction at 150, 200, and 250 °C were Pt/β-MnO2-fresh (0.32) < Pt/β-MnO2-used-150 °C (0.48) < Pt/β-MnO2-used-200 °C (0.51) < Pt/β-MnO2-used-250 °C (0.64). Pt4+ can convert to Pt+2 by heating;The ratio of Pt(3)β-MnO2 than on Pt/β-MnO2. When the temperature is higher than 210 °C, more Mn=O was consumed on Pt/β-MnO2 than on β-MnO2. It is speculated that the inhibition of activity below 200 °C after Pt loading is due to Pt+4, which causes less Mn=O consumption for CO oxidation;The Operando DRIFTS-MS results show that Mn=O plays an important role in CO oxidation. Below 210 °C, more Mn=O was consumed on (4)β-MnO2 interaction is proposed, where the Pt+4 and Pt+2 functions are clearly illustrated.The mechanism of Pt and


Here, we present the improved and more efficient synthesis of both Fmoc-GlcAPC-OH and Fmoc-GlcAPC(Ac)-OH, β-SAAs consisting of only 4–5 synthetic steps. Their active ester and amide bond formation with glycine methyl ester (H-Gly-OMe) was completed and monitored by 1H NMR. The stability of the pyranoid OHs protecting the acetyl groups was investigated under three different Fmoc cleavage conditions and was found to be satisfactory even at high piperidine concentration (e.g. 40%). We designed a SPPS protocol using Fmoc-GlcAPC(Ac)-OH to produce model peptides Gly-β-SAA-Gly as well as Gly-β-SAA-β-SAA-Gly with high coupling efficiency. The products were deacetylated using the Zemplén method, which allows the hydrophilicity of a building block and/or chimera to be fine-tuned, even after the polypeptide chain has already been synthesized.The synthesis of The online version contains supplementary material available at 10.1007/s00726-023-03278-1. Recently, we have investigated and optimized different coupling reagents to generate GXXG model peptides  and remain stable for at least 6–24 h, significantly longer than the duration of a typical coupling cycle (3 h).Although a large number of coupling reagents and active esters have been used in the past  and its per-O-acetylated derivative Fmoc-GlcAPC(Ac)-OH (2). Although O-acetyl protection is common in carbohydrate chemistry, and commonly used in the synthesis of glycopeptides and related compounds -OH (3). However, we found that this route—which consists of 9 steps—is too long and that it could be shortened and optimized. Furthermore, we wanted to prepare building blocks suitable for SPPS  was obtained in four steps and the final product 3 was obtained in five consecutive steps. The chain elongation leading to the nitrile derivative (5), made by OAc → Br → CN exchange, are the key steps of this synthesis. Subsequently, 5 was hydrolyzed in three steps to give the fully unprotected d-glucosamine-1-carboxylic acid , which was then acetylated and N-protected by Boc to give Boc-GlcAPC(Ac)-OH (3), a molecule finally suitable for synthesis using solution-state protocols. This building block was used to complete the homooligomer synthesis with BOP/DIEA, a stepwise synthesis in solution, resulting in the dimer, trimer, etc. up to the hexamer -Ac (10), chain elongation with TMSCN led directly to the nitrile derivative 5 (Scheme 5) gave the HCl salt of the unprotected H-GlcAPC-OH (8) in two steps: (a) Zemplén deacetylation removed the acetyl protection; (b) hydrolysis with 12% aqueous NaOH was carried out to partially remove