In the crystal structure, the cations form hydrogen-bonded zigzag chains through centrosymmetric cyclic  4H8N5+·C2Cl3O2−·H2O, coomprises a 2,4-diamino-6-methyl-1,3,5-triazin-1-ium cation, a trichloroacetate anion and a water molecule of solvation. The protonated N atom of the cation forms a hydrogen bond with a carboxyl O atom of the anion, which also acts as a hydrogen-atom acceptor with the water molecule. The cations form centrosymmetric dimeric units through R22(8) N—H⋯N bond pairs and are extended into zigzag chains along the c-axis direction, also through similar cyclic R22(8) dual N—H⋯N hydrogen-bonding interactions. The water molecule acts as a dual acceptor forming N—H⋯O hydrogen bonds between the amine groups of the cations, forming cyclic R23(8) motifs. The second H atom of the water molecule also acts as a donor in an O—H⋯O hydrogen bond with the second carboxyl O atom, linking the chains along the b-axis direction. These interactions give rise to an overall three-dimensional supramolecular structure. A Hirshfeld surface analysis was employed in order to study the intermolecular interactions.The asymmetric unit of the title molecular salt, C Proton transfer occurs from one of the carboxylic acid oxygen atoms (O1) to atom N5 of the cation, with a resulting N5—H1N5⋯O1 hydrogen bond  and QUWXIQ (with 3-hydroxypyridine-2-carboxylic acid) ], ZUDSUO [with 6-chlorouracil-1-ide ]  of 2,4-diamino-6-methyl-1,3,5-triazine (1.25 mg) and an aqueous solution (10 ml) of trichloroacetic acid (1.63 mg) in a 1:1 molar ratio. The reaction mixture was warmed over a water bath for a few minutes. The resultant solution was then allowed to cool slowly at room temperature. After a few days, colourless block-shaped crystals of the title compound were separated out.2), with Uiso(H) set to 1.2Ueq. The water-bound H atoms were located in a difference-Fourier map and were freely refined [O—H = 0.78 (4) and 0.86 (4) Å].Crystal data, data collection and structure refinement details are summarized in Table 210.1107/S2056989018008307/zs2400sup1.cifCrystal structure: contains datablock(s) global, I. DOI: 10.1107/S2056989018008307/zs2400Isup2.hklStructure factors: contains datablock(s) I. DOI: Click here for additional data file.10.1107/S2056989018008307/zs2400Isup3.cmlSupporting information file. DOI: 1587723CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


The molecular Hirshfeld surfaces analysis were explored with two-dimensional fingerprint plots for the title compounds and other known structures from the literature. Additionally, the lipophilicity parameters (logP) were determined and related to the C⋯H contact contribution in the Hirshfeld surface.Two biologically active compounds were synthesized and their crystal structures were determined. The characteristic feature of both structures is molecular layers in the crystal lattice formed  E)-3-(4-hydroxybenzylidene)chroman-4-one, C16H12O3, I, and (E)-3-(3-hydroxybenzylidene)-2-phenylchroman-4-one, C22H16O3, II, are reported. These compounds are of interest with respect to biological activity. Both structures display intermolecular C—H⋯O and O—H⋯O hydrogen bonding, forming layers in the crystal lattice. The crystal structure of compound I is consolidated by π–π interactions. The lipophilicity (logP) was determined as it is one of the parameters qualifying compounds as potential drugs. The logP value for compound I is associated with a larger contribution of C⋯H interaction in the Hirshfeld surface.The synthesis and crystal structures of ( The dihedral angles between the hydroxybenzylidene rings and the main chroman skeleton are 47.54 (8) and 69.46 (12)°, respectively, for I and II —C2—C3—C4 torsion angle in the range 82.44–107.90°. In both chromanone and flavanone structures, the pyran ring adopts a slightly distorted envelope conformation. In the 41 chromanone derivatives, the bond distances and angles within the chroman moiety are in good agreement with those found in compound I.A search of the Cambridge Structural Database  of compounds I and II was determined using the RP–TLC method. The values of logP obtained are 2.95 and 3.98, respectively for I and II, the difference being due to the different, bulky substituent at the C2 position of the pyran ring. The theoretical values of lipophilicity (miLogP) also show the same trend, the value for compound I is lower (miLogP = 3.14) than that for compound II (miLogP = 4.70). This is in agreement with the values previously reported for similar arylidenochromanone/flavanone derivatives . According to the ‘rule of five’ proposed by Lipinski et al.  and molar mass (< 500 Da).The experimental lipophilicity -2-phenyl-2,3-dihydro-4H-chromen-4-one -3-chroman-4-one -3--2-phenylchroman-4-one  analysis may provide useful descriptors for QSAR study , we found a relationship between the logP value and the fraction of the Hirshfeld surface covered by different intermolecular interactions. The increase of logP corresponds in fact to increasing the C⋯H contribution in the Hirshfeld surface. Furthermore, for compounds I–V, the contribution of the O⋯H interaction in the Hirshfeld surface is inversely proportional to the value of logP.As in our previous studies  and five drops of piperidine was heated at 413 K in an oil bath for four h. The progress of the synthesis was controlled by thin layer chromatography (TLC) using toluene/methanol (9:1 v/v) as eluent. After cooling the reaction mixture was left for 24 h at room temperature. The solidified product was filtered and crystallized from methanol. Compound I was obtained as a yellow powder. The isolated solid was further recrystallized by slow evaporation at room temperature of an acetone solution. Yield: 64%, M.p.: 501–502.5 K. MS (ESI+): m/z 253.3 C16H12O3 [M+H]+. IR (KBr): 3126 (O—H), 2809 (C—Haromat), 1652 (C=O), 1608, 1578 (C=C), 1164 (C–O—C), 751 (=C—H). 1H NMR  δ (ppm): 5.42 , 6.86–7.86 , 7.87 , 10.12 . Analysis calculated for C16H12O3 (M = 252.23 g mol−1) % C: 76.18; % H: 4.81; % O: 19.01. Found % C: 75.3; % H: 5.01; % O: 19.69.The synthesis of compounds ne Fig. 8. CompounII was synthesized according to the procedure described by Pijewska et al., (1993v/v). After cooling at room temperature, the mixture was dissolved in methanol. After 24 h compound II precipitated as a light-cream fine crystalline powder and was purified by crystallization from methanol. Crystal suitable for X-ray analysis were obtained by slow evaporation of an ethanol solution at room temperature. Yield: 52.4%. M.p.: 482–483 K. MS (ESI+): m/z 329.2 C22H16O3 [M+H]+. IR (KBr): 3297 (O—H), 3054 (C—Haromat), 2351 , 1663 (C=O), 1608, 1590, 1504 (C=C), 1141 (C—O—C), 757 (=C—H). 1H NMR  δ (ppm): 6.57 , 5.69 , 6.89–7.91 , 8.12 . Analysis calculated For C22H16O3 (M = 328.19 g mol−1) %C: 80.51; %H: 4.87; % O: 14.62. Found %C: 79.99; %H: 5.11; % O: 14.90.Compound al., 1993. A mixtuUiso(H) = 1.5Ueq(O) or 1.2Ueq(C). A rotating model was used for the hydroxy groups.Crystal data, data collection and structure refinement details are summarized in Table 410.1107/S2056989019015639/rz5266sup1.cifCrystal structure: contains datablock(s) I, II. DOI: 10.1107/S2056989019015639/rz5266Isup4.hklStructure factors: contains datablock(s) I. DOI: 10.1107/S2056989019015639/rz5266IIsup5.hklStructure factors: contains datablock(s) II. DOI: Click here for additional data file.10.1107/S2056989019015639/rz5266Isup4.cmlSupporting information file. DOI: Click here for additional data file.10.1107/S2056989019015639/rz5266IIsup5.cmlSupporting information file. DOI: 1966749, 1966750CCDC references:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


However, a direct experimental test of this mechanism has not yet been performed. To evaluate the potential control of microbial N mineralization on the natural N isotope composition, we cultured fungi (Aspergillus oryzae) in five types of media of varying C:N ratios of 5, 10, 30, 50, and 100 for 4 d, and tracked changes in δ15N in the microbial biomass, NH4+, and dissolved organic N (DON: glycine) over the course of the experiment. High rates of NH4+ excretion from A. oryzae were accompanied by an increase in δ15N in the microbial biomass in low C:N media . In contrast, NH4+ was strongly retained in higher C/N treatments with only minor  changes being detected in δ15N in the microbial biomass. Differences in δ15N in the microbial biomass were attributed to the loss of low-δ15N NH4+ in low, but not high C/N substrates. We also detected a negative linear correlation between microbial nitrogen use efficiency (NUE) and Δ15N (δ15N-biomass–δ15N-glycine). These results suggest an isotope effect during NH4+ excretion in relatively N-repleted environments in which microbial NUE is low, which may explain the vertical patterns of organic matter δ15N in soil profiles.Changes in  The microbial decomposition of soil organic matter (SOM) aids in the mineralization of N in plants, which maintains primary productivity , 53 and  N cycle , 30, 51, N cycle . Therefo13C and 15N NMR analyses revealed that SMB in soil minerals contributed to SOM pools (e.g. the dominant pathway (62%) through which C enters the SOM pool, exceeding C input via leaf litter and fine root turnover, with potentially similar patterns for N  accounts for a significant fraction of the soil N pool  and contOM pools , 33. TheOM pools , 17, 31,ns for N .15N/14N  was previously used to examine relationships among SOM and SMB  SOM markedly varies among ecosystems, from −7.8‰ in acidic tundra , and Sollins et al.  varying widely across soils , while δ15N-SMB shows markedly less enrichment when net N is immobilized (incorporate) at high C/N (N limited conditions).δrocedure , 62, wasss soils . Furthersoil C/N , 41. Dij  being lower than that of the biomass in cows . NUE is defined as the ratio of N incorporation into the microbial biomass to organic N uptake , a well-studied, cosmopolitan strain of fungi, was used in the present study. Five milliliters of A. oryzae (ca. 8×107 cells) was inoculated into 500-mL flasks with 250 mL of medium. Cultures were incubated at 27°C with shaking (180 rpm) for 96 h. Porous silicon plugs were used to maintain aerobic conditions. We used glycine and glucose as the sole C and N sources to adjust the C:N ratio of medium. Five types of media with C:N ratios of 5, 10, 30, 50, and 100  contained glycine and D(+)-glucose at the following amounts: 3.26 and 4 g L−1, 1.63 and 5.2 g L−1, 0.5 and 5.6 g L−1, 0.25 and 4.8 g L−1, and 0.2 and 7.9 g L−1, respectively. The actual initial C:N ratios of media were 5.4±0.1, 11.1±0.2, 31.3±0.5, 52.5±2.0, and 103.6±3.8 . All media contained 0.5 g L−1 MgSO4·7H2O, 0.07 g L−1 CaCl2·2H2O, 0.01 g L−1 FeSO4·7H2O, 4.65 mg L−1 MnSO4·5H2O, 5.0 mg L−1 ZnSO4·7H2O, and 1.0 mg L−1 CoCl·6H2O. pH was adjusted to between 5.7 and 5.9 by adding 0.92 g L−1 K2HPO4 and 6.86 g L−1 KH2PO4. Media were filtered through pre-sterilized membrane filters , and filtrates were poured into pre-combusted  500-mL flasks. A total of 1.0 mg L−1 FeSO4·7H2O was added to medium without sterilization because of its oxidization during autoclaving as well as its precipitation. We had 3 replicates, except for CN 5 which had 4 replicates.A. oryzae was calculated from the freeze-dried biomass weight.Samples were collected every 24 h for a total of 4 d. Five milliliters of media, including the biomass, was collected using a 5-mL pipette and filtered through pre-combusted  glass fiber filters  with a pore size of 0.22 μm , while all media in the flasks were filtered with a membrane filter  to completely recover the biomass in the flask at 96 h. The filtrate was frozen for later analyses. The biomass on the filter was washed 4 times with 5 mL  of 0.9% (w/v) NaCl solution, separated from the filter, transferred to a microtube, and freeze-dried. The growth of 4+ and NO3−  in filtrates were assessed by colorimetric methods using an autoanalyzer . [NO3−] was not detected in any samples (the detection limit of 0.1 μM for the sum of NO2− and NO3−). The concentrations of dissolved organic C ([DOC]) and total dissolved N ([TDN]) were measured using a TOC analyzer with a TN unit . The concentration of dissolved organic N ([DON]) was calculated as follows: (DON)=(TDN)–(NH4+). The C:N ratio of medium (DOC/TDN) was calculated as DOC/TDN=(DOC)/(TDN).The concentrations of NHinitial)×100, where  is the concentration of TDN at 0 h and (biomass N) is the concentration of the biomass N. Microbial NUE was calculated as ([DONconsumption]–[NH4+production])/(DONconsumption), where (DONconsumption) represents the difference between initial DON and DON concentrations over the course of the experiment, and (NH4+production) is the difference between NH4+ concentrations and vs. initial NH4+ measured.The recovery % of N during the incubation was calculated as follows: Recovery N (%)=([TDN]+[biomass N])/ were measured using an elemental analyzer  interfaced with an isotope ratio mass spectrometer . Calibrated in-house standards, such as DL-alanine (δ15N=−1.7‰), glycine (δ15N=10.0‰), and L-histidine (δ15N=−8.0‰), were used to obtain the calibration curve to correct the measured isotopic values. The average standard deviations for a replicate analysis of individual samples were ±0.4‰ for δ15N after blank corrections. The biomass C/N (mol mol−1) was calculated as biomass C/N=(biomass C)/(biomass N).The concentrations of biomass carbon ([biomass C]) and (biomass N) and the δ15N values of NH4+ (δ15N-NH4+) were measured for samples with CN5 and CN10 only after 48 h with sufficient concentrations of (NH4+) that allowed for the assessment of δ15N-NH4+. δ15N-NH4+ was measured using the methods described by Koba et al.  glass fiber filter  using the diffusion method (4+ was then oxidized to NO3− using persulfate (15N of converted NO3− from NH4+ was measured using the denitrifier method (Pseudomonas aureofaciens (ATCC#13985), which lacks nitrous oxide reductase activity, was used to convert NO3− into N2O gas before the isotope analysis. The isotopic standards, USGS25 (δ15N=−30.4‰), USGS26 (δ15N=53.7‰), and IAEA N2 (δ15N=20.3‰), were used for data calibration. The average standard deviations from the replicate analysis of individual samples were large (±1.4‰) for δ15N after blank corrections, and this was attributed to samples with low (NH4+) (CN10 at 48 h). The average standard deviations from the replicate analysis were ±0.1‰ without these low (NH4+) samples after blank corrections.The δa et al. . In brien method , and conrsulfate . The δ15r method  with an 15N of glycine at 0 h (δ15N-glycine) and δ15N of TDN (δ15NTDN) were measured via the denitrifier method, in which total N was digested to NO3− using the persulfate method (15N=−1.7‰), glycine (δ15N=10.0‰), and L-histidine (δ15N=−8.0‰) were used for the calibration. The average standard deviations for the replicate analysis of individual samples were ±0.2‰ for δ15N after blank corrections. Δ15N was calculated as the difference between δ15N-biomass and δ15N-glycine (Δ15N=δ15N-biomass–δ15N-glycine).The δe method . Calibra4+) because of the inaccurate assessment of high (NH4+), and we were unable to calculate δ15N-DON with the concentrations and δ15N data of TDN and NH4+. On the other hand, the concentration of Glyused, glycine utilized by A. oryzae, (Glyused), was calculated as (Glyused)=(biomass N)+(NH4+). The δ15N of Glyused (δ15N-Glyused) was then calculated with the concentrations and δ15N values of biomass N and NH4+ as δ15N-Glyused= δ15Nmicrobe×fmicrobe+δ15N-NH4+×fNH4+ and fmicrobe+fNH4+=1, where fNH4+ is the ratio of the concentration of initial NH4+ to TDN and fmicrobe= 1–fNH4+. δ15N-Glyused was not calculated in CN30, CN50, and CN100 because the concentrations of NH4+ were too low to measure δ15N-NH4+. The standard deviations of δ15N-Glyused were evaluated using the Monte Carlo method described by Koba et al.  was lower than /(TDN) and biomass C/N on each d in all treatments, between δ15N-biomass at 96 h and initial δ15N-glycine (0.1±0.3‰) in all treatments, and between δ15N-Glyused in CN5 and CN10 and initial δ15N-glycine (0.1±0.3‰) on each d. The paired Student’s t-test was used to clarify whether concentrations or δ15N values were significantly different between δ15N-biomass at 24 and 96 h in all treatments, and between δ15N-NH4+ and δ15N-biomass on each d in CN5 and CN10. Pearson’s correlation test was used to examine the relationship between (DOC)/(TDN).Statistical analyses were conducted using R software . An α level of 0.05 was considered to indicate significance. The Student’s A. oryzae grew , while less than 2% of consumed DON was mineralized to NH4+ in other treatments (P<0.001). On the other hand, DOC/TDN was significantly higher than biomass C/N in CN30, 50, and 100 (P<0.01). Moreover, biomass C/N in CN50 and CN100 increased to 21.3±2.5 and 26.6±1.2, respectively, values that were significantly higher than the C/N of soil fungi (e.g. in lipids [47]), thereby increasing C/N. Another possibility is that the microbial biomass incorporated the degraded necromass at low N concentrations, resulting in the high C/N observed in the total biomass.As zae grew , the majzae grew , therebyzae grew . Large aeatments . Nitrateeatments ; P<0.001 and 100 ; P<0.01.il fungi  when incil fungi . Althoug4+ production). Between 24 and 72 h, underestimated biomass N was between 36 and 60%, while only 10–12% of biomass N was underestimated at 96 h in CN30, CN50, and CN100. We measured biomass N at 96 h at the end of the incubation more accurately than at other time points because we collected all of the microbial biomass left in media using large membrane filters , while we collected microbial biomass with a small ratio for 5 mL medium solution using the 5-mL pipette until 72 h. Since DON was nearly consumed by 48 h in CN30 and by 24 h in CN50 and CN100  from 24 to 72 h in CN30, CN50, and CN100 . δ15N-biomass at 96 h was significantly higher than δ15N-glycine (0.1±0.3‰) (P<0.001 and P<0.001), and Δ15N was 3.1±0.5 and 2.7±0.2‰ in CN5 and CN10, respectively. On the other hand, in CN30 and CN100, δ15N-biomass was not significantly different between 24 h (−0.2±0.1 and −0.3±0.9‰) and 96 h (0.4±0.2 and 0.5±0.0‰) (P=0.076 and P=0.279). In CN50, δ15N-biomass was significantly different between 24 h (−0.1±0.3‰) and 96 h (0.6±0.1‰) (P=0.037); however, this difference (0.7±0.2‰) was markedly smaller than that observed for CN5 and CN10. In CN30, CN50, and CN100, δ15N was not significantly different between glycine and biomass N at 96 h , and Δ15N  was smaller than those for CN5 and CN10.δ.6±0.1‰) , P=0.03715N-NH4+ was below the limit of quantification in CN30, CN50, and CN100 (15N-NH4+ was significantly lower than δ15N-biomass (15N-Glyused and δ15N-glycine (0.1±0.3‰), significant differences were observed at 48 and 72 h in CN5 and at 48 h in CN10 (15N-Glyused and δ15N-glycine (average of δ15N-Glyused–average of δ15N-glycine) remained small at 48, 72, and 96 h .δnd CN100 . In CN5 -biomass ; the dif in CN10 . However15N  and CN10 , with N functioning as more of an excess nutrient and, thus, being more rapidly mineralized from organic substrates. The present results confirmed these hypotheses; the mineralization (excretion of NH4+) of DON by A. oryzae in CN5 and CN10   . A smallss of 30 . In contand CN10 , with li-Glyused . These r-Glyused . The neg001, 15N , 41, supial δ15N .15N-NH4+ by A. oryzae may decrease, δ15N-biomass may, in principle, decrease δ15N in the microbial biomass (−14.1±0.8‰ [60]); however, this was not the case in the present study. Although DON and NH4+ absorption both occur in ecosystems , while δ15N-biomass and δ15N-NH4+ approached the δ15N of substrates in the later stages (after 50 h) in CN5 media. These findings are in contrast to the present results showing that δ15N-biomass increased over time , such that δ15N-biomass decreased and δ15N-NH4+ approached the δ15N of substrates. On the other hand, in our experiment, NH4+ re-assimilation did not occur in the later stages when A. oryzae actively mineralized organic N. Furthermore, the glycine cleavage system may have caused the enrichment of δ15N-NH4+ and the depletion of δ15N-biomass in the study by Collins et al. , with a constant and large difference (3–4.8‰) between the δ15N of animals and substrates (15N enrichment; relatively constant 15N enrichment and NUE for animals and variable 15N enrichment and NUE for microbes.The present results on and Δ15N , indicatbstrates , 52, 57.15N-SOM patterns in the field; however, further research is needed to test this hypothesis under natural ecosystem settings. δ15N-SOM has been shown to markedly change with soil depth, with more substantial 15N enrichment being observed from shallow to deep soil layers (15N of SOM is also affected by δ15N in the microbial biomass, which is controlled by microbial NUE. Further research is warranted to understand the control of soil microbial stoichiometry on δ15N-SOM, and the present results suggest a hypothesis that has the potential to link microbial N demands to δ15N, thereby offering a non-intrusive and integrative tool through which to understand nutrient limitations in soil decomposer communities.These results imply similar control over δl layers , 48, 50.l layers , 49, the