the N-Phth protection and hydrolysis with 2 M HCl gave H-GlcAPC-OH (8). Then, this was converted to the Fmoc protected derivative, Fmoc-GlcAPC-OH (1). In the final step, the OH groups were conventionally acetylated (Ac2O in pyridine) to give the fully protected β-SAA, Fmoc-GlcAPC(Ac)-OH (2), which was shown (as detailed below) to be suitable for SPPS. On the other hand, our fine-tuned procedure is based on 5 Scheme .The hydr1) and obtained Fmoc-GlcAPC(Ac)-OH (2) in four and five steps, with overall yields of 36% and 29%, respectively. The process is also more environmentally friendly by reducing the number of synthetic steps. The key step, chain elongation, is performed in a single step. After their formation, both 1 and 2 can be precipitated with 2 M HCl. Molecules 5 and 2 were both successfully recrystallized from ethanol, eliminating column chromatography and providing a greener synthetic route.Our method is more efficient than that of Suhara et al. who obtain their final compound, Boc-GlcAPC(Ac)-OH, in 9 steps with an overall yield of 26%. We achieved our target molecules Fmoc-GlcAPC-OH -OH) as a function of time  and Fmoc-GlcAPC(Ac)-OBt (12), whose formation was indicated by the characteristic chemical shift changes of the aromatic HD with respect to the parent HD’  requires ~ 1 h, resulting in an almost quantitative (96%) conversion  was slower (~ 3 h), but again the conversion was almost quantitative (~ 99%) and the product was stable even after 24 h. Note how useful the NMR monitoring of the reaction is for these β-SAAs, we were able to determine that the activation is slower than expected (1–3 h), but in return the active ester formations are almost quantitative. We have previously investigated several coupling reagents and conditions to obtain the best method for β-SAAs coupling  and Fmoc-GlcAPC(Ac)-OBt (12), was investigated with glycine, the simplest and most flexible proteinogenic amino acid residue. The C-terminal of Gly was protected as a methyl ester: H-Gly-OMe. Slow amide bond formation was observed for both Fmoc-GlcAPC-OBt (11) and Fmoc-GlcAPC(Ac)-OBt (12) in the presence of 2 equivalents of H-Gly-OMe and 6 equivalents of DIEA: even after 24 h the conversion was only 63% and 17%, respectively. -OBt 1, was invN-phthaloyl-protected model compound Phth-GlcAPC(Ac)-NH2 (13) was used, as the Phth-group is an inexpensive and simple amide protecting group  the chimera synthesis is not too long, (ii) the sequence is not too long and (iii) the sugar amino acid is not too close to the C-terminus of the sequence.As our intention was to develop Fmoc chemistry suitable β-SAAs using acetylated derivatives, e.g. Fmoc-GlcAPC(Ac)-Gly-OMe, the question arises to what extent an acetyl protecting group resists cleavage conditions. Therefore, we investigated the stability of this group under conventional Fmoc cleavage conditions and beyond. The Ac)-NH2 1 was usedAc)-NH2 1 was usedoup Fig. , 40% pip1, 2) are promising candidates for coupling to SPPS. Having both the O-acetylated Fmoc-GlcAPC(Ac)-OH (2) and its “free” derivative Fmoc-GlcAPC-OH (1) in hand, SPPS was investigated for both. Note that the free β-SAA 1 has a total of 3 free -OH groups as nucleophiles, which could cause miscoupling, especially that of the primer -OH of the C-6 position. Armed with the