E configuration at the C=C double bond. The dihedral angle between the benzene ring and the prop-1-enyl group is 25.4 (3)°. In the crystal, molecules are linked via N—H⋯F hydrogen bonds, forming inversion dimers which are linked into ribbons along the b axis by C—H⋯N hydrogen bonds. The ribbons are linked by N—H⋯π and C—H⋯π interactions, generating a three-dimensional network.The molecule adopts an  9H8F3N, adopts an E configuration with respect to the C=C double bond. The dihedral angle between the benzene ring and the prop-1-enyl group is 25.4 (3)°. In the crystal, molecules are linked via pairs of N—H⋯F hydrogen bonds into inversion dimers with an R22(16) ring motif. The dimers are linked by C—H⋯N hydrogen bonds, forming a ribbon structure along the b-axis direction. The ribbons are linked by N—H⋯π and C—H⋯π interactions, generating a three-dimensional network.The molecule of the title compound, C Their derivatives exhibit strong fluorescence in aqueous solution and so can be utilized as ion-probes and in bio-imaging . Since aniline derivatives with 2,4-bis have been used as fluorogenic substrates for dipepeptidyl peptidase-4 °. The C5—C10—C11—C12 and C9—C10—C11—C12 torsion angles are −158.9 (3) and 24.6 (4)°, respectively. The bond lengths and angles in the title compound are normal and agree with those in other trifluoropropenylaniline compounds  chain motif along the b-axis direction. The ribbons are linked by N—H⋯π and C—H⋯π interactions -3,3,3-trifluoroprop-1-enyl and 2-aminophenyl-1-enyl fragments, respectively. Of these structures, those that resemble the title compound are 4-[2-phenyl]morpholine -2-chloro-3,3,3-trifluoroprop-1-enyl]phenyl}acetamide -1,4-dipiperidino-2,5-bisbenzene  and 2,2′-bipyridyl (2.0 mmol) were placed in a flask. To the flask were added anhydrous DMF (6.0 ml) and (E)-trimethyl-silane (2.0 mmol), and the mixture was stirred at 353 K. After the reaction mixture had been stirred for 4 h, it was poured into ice–water. The mixture was extracted with CH2Cl2, and the organic layer was dried over anhydrous MgSO4. After the solid had been filtered off, the solvent was removed in vacuo, and the residue was purified by silica gel column chromatography to give the product in 68% yield. Colourless single crystals were obtained by recrystallization from an ethyl acetate–hexane  solution (m.p. 321–322 K). 1H NMR (CDCl3) δ: 3.81 , 6.13 , 6.72 , 6.80 , 7.18 , 7.24 , 7.29 . 13C NMR (CDCl3) δ: 116.6 , 116.8, 119.2, 119.4, 123.6 , 127.9, 130.9, 133.3 , 144.8. 19F NMR (CDCl3) δ: 12.07 . MS m/z 187 (M+), HRMS calculated for C9H8F3N 187.1617 (M+), found 187.0603.The title compound was prepared by a modification of a reported procedure (Omote Uiso(H) = 1.2Ueq(C). One outlier  global, I. DOI: 10.1107/S2056989018012756/is5499Isup2.hklStructure factors: contains datablock(s) I. DOI: 1866671CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


II atom in the mononuclear title complex has an almost regular octahedral N4O2 coordination geometry. In the crystal, the complex molecules are linked in a three-dimensional network through strong O—H⋯N hydrogen bonds.The Ni 2·6H2O, produced the title complex, [Ni(NCS)2(C12H8N4O)2(H2O)2]. The NiII atom is located on an inversion centre and is octahedrally coordinated by four N atoms from two ligands and two pseudohalide ions, forming the equatorial plane. The axial positions are occupied by two O atoms of coordinated water molecules. In the crystal, the molecules are linked into a three-dimensional network through strong O—H⋯N hydrogen bonds. Hirshfeld surface analysis was used to investigate the intermolecular interactions in the crystal packing.The reaction of 2,5-bis(pyridin-4-yl)-1,3,4-oxadiazole (4-pox) and thiocyanate ions, used as co-ligand with nickel salt NiCl The Ni1x, 1 − y, 2 − z] are also observed between pyridine rings (N4/C8–C12) coordinated to adjacent metal centres.In the crystal, the molecules are linked through strong O—H⋯N hydrogen bonds Table 1, forminget al., 2007CrystalExplorer17.5  analysis  also have a nearly symmetrical distribution of points, with thick edges at de + di ≃ 3.1 Å. The H⋯H contacts  appear in the central region of the fingerprint plot with de = di ≃ 1.0 Å. The S⋯H/H⋯S contacts  indicate that the interatomic separations are greater than the sum of the van der Waals radii, suggesting they have a limited influence on the molecular packing. The C⋯C contacts  are a measure of the π–π stacking interactions and have an arrow-shaped distribution of points with the tip at de = di ≃ 1.7 Å. π–π Interactions are indicated by adjacent red and blue triangles in the surface mapped over shape-index  crystal in the range 500–4000 cm−1 in addition to another sharp and strong band at 2083 cm−1, attributable to the water molecules  where M = CdII . After 10 min of reaction, the precipitate obtained was filtered and washed with distilled water and dissolved in 15 ml of DMF. After one month of slow evaporation of the solvent, the obtained green single crystals were washed with water and dried under vacuum (80%). These crystals were used as isolated for single crystal X-ray analysis. Analysis calculated for C26H20N10NiS2O4. C, 47.36; H, 3.06; N, 21.24; S, 9.73; found: C, 47.51; H, 3.13; N, 21.29; S, 9.59. IR–ATR (cm−1): 3055 (w), 1618 (m), 1569 (m), 1551 (m), 1488 (m), 1418 (m), 1330 (w), 1279 (w), 1237 (w), 1213 (w), 1125 (w), 1097 (w), 1062 (m), 1019 (w), 1008 (m), 972 (w), 842 (s), 750 (m), 728 (s), 715 (s), 697 (m). UV–vis : 274 (28920).The 2,5-bis(4-pyridin-4-yl)-1,3,4-oxadiazole (4-pox) ligand was synthesized as described previously  set to 1.5 Ueq(O). All other H atoms were located in a difference-Fourier map and refined as riding, with C—H = 0.93 Å and with Uiso(H) = 1.2Ueq(C).Crystal data, data collection and structure refinement details are summarized in Table 410.1107/S2056989019008727/rz5259sup1.cifCrystal structure: contains datablock(s) I. DOI: 10.1107/S2056989019008727/rz5259Isup2.hklStructure factors: contains datablock(s) I. DOI: 1911306CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


Many compounds were shown to exhibit a number of different polymorphic structures , although the bulk material studied by solid‐state NMR spectroscopy often contained just one major polymorph. For the unoxidised heterocycles, the presence of weak intermolecular J couplings was also investigated by DFT calculations.A series of P−E‐containing heterocycles  with aromatic backbones were synthesised and characterised by single‐crystal and powder XRD, microanalysis and mass spectrometry. Solution‐ and solid‐state  The J values calculated by periodic DFT confirm that a larger interaction is expected for 13, as a consequence of the different packing motifs of the two compounds. To understand the effect of these unusual interactions on the stability, conformation and solid‐state packing of the compounds, the series has been extended here, both to include a different chalcogen (S) and to vary the oxidation state of the P atom, potentially precluding this atom's participating in additional interactions. A similar approach was taken in previous work by Woollins and co‐workers to monitor the resulting molecular distortion and effect on the though‐space interactions between the peri positions for compounds of the form Nap[P(E′)(Ph2)(ER)] .In 2015, Sanz Camacho et al.δiso for 77Se and 31P were compared for solution and solid‐state samples, and differences were related to conformational changes.Herein, we present a study on the properties and structural features of a series of new chalcogen–phosphorus heterocycles. These compounds exhibit extensive polymorphism, which was investigated not only by single‐crystal XRD, but also by studying the bulk material by solid‐state NMR spectroscopy and powder XRD (PXRD). Structural characterisation was completed by using solution‐state NMR spectroscopy, IR spectroscopy, mass spectrometry and elemental analysis. The isotropic chemical shifts cd]1,2‐dithiole isopropylphosphine (1) and naphtho1,2‐dithiole tert‐butylphosphine (9) were prepared according to reference cd]1,2‐diselenole isopropylphosphine (5) and naphtho1,2‐diselenole tert‐butylphosphine (13) have already been described in recent work.2O2 at room temperature or 0 °C in air. Yields and compound numbering are given in Table Scheme III compounds 1, 5, 9 and 13 are stable in solution for up to about one week, after which decomposition to the NapE2 precursor is observed, rather than oxidation to the PV=O‐containing system. In the solid state, these compounds are stable upon exposure to air for at least 12 months. The oxidised heterocycles have high stability upon exposure to air in the solid form, but decompose quickly in solution to the E−E analogue (rather than the corresponding unoxidised heterocycle).The P1–16 show some interesting features (see Table S1 in the Supporting Information). Six of the compounds  adopt different polymorphic forms . There are a number of isomorphous structures . The basic molecular geometry is an open envelope conformation for the C3E2P ring with hinge angles between the C3E2 and E2P planes of about 55°. The molecules adopt two conformations. The substituent(s) on phosphorus  can be approximately perpendicular or collinear with the naphthalene plane. This can be thought of as similar to the axial/equatorial arrangements in a cyclohexane ring. The most obvious common feature is that all of the structures where X=O have the P=O bond approximately perpendicular to the naphthalene ring plane. Interestingly, in most cases , for the heavier chalcogens, when R=iPr the structures have the P−C bond perpendicular to the naphthalene plane, whereas the tBu analogues have the P−C bond collinear.The single‐crystal X‐ray structures of 31P and 77Se NMR parameters for all compounds are given in Table 31P{1H} NMR spectra of all the complexes exhibit singlets with satellites due to 1J(P−Se) or/and 1J(P=Se) coupling for the selenium compounds. The 31P{1H} NMR spectra of the sulfur compounds appear to be the most deshielded of the series, in agreement with previous literature.tBu compounds are deshielded in comparison to the corresponding iPr compounds, with the exception of 10. The nature of the X substituent has an influence on the 1J(31P−77Se) coupling constant, which decreases in the order O≥Se>S. The R group also has an impact on the 1J(31P−77Se) coupling, with tBu>iPr. However, the opposite trend is observed for the 1J(31P=77Se) coupling, which decreases in the order iPr>tBu. The 77Se isotropic chemical shifts are also affected by the nature of X, with δiso(77Se) decreasing for the iPr compounds in the order Se≥S>O. However, for the tBu compounds, the S‐containing system is the most deshielded. In contrast to the 31P{1H} NMR spectra, δiso(77Se) are, as expected, more deshielded for the iPr analogues.The 5 and 13 have previously been characterised and studied by 31P and 77Se solid‐state NMR spectroscopy. The presence of an intermolecular J coupling between 31P in one molecule and 77Se in an adjacent molecule was observed for 13 .77Se NMR spectrum of 5 (where R=iPr), as the different packing motifs of the two result in a greater distance between the two atoms and, therefore, a smaller coupling. In contrast, the sulfur compounds (1 and 9) have not been previously reported. Compound 1 exhibits three different polymorphs. The asymmetric units and packing motifs for these three structures and the single polymorph observed for 9 are shown in Figure 1 differ not only in their asymmetric units, but also in the packing motifs. Polymorph 1 b has four molecules in the asymmetric unit. Each molecule is more isolated and stacks in an antiparallel arrangement along the c axis. Both 1 a and 1 c have only two molecules in the asymmetric unit, although the extended packing is different between the two polymorphs. In 1 a, the molecules form triangles that stack along the c axis, whilst in 1 c the chains of molecules stack perpendicularly along the a axis. To determine which polymorph(s) were present in the bulk sample, a PXRD pattern was collected and compared to those simulated for each polymorph individually. The experimental and simulated PXRD patterns are shown in Figure S2.1 of the Supporting Information. The relative intensities and position of the reflections in the experimental pattern agree with those simulated for 1 b.Compounds 31P MAS NMR spectra of 5 and 13 have previously been reported by Sanz Camacho et al., in 2016.13, with a 31P J‐resolved spectrum showing an intermolecular homonuclear 31P–31P coupling (≈88 Hz). Interestingly, this interaction was between P species that are crystallographically equivalent but are rendered magnetically inequivalent in a fraction of the molecules by heteronuclear coupling to 77Se. This interaction was not resolved for 5  as a consequence of the different packing motifs.The 31P MAS NMR spectrum of 1, shown in Figure S2.2 of the Supporting Information, contains three isotropic resonances, each with a significant sideband manifold as a result of CSA. The three resonances exhibit an integrated intensity ratio (including sidebands) of about 1:1:2, suggesting that the resonance at lowest shift might correspond to two P atoms with a very similar environment. This suggests the presence of four molecules in the asymmetric unit and hence the presence of polymorph 1 b in the bulk sample, in agreement with the PXRD results. Unfortunately, due to the nature of the sample of 9 (a sticky solid), it was not possible to study the bulk compound by solid‐state NMR spectroscopy or PXRD.The 1, 5, 9 and 13 exhibit different crystal packing motifs, resulting in different internuclear chalcogen–P distances, as shown in Figure 13 this leads to the observation of intermolecular (31P–77Se and 31P–31P) J couplings. Figure 31P–33S couplings would not be present .Despite their chemical similarity, 13 also leads to the observation of 31P–31P homonuclear intermolecular coupling, as discussed above. Table JPP coupling constants predicted for compounds 1, 5, 9 and 13 by periodic DFT. Values of J are generally larger for shorter P−P distances, though, notably in 5, a larger J value is predicted for the longer P−P distance owing to the relative orientation of the paired molecules. A significant through‐space J coupling is computed only for 13, and this reflects the shorter internuclear P−P distance and particular packing arrangement found for this compound.The intermolecular proximity in 2, 6, 10 and 14) were the most difficult to synthesise, due to their tendency to decompose, and only a single crystal structure was obtained for each compound. The asymmetric unit and packing motifs are shown in Figure 6 and 14, respectively, and in Figures S3.1 and S3.2 in the Supporting Information for 2 and 10. The PXRD patterns for bulk samples of 2 and 10  are in good agreement with those simulated from the structural models derived from single‐crystal diffraction. The 31P MAS NMR spectra (Supporting Information) also confirm the presence of one distinct P species. For selenium compounds 6 and 14, the simulated PXRD patterns do not agree with those obtained experimentally for the bulk material, as shown in Figure The four P=O oxidised heterocycles . This can be confirmed by the application of 31P decoupling, as in Figure 31P MAS NMR spectrum , as seen in Figure 14. However, this clearly demonstrates the need for multiple characterisation techniques when synthesising new materials.Solid‐state 3, 7, 11 and 15. The asymmetric units, crystal packing motifs, PXRD data and 31P MAS spectra of 7, 11 and 15 are shown in Section S4 of the Supporting Information. Only one polymorph is observed for each of 7, 11 and 15, with two, two and one distinct molecules in their respective asymmetric units. Their PXRD patterns are also in good agreement with those simulated from the structural models derived from single‐crystal XRD. For 3, three different polymorphs are observed; 3 a and 3 c have two distinct molecules in the asymmetric unit, while only one is present for 3 b. The difference in crystal packing motifs for the three polymorphs is shown in Figure 31P MAS NMR spectrum of 3 shows three resonances, each with different intensity, as shown in Figure 3 a and 3 c and one for 3 b), although the chemical similarity of the environments may well result in some overlap of the spectral resonances.The four P=S oxidised heterocycles are 31P MAS NMR spectra (isotropic region only) are shown in Figure The presence of significant sideband manifolds for each resonance hinders the accurate determination of relative signal intensities, and so a second spectrum was acquired at a faster MAS rate of 55 kHz (with a 1.3 mm rotor), as shown in Figure 3 a , which suggests loss of 3 b and 3 c from the bulk material. This suggests that the two resonances observed at about 62 and about 66 ppm correspond to the two distinct P species in 3 a, while the signal at about 64 ppm most likely results from the overlap of three signals (from 3 b and 3 c).The changes appear to be irreversible, with no further change in the spectrum as the temperature is reduced back to 273 K. PXRD measurements also confirmed a change in the bulk sample after the VT NMR experiments, as shown in Figure 4, 8, 12 and 16). The asymmetric units, crystal packing motifs and PXRD data for 4, 8 and 16 are shown in Section S5 of the Supporting Information. The two polymorphs of 4 have one (4 a) and two (4 b) distinct molecules in the asymmetric unit and have very different predicted PXRD patterns. Comparison of these to the experimental PXRD pattern suggests that 4 a makes up the majority of the bulk material. The 31P MAS and 77Se CP MAS NMR spectra  exhibit one 31P and one 77Se species, in agreement with the structure of 4 a, together with low‐intensity resonances that suggest 4 b is present only in a very small amount. The 1J(31P=77Se) coupling can be resolved in both the 31P and 77Se spectra (≈779 Hz). Upon application of 31P decoupling, the 77Se spectrum displays a very small residual coupling, most probably as a result of the low decoupling power that can be applied and the magnitude of the CSA present. The two polymorphs of 8  have one (8 a) and two (8 b) distinct molecules in the asymmetric unit, again with very different predicted PXRD patterns. Comparison with the experimental PXRD pattern suggests that the bulk material is primarily 8 a. The 31P MAS and 77Se CP MAS NMR spectra  show a single P site and three distinct Se sites, as expected, with 1J(31P−77Se)=380–390 Hz and 1J(31P=77Se)=749 Hz.Two polymorphs were identified for each of the P=Se oxidised