previously established NMR data on active ester formation and coupling data, SPPS was used to generate the Ac-GXXG-NH2 model chimera (X = β-SAA: -GlcAPC- and -GlcAPC(Ac)-). Using this model peptide, we investigated all 3 different conditions of amide bond formation, namely those of α-β, β-β and β-α amide formation. For Fmoc-Gly-OH coupling, DIC/HOBt was used (1 h coupling time), whereas for β-SAAs PyBOP/DIEA was used (3 h). For Fmoc cleavage, 2% piperidine and 2% DBU in DMF protocol was used. When using Fmoc-GlcAPC-OH, a building block with free -OH groups, we found low coupling efficiencies -OH was successfully incorporated into the Ac-GXXG-NH2 model chimera: both coupling efficiencies (84–91%) and yields (69–90%) are acceptable . Thus, both the milder and the harsher TFA conditions allow to obtain the desired peracetylated chimera.On the other hand, the per-cies 84–9% and yiele Table . Using tO-acetyl protecting groups, Ac-GXG-NH2 was synthesized and studied, as partial deacetylation is easy to detect on this simple chimera . The final cleavage was carried out at 40 °C with 95% TFA and we found that the fully O-acetylated chimera is intact. Neither the MS spectrum nor the HPLC chromatogram show any sign of decomposition .To further investigate the stability of the 2. As the deacetylation was performed in MeOH, which induces the collapse of the S RAM Tentagel® resin, the reaction was performed after the final cleavage. The purified per-O-acetyl Ac-GXG-NH2 was dissolved in MeOH and a few drops of 2 M NaOMe/MeOH solution were added. After 1 h at room temperature, the reaction was stopped by adding TFA to the mixture. The result was a fully -OH deprotected Ac-GXG-NH2 chimeric peptide . In summary, the O-acetyl groups were successfully removed by the Zemplén deacetylation method, whereas the conventional 50% or 95% TFA final cleavage mixture did not interfere with the O-acetyl groups.To obtain the fully unprotected chimera, the Zemplén deacetylation method, a reaction widely used in carbohydrate chemistry to remove acetyl protection, was performed on Ac-GXG-NH1) and Fmoc-GlcAPC(Ac)-OH (2), suitable for SPPS were synthesized in only four and five synthetic steps. Using NMR measurements, we found that the corresponding active esters are formed with PyBOP/DIEA in 1–3 h and when coupled with an α-amino acid, H-Gly-OMe, they form a β-α-amide bond. The stability of the O-acetyl protecting groups was confirmed by 1H NMR under Fmoc cleavage conditions using Phth-GlcAPC(Ac)-NH2 (13) as a model. Both Ac-GXG-NH2 and Ac-GXXG-NH2 chimeras with O-acetyl protected β-SAA (2) were successfully synthesized in SPPS: both coupling efficiencies and overall yields were found to be sufficient. The acetyl groups were removed by the Zemplén deacetylation method after final resin cleavage. In conclusion, it is possible to prepare chimeric peptides containing β-α-, α-β- and/or β-β-amide linkages using the new and efficiently synthesized β-SAA, Fmoc-GlcAPC(Ac)-OH (2). In addition, this biocompatible synthetic product is even more versatile and tunable, because the O-Ac groups of the β-SAA can be retained or removed at the end of the synthesis.In the present work, new β-sugar amino acids, namely Fmoc-GlcAPC-OH  can be found in Supporting information, in the online version.2. Solvents and reagents