compounds (16 have one (16 a) and two (16 b) molecules in their asymmetric units but have very similar predicted PXRD patterns . The 31P MAS NMR spectrum  contains two closely spaced resonances with similar intensities, suggesting that either 1) 16 b makes up the majority of the bulk material, with 16 a present only in small amounts, or 2) the two P atoms in 16 b have an identical chemical shift and there is a 2:1 ratio of 16 a and 16 b. The 77Se CP MAS NMR spectra  confirm that six Se sites are most likely to be present, again in support of the presence of only 16 b in the bulk sample, although resonances overlap even under 31P decoupling. Final confirmation was obtained by acquiring the 77Se CP MAS NMR spectrum of a powder obtained from a single crystal whose structure was determined by single‐crystal XRD to be 16 b. This was in excellent agreement with the spectra obtained on the bulk material.The two polymorphs of 12 a and 12 b both contain two molecules in the asymmetric unit, although the two structures have different crystal packing motifs, as shown in Figure 12 a resulted from crystallisation from dichloromethane/ethanol, while 12 b crystallised from dichloromethane/methanol. The very similar simulated PXRD patterns  couplings of about 835 and about 826 Hz. Additional peaks are also observed as shoulders on each resonance, possibly arising from the second polymorph. The application of 31P decoupling suggests that four Se sites are present, although the low decoupling power available does limit resolution. From this, it seems likely that a mixture of both polymorphs is present in the bulk sample.Polymorphs s Figure  b make im Figure  a, possim Figure  b exhibi31P , lower δiso is found for unoxidised heterocycles, with an increase on moving from P=O to P=Se and, finally, P=S. For compounds in which E=Se (circles), unoxidised heterocycles again have the lowest δiso, with an increase upon =Se oxidation. Larger, but similar, shifts are seen for P=O and P=S compounds. The range of shifts is much greater when E=S than when E=Se. However, Figure 31P species in the same material . There is a good correlation between solution‐state and solid‐state 77Se chemical shifts  all lie slightly above the ideal correlation.Figure P Figure  a, the ral Table  and in dal Table . Three ps Figure  b. The sc‐Napc dihedral angle  is approximately 0°, and B, in which the X‐P‐Ec‐Napc dihedral angle is about 180°. Interestingly (and as shown in Section S1 of the Supporting Information), all P=O compounds adopt conformation A. For compounds in which R=iPr (1–9), only the two P=O compounds (2 and 6) adopt this conformation. When R=tBu, all S compounds adopt the A conformation (with the exception of unoxidised heterocycle 9), while the corresponding Se compounds exhibit the B conformation, with the exception, as stated above, of P=O compound 14. For compounds that exhibit polymorphism, all polymorphs adopt the same conformation, with variations only in the crystal packing. It would seem, therefore, that some of the differences in chemical shift observed between solid‐ and solution‐state NMR spectra, may result from the fact that two distinct conformations are possible in the solid state , as opposed to the more dynamic conformational averaging that likely exists in solution. To explore the effect of conformation on the NMR parameters, calculations were performed for (optimised) isolated molecules of 12, 14 and 16 in both A and B conformations. Values of the isotropic shielding σisocalcdand span Ωcalcd for 31P and 77Se are given in Table 31P, small differences in σisocalcd(between 1 and 17 ppm) are seen, while for 77Se, more significant differences in σisocalcdare observed (135–323 ppm for P−Se and 249–345 ppm for P=Se).As discussed earlier . Differences in 77Se Ωcalcd are more significant, with changes of 300–600 ppm and about 1100 ppm, for P−Se and P=Se species, respectively. Although this information is lost in solution‐state NMR measurements, owing to the rapid tumbling motion of the molecules, Table Ω  can be used to indicate the molecular conformation adopted in the bulk powder. Figure Ω against δiso for 31P and 77Se, respectively.Clearly there are more significant variations in Ω, κ, and the principal components of the shielding tensors iiδ are given in Tables S6.1 and S6.2 of the Supporting Information. There is a clear distinction in 31P Ω . Although the 77Se Ω value  and the B conformation (Ω≈200 ppm), in excellent agreement with the DFT calculations. As shown in Figure S7.1 of the Supporting Information, there is also good agreement between 1J(31P−77Se) in solution and in the solid state. There is a clear distinction in the magnitude of the J coupling seen in the solid state for P−Se species in unoxidised (≈270–300 Hz) and oxidised (≈400 Hz) heterocycles, although the ranges have a little more overlap in solution. The J coupling for P=Se species is much greater in both solution and in the solid state, as expected. There are no significant differences in the magnitude of any J couplings between molecules adopting the A or B conformations in the solid state.Values of Ω Figure  b for coe Figure  c for P−31P and 77Se solid‐state NMR spectra are very sensitive to changes in the local environment, even for polymorphs with structures that are quite similar, as seen for 12. Furthermore, solid‐state NMR spectroscopy provides a quantitative determination of the fractions of each polymorph in the bulk sample from the relative intensities of the resonances. For the heterocycles studied here, the tendency to exhibit polymorphism increases as the relative content of more polarizable atoms increases, for example, the two compounds that contain three Se atoms, 8 and 16, exhibit the greatest number of polymorphs.The presence of extensive polymorphism in a series of heterocycles was confirmed by using the complementary techniques of single‐crystal and powder XRD, as well as solid‐state NMR spectroscopy. It is clear that although single‐crystal XRD is essential to understand the detailed structure of each polymorph, methods such as powder XRD and solid‐state NMR spectroscopy are required to determine the fraction of each of these in the bulk simple. PXRD can provide a good indication of the polymorphs present in many cases, although for some compounds the PXRD patterns of the different polymorphs are very similar. In other cases, the complex mixture of polymorphs in the bulk material also hinders detailed analysis of the PXRD patterns. In contrast, δiso and Ω), particularly for 77Se, are very sensitive to the conformation adopted. This was also confirmed by DFT calculations of NMR parameters for isolated molecules of 12, 14 and 16 adopting the two different conformations. Comparison of the isotropic chemical shifts in solution and in the solid state show differences for some compounds, and the DFT calculations confirmed that these differences do not generally arise as a result of the crystal packing and intermolecular interactions, but probably as a result of the different conformation found in the solid state and the rapid averaging that likely occurs in solution. Intermolecular interactions are observed  for the unoxidised compounds, but these are limited in the oxidised analogues with occupation of the P lone pair.Two different arrangements of the P=X bond (and consequently the P−C bond) were observed for the oxidised compounds, and it was shown that the NMR parameters , atmospheric pressure chemical ionisation (APCI+), atmospheric solids analysis probe (ASAP+) and nano‐electrospray (NSI) mass spectra were carried out by the EPSRC National Mass Spectrometry Service, Swansea. 1H and 13C solution‐state NMR spectra were recorded with a Bruker Avance 400 MHz or a Bruker Avance 300 MHz spectrometer with chemical shifts referenced to residual solvent peaks. 77Se and 31P solution‐state NMR spectra were recorded with a Jeol GSX 270 MHz spectrometer with chemical shifts referenced to external (CH3)2Se and 85 % H3PO4, respectively. Assignments of 13C and 1H NMR spectra were made with the help of 1H–1H COSY, 1H–13C HSQC and 1H–13C HSBC experiments. The naphtho1,2‐dithiole and naphtho1,2‐diselenole precursors were prepared by literature procedures.5 and 13 have been reported elsewhere.All syntheses were carried out under an oxygen‐ and moisture‐free nitrogen atmosphere by using standard Schlenk techniques and glassware. Reagents were obtained from commercial sources and used as received. Dry solvents were collected from an MBraun solvent purification system. Elemental analyses were performed by Stephen Boyer at the London Metropolitan University. IR spectra were recorded for solids as KBr discs and oils on KCl plates in the range 4000–300 cmm solution of lithium triethylborohydride (superhydride) in THF  was added dropwise to a solution of naphtho1,2‐dithiole  in THF (100 mL). The mixture was stirred at room temperature for 15 min, after which a solution of dichloroisopropylphosphine  in THF (10 mL) was added dropwise to the mixture. The resulting mixture was heated to about 66 °C and left overnight. After the solvent was removed in vacuo, the reaction mixture was extracted with hexane (125 mL), washed with distilled water (200 mL) and the organic layer dried with magnesium sulfate and concentrated under reduced pressure. Column chromatography on silica gel (hexane) was performed to afford the purified target compound as a white solid. Crystals suitable for X‐ray diffraction were grown from hexane . IR (KBr disc): ν˜=2951w, 2916, 2956w, 1548s, 1494s, 1463w, 1360s 1317w, 1232w, 1203vs, 1192s, 1148w, 1082w, 1030s, 888w, 868w, 813vs, 755vs, 639s, 546w, 533w, 508s, 498 cm−1 s; 1H{31P} NMR : δ=7.8  7.6  7.4  1.9  1.1 ppm ; 13C{1H} NMR : δ=135.4  131.0  129.6  127.7  125.7  124.4  28.8  18.9 ppm ; 31P{1H} NMR : δ=4.72 ppm (s); MS (APCI+): m/z (%) 265.02 (100) [M+H]+; elemental analysis calcd (%) for C13H13PS2 (264.35): C 59.07, H 4.96; found: C 59.21, H 4.87.A 1 1  in dichloromethane (40 mL) and stirring continued for 5 h. Removal of the volatile substances afforded a pale yellow solid. Crystals suitable for X‐ray diffraction were grown by layering a hexane solution of 2 with dichloromethane . M.p. 114–118 °C; IR (KBr disc): ν˜=2966w, 2362w, 1546w, 1460w, 1365w, 1213vs, 1035s, 877s, 820vs, 758vs, 667s, 571vs, 554vs, 535 cm−1 s; 1H{31P} NMR : δ=7.8 , 7.6 , 7.4 , 2.3 , 1.4 ppm ; 13C{1H} NMR  δ=136.2 , 132.3 , 130.1 , 127.0 , 126.7 , 126.4 , 34.8 , 15.5 ppm ; 31P{1H} NMR : δ=52.0 ppm (s); MS (APCI+): m/z (%) 281.0223 (56) [M+H]+, 220.9647 (100) [C10H6PS2], 189.9909 (67) [C10H6PS]+; elemental analysis calcd (%) for C13H13OPS2 (280.34): C 55.7, H 4.7; found: C 55.5, H 4.7.Hydrogen peroxide  was added to solution of 1  and sulfur flowers  in toluene (50 mL) was heated at 110 °C for 48 h. The resulting solution was allowed to cool to room temperature and, after removal of the volatile substances, column chromatography on silica gel hexane/CH2Cl2 1:1 was performed to afford a pale pink solid. Crystals suitable for X‐ray diffraction were grown from diethyl ether . M.p. 175–177 °C; IR (KBr disc):ν˜=2965w, 2922w, 2862w, 1546s, 1494w, 1447w, 1361w, 1326w, 1262s, 1200s, 1092vs, 1031vs, 878w, 817vs, 757vs, 715vs, 614vs, 566s, 483 cm−1 w; 1H{31P} NMR : δ=7.8 , 7.6 , 7.4 , 2.3 , 1.4 ppm ; 13C{1H} NMR : δ=136.1 , 131.2 , 130.5 , 128.1 , 126.5 , 35.6 , 15.5 ppm ; 31P{1H} NMR : δ=67.2 ppm (s); MS (APCI+): m/z (%) 296.9992 (100) [M+H]+.A solution of 1  and elemental selenium  in toluene (50 mL) was heated at 110 °C and left overnight. The resulting solution was allowed to cool to room temperature and was filtered through a silica plug with hexane (250 mL) and dichloromethane (250 mL). Removal of the volatile substances afforded a pink solid. Crystals suitable for X‐ray diffraction were grown by layering a solution of 4 in dichloromethane with hexane . M.p. 191–197 °C; IR (KBr disc): ν˜=2964s, 2922w, 1949w, 1546w, 1493w, 1443w, 1360w, 1261vs, 1202s, 1094vs, 1030vs, 877w, 814vs, 819.1vs, 751s, 664s, 564vs, 428 cm−1 w; 1H{31P} NMR : δ=7.8 , 7.6 , 7.4 , 2.4 , 1.2 ppm ; 13C{1H} NMR : δ=136.1 , 130.8 , 130.5 , 127.8 , 126.5 , 126.3 , 35.2 , 16.0 ppm ; 31P{1H} NMR : δ=51.6 ppm =797 Hz); 77Se{1H} NMR : δ=−310.6 ppm =797 Hz); MS (APCI+): m/z (%) 343.9350 (3) [M]+, 220.9641 (100) [C10H6S2P]+, 189.9904 (82) [C10H6S2]+; elemental analysis calcd (%) for C13H13SePS2 (343.3): C 45.5, H 3.8; found: C 45.4, H 3.9.A solution of 2O2 (30 % solution in water)  was added dropwise (60 μL per 20 min) to a solution of 5  in toluene (80 mL) in an ice bath. Stirring was continued until complete consumption of the starting material, monitored by 31P NMR spectroscopy. The reaction mixture was washed with water (100 mL), and the organic layer dried with magnesium sulfate and concentrated under reduced pressure. Crystals suitable for X‐ray diffraction were grown by layering a dichloromethane solution of 6 with hexane . M.p. 79–82 °C; IR (KBr disc):ν˜=3433.6s, 2964s, 1639.5s, 1538.8s, 1460.5s, 1349.6s, 1261.6vs, 1195.6vs, 1096.5vs, 1028.6vs, 874s, 798.3vs, 754.5vs, 653.7s, 498vs, 383.9s, 340.4s, 309.2s, 294.8s, 260.2 cm−1 vs; 1H{31P} NMR : δ=7.8 , 7.7 , 7.3 , 2.37 , 1.4 ppm ; 13C{1H} NMR : δ=136.5 , 134.3 , 131.0 , 128.6 , 126.4 , 38.9 , 16.1 ppm ; 31P{1H} NMR : δ=40.4 ppm =398.4 Hz); 77Se{1H} NMR : δ=403.8 ppm =396.6 Hz); MS (APCI+): m/z (%) 376.9107 (100) [M+2 H]+, 286.8873 (67) [C10H6Se2+H]+; elemental analysis calcd (%) for C13H13OPSe2 (374.1): C 41.7, H 3.5; found: C 41.6, H 3.4.H5  and elemental sulfur  in toluene (30 mL) was heated at 80 °C for several hours. The resulting solution was allowed to cool to room temperature and then the solvent was removed in vacuo. Column chromatography on silica gel with hexane/dichloromethane (4:1) was performed to afford the purified target compound as a purple solid. Crystals suitable for X‐ray diffraction were grown by layering a dichloromethane solution of 7 in methanol . M.p. 147–150 °C; IR (KBr disc): ν˜=3424w, 2921w, 1655w, 1539s, 1488w, 1441w, 1357s, 1315w, 1192s, 1032s, 816vs, 753s, 700vs, 590vs, 482s, 384 cm−1 w; 1H{31P} NMR : δ=7.8 , 7.8 , 7.4 , 2.4 , 1.3 ppm ; 13C{1H} NMR : δ=136.3 , 132.9 , 131.2 , 128.3 , 128.0 , 126.3  39.1  15.8 ppm ; 31P{1H} NMR : δ=43.3 ppm =385 Hz); 77Se{1H} NMR : δ=438.7 ppm =385 Hz); MS (EI+): m/z (%) 391.9 (15) [M+H]+, 285.8 (100) [C10H6Se2]+, 237.9 (33) [C10H6SeP].+, 205.9 (33) [C10H6Se].+, 126.0 (32) [C10H6].+; elemental analysis calcd (%) for C13H13SPSe2 (390.20): C 40.02, H 3.36; found: C 40.14, H 3.31.A solution of 5  and elemental selenium  in toluene (30 mL) was heated to 80 °C and left overnight. The resulting solution was allowed to cool to room temperature and then the solvent was removed in vacuo. Column chromatography on silica gel with hexane/dichloromethane (4:1) was performed to afford the purified target compound as an orange‐pink solid. Crystals suitable for X‐ray diffraction were grown by layering a dichloromethane solution of 8 in methanol . M.p. 150–153 °C; IR (KBr disc): ν˜=3450w, 2962w, 2858w, 1539s, 1487w, 1438s, 1355s, 1312w, 1237w, 1191s, 1135w, 1084w, 1027s, 871w, 845w, 812vs, 750vs, 686w, 648vs, 515vs, 475vs, 424w, 374 cm−1 w; 1H{31P} NMR : δ=7.9 , 7.8 , 7.4 , 2.5 , 1.2 ppm ; 13C{1H} NMR : δ=136.3 , 132.5 , 131.3 , 128.3 , 127.9 , 126.3 , 38.2  16.3 ppm ; 31P{1H} NMR : δ=22.0 ppm =391 Hz, 1J (31P–77Se)=773 Hz); 77Se{1H} NMR ; δ=439.2 =391 Hz), −260 ppm =773 Hz); MS (APCI+): m/z (%): 438.8339 (53) [M+H]+, 360.9161 (38) [C13H13PSe2]+; elemental analysis calcd (%) for C13H13PSe3 (437.1): C 35.7, H 3.0; found: C 35.7, H 2.9.A solution of m solution of superhydride in THF  was added dropwise to a solution of naphtho1,2‐dithiole  in THF (100 mL). The mixture was stirred at room temperature for 15 min, after which a solution of dichloro‐tert‐butylphosphine  in THF (10 mL) was added dropwise to the mixture. The resulting mixture was warmed to about 66 °C and left overnight. After the solvent was removed in vacuo, the reaction mixture was extracted with hexane (125 mL), washed with distilled water (200 mL) and the organic layer dried with magnesium sulfate and concentrated under reduced pressure. Column chromatography on silica gel (hexane) was performed to afford the purified target compound as a yellowish sticky solid. Crystals suitable for X‐ray diffraction were grown from hexane . 1H{31P} NMR : δ=7.7  7.6  7.3  1.1 ppm ; 13C{1H} NMR : δ=135.2  129.8  129.1  128.3  125.8  125.5  39.6  27.4 ppm ; 31P{1H} NMR : δ=24.1 ppm (s); MS (EI+): m/z (%) 278.0 (10) [M⋅]+, 189 (100) [C10H6S2−H]+; elemental analysis calcd (%) for C14H15PS2 (278.3): C 60.4, H 5.43; found: C 60.55, H 5.35.A 1 m) was added. to a solution of 9  in dichloromethane (50 mL) The mixture was stirred overnight to give a yellowish solution. Removal of the volatile substances afforded a pale yellow solid. Crystals suitable for X‐ray diffraction were grown by layering a solution of 10 in dichloromethane with hexane . M.p. 230–236 °C (decomp); IR (KBr disc): ν˜=2962w, 2926w, 2857w, 2361w, 1546w, 1457w, 1362w, 1262w, 1206vs, 1185vs, 1146s, 883w, 822vs, 761vs, 624vs, 572vs, 510s, 488w, 406 cm−1 w; 1H{31P} NMR : δ=7.7 , 7.6 , 7.4 , 1.4 ppm ; 13C{1H} NMR : δ=136.1 , 132.5 , 130.0 , 127.1 , 126.3 , 125.6 , 40.5 , 24.8 ppm ; 31P{1H} NMR : δ=51.7 ppm (s); MS (APCI+): m/z (%) 295.0375 (100) [M+H]+.Hydrogen peroxide  and sulfur flowers  at 90 °C for 72 h. A pinkish orange solid was afforded. Crystals suitable for X‐ray diffraction were grown by layering a solution of 11 in dichloromethane with hexane . M.p. 198–203 °C; IR (KBr disc): ν˜=2957s, 2922w, 2859s, 1550w, 1495w, 1470w, 1456w, 1365w, 1324w, 1261w, 1203s, 1094br, 1015w, 883w, 813vs, 755vs, 689vs, 601vs, 551vs, 472w; 1H{31P} NMR  δ (ppm)=7.8 , 7.6 , 7.4 , 1.4 ; 13C{1H} NMR : δ=134.8 , 130.0 , 129.2 , 127.05 , 125.3 , 125.1 , 44.8 , 24.2 ; 31P{1H} NMR : δ=70.2 (s); MS (APCI+): m/z (%) 311.0145 (100) [M+H]+, 279.0425 (48) [C14H15PS2]+; elemental analysis calcd (%) for C14H15PS3 (310.43): C 54.2.1, H 4.9; found: C 53.8, H 5.0.Compound 12 was prepared by the procedure described previously for 4, with 9  and elemental selenium  yielding a white solid. Crystals suitable for X‐ray diffraction were grown by layering a solution of 12 in dichloromethane with hexane . M.p. 203–206 °C; IR (KBr disc): ν˜=2964s, 2921w, 1548w, 1494w, 1469w, 1454s, 1364w, 1261vs, 1202s, 1170w, 1094vs, 1016vs, 882w, 812vs, 754s, 614s, 578vs, 548vs, 445 cm−1 w; 1H{31P} NMR : δ=7.8 , 7.6 , 7.4 , 1.4 ppm ; 13C{1H} NMR : δ=135.9 , 130.6 , 130.4 , 128.3 , 126.3 , 126.0 , 46.3 , 25.6 ppm ; 31P{1H} NMR : δ=53.8 ppm =794 Hz); 77Se{1H} NMR : δ=−1520.5 ppm =794 Hz); MS (APCI+): m/z (%): 358.9588 (100) [M+H]+, 279.0427 (64) [C14H15S2P]+; elemental analysis calcd (%) for C14H15SePS2 (357.3): C 47.1, H 4.2; found: C 47.2, H 4.3.Compound 2O2  was added dropwise to a solution of 13  in dichloromethane (40 mL) and stirring was continued for 1 h. The reaction mixture was washed with water (100 mL), and the organic layer dried with magnesium sulfate and concentrated under reduced pressure. Crystals suitable for X‐ray diffraction were grown from dichloromethane . M.p. 199–201 °C; IR (KBr disc): ν˜=3422.9s, 2957.2s, 1592.5w, 1541.6s, 1490.2w, 1455.7s, 1362s, 1317.3w, 1196.6vs, 1175vs, 1137.9s, 1008.6w, 819.1vs, 804s, 758.3vs, 689.1w, 616.1s, 505.5vs, 468.3vs, 396.1w, 318.6w, 286.6w, 259.7 cm−1 vs; 1H{31P} NMR : δ=7.8 , 7.3 , 1.4 ppm ; 13C{1H} NMR : δ=136.3 , 134.0 , 130.8 , 128.6 , 126.3 , 44.0 , 25.0 ppm ; 31P{1H} NMR : δ=44.1 ppm =406.7 Hz); 77Se{1H} NMR : δ=392.9 ppm =406.7 Hz); MS (APCI+): m/z (%): 390.9268 (28) [M+H]+, 316.8537 (42) [C10H6Se2P]+, 285.8798 (84) [C10H6Se2]+, 253.9396 (100) [C10H6SePO]+, 236.9370 (82) [C10H6SeP]+, 206.9710 (31) [C10H6Se]+, 128.0620 (24) [C10H8]+; elemental analysis calcd (%) for C14H15OPSe2 (388.2): C 43.3, H 3.9; found: C 43.2, H 3.8.H15 was prepared by the procedure described previously for 7, with 13  and elemental sulfur  yielding a white‐green solid. Crystals suitable for X‐ray diffraction were grown by layering a dichloromethane solution of 15 with hexane . M.p. 199–202 °C; IR (KBr disc): ν˜=3417w, 2965w, 1638w, 1538w, 1491w, 1455w, 1356s, 1191s, 1013w, 847w, 808s, 750s, 669vs, 581s, 489s, 430 cm−1 w; 1H{31P} NMR : δ=7.8 , 7.8 , 7.4 , 1.3 ppm ; 13C{1H} NMR : δ=135.8 , 131.3 , 130.7 , 128.9 , 126.8 , 125.8 , 47.6 , 24.2 ppm ; 31P{1H} NMR : δ=48.6 ppm =398 Hz); 77Se{1H} NMR : δ=413.2 ppm =398 Hz); MS (EI+): m/z (%): 405.8 (44) [M].