for MS measurements were purchased from VWR. TLC was performed on silica gel 60 F254, 230 mesh (E. Merck) and spots were detected by UV light (254 nm), charring with 5% H2SO4 solution.Reagents, materials and solvents were purchased from Sigma-Aldrich, Merck, Reanal, VWR and Iris Biotech. Moisture-sensitive solvents were dried on molecular sieve (3 Å), while acetonitrile was distilled from CaH2O (A) and 0.08% TFA, 95% ACN/5% H2O (B), flow rate 0.9 ml/min and UV detection at 220 and 280 nm. Gradient was as follow: 0 min: 5% B, 30 min: 95% B, 33 min: 95% B, 33.1 min: 5% B, 45 min: 5% B.Chimera peptides were measured by RP-HPLC on Aeris™ 3.6 μm peptide XB-C18 100 Å, LC Column 250 × 4.6 mm with eluents 0.1% TFA in H1H and 125.76 MHz for 13C. The sample concentrations ranged from 0.1 M to 0.2 M. Spectra were recorded in DMF-d7 using the solvent residual peak as the 1H internal reference. Spectra evaluation was done with TopSpin 4.1.1 software.NMR experiments were performed at 298 K on Bruker Avance DRX 500 MHz spectrometer equipped with TXI probe with z-gradient, operating at 500.13 MHz for The mass spectrometry measurements were performed using a Bruker Amazon SL ion trap mass spectrometer equipped with an electrospray ion source.3,4,6-Tri-O-acetyl-2-deoxy-2-phthalimido-β-d-glucopyranosyl-1-carboxamide (Phth-GlcAPC(Ac)-CN, 5) Tetra-O-acetyl-2-deoxy-2-phthalimido-d-glucopyranose  was dissolved in anhydrous acetonitrile (25 ml). Then, TMSCN  and BF3‧OEt2  were added to this solution. The reaction mixture was stirred at room temperature overnight. After TLC (toluene:EtOAc 3:2) indicated the completion of the reaction, water (10 ml) was added to the reaction mixture, followed by 30 min of stirring. The solution was concentrated in vacuo. Then, CHCl3 (40 ml) was added to the residue orange oil, and the solution was washed three times with water. The organic phase was dried (MgSO4), filtered and concentrated in vacuo. The remaining crude product was crystallized from EtOH, twice. Yield: 1.55 g (40%). TLC : Rf = 0.51; [α]D22 =  + 65.3 , lit. (Myers 1984) [α]D22 = 68.8 1H NMR  δ 7.89–7.74 , 5.72 , 5.35 , 5.17 , 4.64 , 4.31–4.13 , 3.89–3.83 .2-Deoxy-2-(9-fluorenylmethoxycarbonylamino)-β-d-glucopyranosyl-1-carboxylic acid  Phth-GlcAPC(Ac)-CN  was suspended in 12 m/m% NaOH solution (40 ml) and refluxed at 18 h. Then, cc HCl was added into the solution to get 2 M HCl concentration, and it was continued boiling at 18 h. After TLC (MeOH:AcOH 6:1) showed completion of the reaction, the mixture was left to cool to room temperature and filtered. The remaining solution was extracted with EtOAc (three times). Then, the water phase was concentrated in vacuo to remove HCl gas, and the remaining solution was lyophilized. Then, the dried product was dissolved in water/methanol . The pH of the solution was set to pH 8 with saturated aqueous NaHCO3 solution. Then, a solution of Fmoc-OSu  in THF (15 ml) was added, and the pH was again set to pH 8 with NaHCO3 solution. The reaction mixture was stirred at room temperature for 2 days. After the reaction was completed, it was concentrated in vacuo to remove organic solvents. The remaining aqueous solution was diluted to 15 ml with water, pH set to pH 8 and extracted with EtOAc (5 times). The pH of the aqueous phase was set to 2 and it was