+, 285.8 (100) [C10H6Se2].+, 205.9 (28) [C10H6Se].+, 126.0 (40) [C10H6⋅]+; elemental analysis calcd (%) for C14H15SPSe2 (404.23): C 41.6, H 3.7; found: C 41.7, H 3.6.Compound 16 was prepared by the procedure described previously for 8, with 13  and elemental selenium  yielding a light purple solid. Crystals suitable for X‐ray diffraction were grown by layering a dichloromethane solution of 16 with methanol . M.p. 182–185 °C; IR (KBr disc): ν˜=3423.7s, 2967s, 2283.9w, 1537.6s, 1490.6w, 1453.4s, 1355.6s, 1327.7s, 1190.3s, 1164.6s, 1012.3s, 846.1w, 807.8vs, 749.1vs, 593.7s, 556.4w, 534.8vs, 520.7vs, 482.2s, 417.4s, 375.4 cm−1 s; 1H{31P} NMR : δ=7.9 , 7.8 , 7.4 , 1.3 ppm ; 13C{1H} NMR : δ=136.1 , 131.1 , 130.8 , 130.3 , 127.4 , 126.1  48.2  25.3 ppm ; 31P{1H} NMR : δ=27.3 ppm =407 Hz, 1J (31P–77Se)=752 Hz); 77Se{1H} NMR : δ=406.1 =407 Hz), −143.7 ppm =752 Hz); MS (NSI+): m/z (%): 919.7195 (10) [2 M+NH4−H] 452.8489 (100) [M+H]+; elemental analysis calcd (%) for C14H15PSe3 (453.8): C 37.3, H 3.3; found: C 37.4, H 3.3.Compound 1 a, 8 a, 9 and 13 were collected at −148(1) °C by using a Rigaku MM007 High‐Brilliance RA generator  and Saturn CCD system. At least a full hemisphere of data was collected by using ω scans. Data for 1 b, 2, 3 a, 4 a, 6, 7, 10, 12 a, 14, 15 and 16 a were collected at −100(1) °C, and those for 1 c, 3 b, 3 c, 4 b and 16 b at −180(1) °C by using a Rigaku FR‐X Ultrahigh‐Brilliance Microfocus RA generator  with XtaLAB P200 diffractometer. At least a full hemisphere of data was collected by using ω scans. Data for 5 were collected at −180(1) °C by using a Rigaku MM007 High Brilliance RA generator  and Mercury CCD system. At least a full hemisphere of data was collected by using both ω and φ scans. Data for 8 b were collected at −100(1) °C by using a Rigaku SCXmini CCD diffractometer . At least a full hemisphere of data was collected by using ω scans. Data for 11 and 12 b were collected at −148(1) °C by using the St Andrews Automated Robotic Diffractometer (STANDARD),Kα radiation, SHINE monochromator) and Saturn 724 CCD system, coupled with a Microglide goniometer head and an ACTOR‐SM robotic sample changer. Data for all compounds were collected and processed  with CrystalClear (Rigaku).X‐ray diffraction data for https://www.ccdc.cam.ac.uk/services/structures?id=doi:10.1002/chem.201800978 1816237 (1 a), 1816238 (1 b), 1816239 (1 c), 1816236 (2), 1816241 (3 a), 1816235 (3 b), 1816246 (3 c), 1816243 (4 a), 1816245 (4 b), 1057058 (5), 1816242 (6), 1816248 (7), 1816249 (8 a), 1816244 (8 b), 1816257 (9), 1816247 (10), 1816252 (11), 1816251 (12 a), 1816254 (12 b), 1057057 (13), 1816253 (14), 1816255 (15), 1816256 (16 a), and 1816258 (16 b) contain the supplementary crystallographic data for this paper. These data can be obtained free of charge from http://www.ccdc.cam.ac.uk/.31P, MAS NMR spectra were acquired at 298 K, 14.1 T and 7.5 kHz MAS with 1H decoupling. For variable‐temperature experiments the sample temperature was controlled with a Bruker BCU‐II chiller and Bruker BVT/BVTB‐3000 temperature controller and heater booster. The sample temperature was calibrated by using the temperature‐dependent shift of an external sample of RbCl.3PO4 (aq.) at 0 ppm, by using BPO4 at −29.6 ppm as a secondary reference. For 77Se, CP MAS experiments (with ramped contact pulse durations of 5–8 ms and TPPM 1H decoupling) were carried out at 298 K at 9.4 and 14.1 T. Chemical shifts were referenced to (CH3)2Se at 0 ppm, by using the isotropic resonance of solid H2SeO3 at 1288.1 ppm as a secondary reference. The position of the isotropic resonances within the spinning sideband manifolds were unambiguously determined by acquiring a second spectrum at a different MAS rate. In some cases, spectra were also acquired with additional 31P continuous wave decoupling. Experimental NMR parameters were determined by line‐shape analysis with Bruker Topspin software, SOLA.Solid‐state NMR measurements were performed with Bruker Avance III spectrometers operating at magnetic field strengths of 9.4 and 14.1 T. Experiments were carried out with conventional 4, 1.9 or 1.3 mm MAS probes at MAS rates between 5 and 55 kHz. For J coupling were carried out with the CASTEP DFT code (version 17.52),k‐point spacing−1. For all calculations, the initial atomic positions and unit‐cell parameters were taken from the single‐crystal X‐ray diffraction structures determined in this work. Prior to the calculation of NMR parameters, geometry optimisations were performed for each structure. All atomic positions and lattice parameters were allowed to vary. All J coupling constant were tested for convergence with supercell size by constructing supercells based on the optimised unit cells; for 1 a, 1 b, 1 c, 5 and 13 a 2×1×1 supercell was adopted, while for 9 a 1×2×1 supercell was used. Calculations performed on isolated molecules in alternative conformations were carried out by using CASTEP 7 . Models were produced by extracting a single molecule from the unit cell and placing it in a 20 Å periodic box before geometry optimisation of all atomic positions. For the alternative X‐P‐Ec‐Napc dihedral angle (see Results and Discussion) a 180° rotation about this dihedral angle was performed manually and the geometry reoptimised. Calculations were performed on two computing clusters at the University of St Andrews; single‐molecule calculations used a cluster of 300×12‐core Intel Westmere nodes connected via QDR Infiniband, and more resource‐intensive calculations were performed on a cluster of 54× 32‐core Intel Broadwell nodes with FDR Infiniband interconnect and 300 TB distributed file system (GPFS).Calculations of The authors declare no conflict of interest.As a service to our authors and readers, this journal provides supporting information supplied by the authors. Such materials are peer reviewed and may be re‐organized for online delivery, but are not copy‐edited or typeset. Technical support issues arising from supporting information (other than missing files) should be addressed to the authors.SupplementaryClick here for additional data file.


N-glycans, an important modification of glycoproteins. Recently, several studies have reported that removal of core α1-6-linked fucose from N-glycans labeled with the reactive N-hydroxysuccinimide carbamate fluorescent labels 6-aminoquinolyl-N-hydroxysuccinimidylcarbamate (AQC) and RapiFluor-MS is severely impeded. We report here the cloning, expression and biochemical characterization of an α-fucosidase from Omnitrophica bacterium (termed fucosidase O). We show that fucosidase O can efficiently remove α1-6- and α1-3-linked core fucose from N-glycans. Additionally, we demonstrate that fucosidase O is able to efficiently hydrolyze core α1-6-linked fucose from N-glycans labeled with any of the existing NHS-carbamate activated fluorescent dyes.Exoglycosidases are often used for detailed characterization of glycan structures. Bovine kidney α-fucosidase is commonly used to determine the presence of core α1-6 fucose on  N-glycosylation) is an abundant and complex form of post-translational modification of eukaryotic secretory proteins. Common techniques like liquid chromatography (LC) coupled to fluorescent detection (FLR) and/or mass spectrometry (MS), or capillary electrophoresis (CE) with laser-induced fluorescence (LIF) have been used to structurally characterize N-glycans. Data from these analyses can be compared to validated mass or mobility data in reference databases to permit assignments of N-glycan structures2. An added measure of glycan structure verification can be gained through the use of exoglycosidases in these analytical workflows4.Asparagine-linked glycosylation  are each coupled to the reducing end GlcNAc of a free N-glycan using Schiff base chemistry )6, InstantAB™, InstantPC™, and 6-aminoquinolyl-N-hydroxysuccinimidylcarbamate (AQC)8 utilize reactive NHS-carbamate chemistry to form a covalent urea linkage between the fluorophore and a glycosylamine moiety that transiently resides on the reducing end GlcNAc following N-glycan release by PNGase F  has historically been the preferred enzyme for confirmation of the presence of α1-6 fucose side-branched to the N-glycan core. Recent studies reported that removal of core α1-6-linked fucose from N-glycans labeled with the aminoquinoline dyes AQC or RFMS by BKF was severely impeded10. Molecular modeling of BKF with a core fucosylated RFMS labeled N-glycan suggested that rigidity of the aminoquinoline label contributed to steric clashes within the BKF active site9. In the present study, we have extended these biochemical observations by showing that BKF is also inefficient at removing core α1-6-linked fucose from N-glycans labeled with NHS-carbamate derivatives of aminobenzamide dyes . Thus, BKF is severely limited as a tool for confirming N-glycan core fucose in workflows using any of the reactive NHS-carbamate fluorescent labels that are currently available.While newer labels have advantages for N-glycans labeled with reactive NHS-carbamate fluorescent dyes. We report here the cloning, expression and biochemical characterization of an α-fucosidase from Omnitrophica bacterium (termed fucosidase O). We show that fucosidase O has a strong preference for hydrolysis of α1-6-linked core fucose over α1-2-, α1-4-linked fucose in several glycans that we tested. The enzyme also can remove α1-3-linked core fucose that can occur on N-glycans of plants, worms and