cooled to 0 °C. The precipitate was filtered and dried. Yield: 0.85 g (88%). TLC : Rf = 0.34; [α]D22 =  + 19.8 , 1H NMR  δ 7.99 ; 7.88 ; 7.44 ; 7.30 ; 4.21 ; 4.21 ; 4.15 ; 3.95 ; 3.82 ; 3.72 ; 3.56 ; 3.36 ; 3.28  13C NMR  δ 170.7 (COOH); 156.3 (NHC(O)); 144.4 (Fmoc Ar2-C1); 144.2 (Fmoc Ar1-C1); 141.2 (Fmoc Ar2-C6); 141.1 (Fmoc Ar1-C6); 128.07 (Fmoc Ar1-C4); 128.04 (Fmoc Ar2-C4); 127.6 (Fmoc Ar2-C3); 127.5 (Fmoc Ar1-C3); 125.9 (Fmoc Ar1-C2); 125.7 (Fmoc Ar2-C2); 120.54 (Fmoc Ar1-C5); 120.5 (Fmoc Ar2-C5); 81.5 (C1); 78.3 (C5); 75.1 (C3); 70.9 (C4); 66.03 (Fmoc CH2); 61.5 (C6); 55.3 (C2); 47.1 (Fmoc C(2Ar)).3,4,6-Tri-O-acetyl-2-deoxy-2-(9-fluorenylmethoxycarbonylamino)-β-d-glucopyranosyl-1-carboxylic acid (Fmoc-GlcAPC(Ac)-OH, 2) Fmoc-GlcAPC-OH  was suspended in pyridine (5 ml) and cooled to 0 °C, then, Ac2O (3 ml) was slowly added to this solution, and it was left to warm to room temperature, and stirred overnight. After TLC (EtOAc:MeOH 4:1) shows completion of the reaction, the reaction mixture was poured to ice-water (25 ml). The product was precipitated and filtered. The crude product was recrystallized from EtOH. Yield: 1.1 g (85%). TLC : Rf = 0.31; [α]D22 =  + 193.5 , 1H NMR  δ 7.89 ; 7.64 ; 7.41 ; 7.31 ; 5.37 ; 4.98 ; 4.35 ; 4.23 ; 4.21 ; 4.19 ; 4.16 ; 4.11 ; 3.98 ; 3.93 ; 2.0 and 1.9  13C NMR  δ 170.2 (COOH); 169.9, 169.6, 169.2 (Ac-C(O)) 156.1 (NHC(O)); 144.4 (Fmoc Ar2-C1); 144.0 (Fmoc Ar1-C1); 141.2 (Fmoc Ar2-C6); 141.1 (Fmoc Ar1-C6); 127.7 (Fmoc Ar1-C4); 127.7 (Fmoc Ar2-C4); 127.2 ; 125.5 (Fmoc Ar1-C2); 125.4 (Fmoc Ar2-C2); 120.1 ; 77.4 (C1); 75.5 (C5); 73.7 (C3); 68.9 (C4); 66.5 (Fmoc CH2); 62.4 (C6); 53.6 (C2); 47.0 (Fmoc C(2Ar)).® resin was used . Resin was swollen in DCM. The first step was the removal of the Fmoc group with common method . The successful cleavage was analyzed by the Kaiser test. After that, the coupling of αAAs to resin was made by reagent pairs HOBt/DIC (αAA:HOBt:DIC 1:1:1) in DMF for 1 h, while that of βSAA either Fmoc-GlcAPC-OH (1) or Fmoc-GlcAPC(Ac)-OH (2) was accomplished by PyBOP/DIEA (βSAA:PyBOP:DIEA 1:1:2) in DMF for 3 h. Finally, resin was acetylated with Ac2O:DIEA:DMF  for 45 min. Resin was washed with 3 × DMF, 3 × DCM, 1 × Et2O and dried in vacuo after finishing coupling and acetylation. During synthesis, the Fmoc group was removed with 2% piperidine and 2% DBU in DMF (3 + 17 min) and indicated by Kaiser test. The capacity of the resin was determined by UV–Vis measurement regarding to Fmoc chromophore amount (Fmoc-piperidine adduct) released by using 50% piperidine in DMF  for 3 h. Resin was washed with 2 × DMF, 3 × DCM and 2 × MeOH and solvent was removed in vacuo. The crude products were precipitated with diethyl ether.For the SPPS S RAM Tentagel2 (20 mg) was dissolved in MeOH (2 ml) and 2 M NaOMe/MeOH (1 ml) solution was added. After 1 h at room temperature, the reaction was stopped with the addition of TFA (1 ml). The acidic solution was concentrated to get white solids (10 mg).The purified Ac-Gly-GlcAPC(Ac)-Gly-NHAc-Gly-GlcAPC(Ac)-Gly-NH2m/z): [M + H]+ calcd. For 489.18; found: 489.15.Rp-HPLC: 11.82 min, MS -GlcAPC(Ac)-Gly-NH2m/z): [M + H]+ calcd. for 804.27; found: 804.21.Rp-HPLC: 16.01 min, MS : [M + H]+ calcd. for 363.05; found: 363.06.Rp-HPLC: 3.61 min, MS Below is the link to the electronic supplementary material.