other non-mammalian eukaryotes. Finally, we demonstrate that fucosidase O is able to efficiently hydrolyze core α1-6-linked fucose on N-glycans labeled with NHS-carbamate activated fluorescent labels. Our study provides an alternative enzymatic solution for confirmation of N-glycan core α1-6 fucose.We sought to identify a novel α1-6-fucosidase with the ability to remove core fucose from Omnitrophica bacterium OLB16 (herein termed fucosidase O) was identified via computational interrogation of sequence repositories. Fucosidase O is a member of CAZy glycoside hydrolase family GH29. Its deduced protein sequence (KXK31601) shows 48% and 45% amino acid sequence identity to those of the bovine tissue (NP_001039500) and plasma (NP_001192747) α-L-fucosidases, respectively. Additionally, the enzyme has a 20 amino acid signal peptide as predicted by the SignalP 4.0 algorithm11. To generate recombinant protein for biochemical characterization, fucosidase O lacking its signal peptide was intracellularly expressed in E. coli. The recombinant protein was purified as described in Materials and Methods   Fig. . Terminase) Fig. . However II Fig. , and no ate Fig. .Figure 3The linkage preference of recombinant fucosidase O was compared to that of native BKF using fucosylated oligosaccharide substrates containing α1-2-linked fucose (2′-fucosyllactose) and α1-6-linked fucose (NA2F). After a 1 hour incubation, fucosidase O released α1-6 and α1-2-linked fucose with 95% and 27% efficiency, respectively; while BKF released 33% of α1-6-linked fucose and 100% of α1-2-linked fucose Fig. . TherefoN-glycans as substrates (see Supplementary Table Nicotiana benthamiana plants12 were labelled with anthranilic acid (2-AA), and were incubated with fucosidase O. The reaction products were analyzed by MALDI-TOF-MS. No hydrolysis of the α1-3 fucosyl Lewis X motif attached to the N-glycan outer arm was detected . Thus, 23-fold higher concentration of fucosidase O (35 U/mL) and extended incubation time (16 hours) at 37 °C were required to achieve complete digestion of RFMS-labeled glycans  by BKF and fucosidase O was evaluated. Complete defucosylation of RFMS-labeled NGA2F was observed after treatment with fucosidase O (35 U/mL) for 16 hours at 37 °C . This experiment was performed by incubating NA2F-InstantAB or NA2F-InstantPC with fucosidase O (35 U/mL) or BKF (2 U/mL) for 16 hours at 37 °C. BKF was able to remove 90% of core fucose from NA2F-InstantAB and only 49% from NA2F-InstantPC, whereas ≥99% of core fucose was released from both substrates by fucosidase O . A standard of human IgG N-glycans labeled with RapiFluor-MS was obtained from Waters . Native bovine kidney fucosidase  fucosidase) was from Prozyme. 2-chloro-4-nitrophenyl α-L-fucopyranoside (CNP-Fuc) was obtained from CarboSynth US .All chemical reagents and solvents were purchased from Sigma-Aldrich. Labeled Omnitrophica bacterium OLB16 (GenBank: KXK31601.1) was identified in the Omnitrophica genome database14. A codon optimized DNA sequence encoding fucosidase O lacking its signal peptide (23-449 amino acids) with terminal vector-specific overlapping sequences was synthesized by Integrated DNA Technologies . Bacterial expression vector pJS11915 was amplified using the primers 5′ ATGTTAACCTCCTAAGCTTAATTC 3′and 5′ GAATTCAGCTTGGCTGTTTTG 3′ and purified by gel extraction. The DNA fragment was cloned into pJS119 using the NEBuilder™ HiFi DNA Assembly Cloning Kit . The resulting pJS119k-FucO plasmid and its sequence is available upon request.A DNA fragment  encoding fucosidase O from E. coli NEB Express cells . An overnight culture of transformed cells was diluted 1:100 in 2 L of LB medium supplemented with 50 μg/mL kanamycin and grown to 0.6 OD600 units at 37 °C. The expression of recombinant fucosidase O was induced by addition of isopropyl-β-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM with shaking for 4 h at 30 °C. The cells were harvested by centrifugation and resuspended in 50 mL of 20 mM Tris-HCl, pH 7.5. The cells were lysed by sonication with six 15 s bursts. Cell debris was removed by centrifugation at 19,000 × g for 60 min at 4 °C. The cleared cell lysate was applied to a 5 mL (bed volume) DEAE column  equilibrated with 20 mM Tris-HCl, pH 7.5 at a flow rate of 2 mL/min. Bound proteins were eluted with a 0-300 mM NaCl gradient in 20 mM Tris-HCl, pH 7.5 and collected in 5 mL fractions. The fractions containing fucosidase O were pooled, ammonium sulfate was added to 1.5 M final concentration at 4 °C, and the protein sample was directly applied to a 5 mL (bed volume) Phenyl Sepharose FF (Low Sub) column  at a flow rate of 3 mL/min. Bound protein was eluted using a reverse gradient of 1.5-0 M ammonium sulfate in 20 mM Tris-HCl, pH 7.5 (5 mL fraction size). Pooled fractions containing pure protein were dialyzed against 20 mM Tris-HCl, pH 7.5 containing 50 mM NaCl, 1 mM EDTA and concentrated using Vivaspin 20 concentrators . The yield of purified enzyme corresponded to 0.8 mg per liter of starting cell culture.For protein expression, the assembled plasmid pJS119k-FucO was introduced into The colorimetric substrate 2-chloro-4-nitrophenyl α-L-fucopyranoside (CNP-Fuc) was used to assay enzymatic activity during purification. Typically, 1 µL α-fucosidase fraction was added to 100 µL of 2 mM CNP-Fuc in 20 mM sodium acetate buffer, pH 5.5, and incubated 1 hour at 37 °C. Light absorbance was read at 405 nm.N-acetylglucosaminidase, β-N-acetylgalactosaminidase, β-galactosidase, α-galactosidase, α-neuraminidase, α-mannosidase, α-glucosidase, β-xylosidase and β-mannosidase  was incubated at 37 °C for 16 h with 3 μg of fucosidase O in 10 μL of reaction buffer . The reaction mix was spotted onto a silica-60 thin layer chromatography (TLC) plate  and separated using a mobile phase of isopropanol-ethanol-water . Reaction products were visualized by UV light at 302 nm. Purified fucosidase O was tested for the following activities: β-N-glycan G0F labeled with 2-aminoacridone (GlcNAcβ1-2Manα1-6(GlcNAcβ1-2Manα1-3)Manβ1-4GlcNAcβ1-4GlcNAc(Fucα1-6)-AMAC), in 1 hour at 37 °C in a total reaction volume of 10 µL. To assay fucosidase O, two-fold dilutions of enzyme were incubated with 1 nmol AMAC-labeled G0F substrate in 50 mM sodium acetate (pH 5.5) containing 5 mM CaCl2 in a 10 µL reaction. The reaction mix was incubated at 37 °C for 1 hour. Separation of reaction products were visualized via thin layer chromatography as described above.One unit of fucosidase O was defined as the amount of enzyme required to cleave >95% of fucose from 1 nmol of the human IgG N-glycan substrates with 2-aminobenzamide (2AB), 10 μL of a fluorescent labeling mix  was added to each tube containing a dried N-glycan sample. The reaction was incubated at 65 °C with agitation at 700 rpm for 120 min. For labeling with 7-amino-4-methylcoumarin (AMC), 10 μL of a labeling mix  was added per 3 nmol of dried N-glycans. The reaction was incubated at 80 °C for 45 min. Excess label was removed by passage over HILIC SPE MacroSpin columns . Briefly, labeling samples (10 μL) were diluted to 300 μL with 90% acetonitrile/10% 50 mM ammonium formate, pH 4.4 (90% ACN/NH4F), loaded on columns equilibrated with the same buffer, and washed with 5 × 350 μL 90% ACN/NH4F. Samples were eluted in 100 μL 50 mM NH4F, pH 4.4.To label N-morpholino)ethanesulfonic acid buffer (pH 6.0-6.5) or sodium phosphate buffer (pH 7.5–8.0). Each reaction had a final volume of 10 μL. After incubation for 30 min at 37 °C, 5 μL of each sample were analyzed by UPLC-HILIC-FLR.To determine the optimum pH of fucosidase O, 2 pmol of 2AB-labeled trimannosyl N-glycan substituted with α1-6-linked core fucose (M3N2F) were incubated with 10 mU fucosidase O in parallel reactions containing 50 mM citrate buffer (pH 3.0), 50 mM sodium acetate buffer (pH 4.0-5.5), 50 mM 2-, after which 5 μL of each sample were analyzed by UPLC-HILIC-FLR.To determine the optimum temperature fucosidase O, 2 pmol 2AB-labeled NA2F were mixed with 10 mU fucosidase O in 50 mM sodium acetate buffer, pH 5.5 containing 5 mM CaClTo examine the effect of cations on fucosidase O activity, 2 pmol 2AB-labeled M3N2F were mixed with 10 mU fucosidase O in 50 mM sodium acetate buffer, pH 5.5 containing appropriate metal salts at the final concentration of 2 mM in a final reaction volume of 10 μL. After incubation for 60 min at 37 °C, 5 μL of each sample were analyzed by UPLC-HILIC-FLR.N-glycan or oligosaccharide substrates containing α1-6-, α1-2-, α1-3- or α1-4-fucose. A list of substrates used in this study is shown in Supplementary Table 2, and 105 mU of fucosidase O or 6 mU GLYKO α fucosidase [BKF] in a total reaction volume of 70 µL . Reactions were incubated at 37 °C and a 10 µL aliquot was harvested at regular time points. To each sample, 100 µL of 20% acetonitrile was added to stop the reaction. Each reaction mixture was transferred to the Nanosep 10 K Omega  centrifugal device and centrifuged for 5 min at 12,000 × g. The samples were dried using a SpeedVac concentrator and were each dissolved in 10 μL of deionized water. For UPLC-HILIC-FLR analysis, 5 μL of each sample was mixed with 11.7 μL acetonitrile . An 10 μL aliquot of this mix was injected for UPLC-HILIC-FLR separation (next section).Substrate specificity and the glycosidic bond preference of purified fucosidase O and BKF were compared using N-glycan containing α1-6-linked fucose substrate used for fucosidase O, and a synthetic substrate 4-nitrophenyl α-L-fucopyranoside for BKF). Since the enzymes exhibit different activity on each of these two substrates, we used equimolar concentrations of each protein (0.15 μg) rather than equal number of units for comparison studies  on a Waters H-Class ACQUITY instrument  equipped with a quaternary solvent manager and a fluorescence detector. Solvent A was 50 mM ammonium formate buffer pH 4.4 and solvent B was acetonitrile. The gradient was 0-1.47 min, 30% solvent A; 1.47-24.81 min, 30–47% solvent A; 25.5-26.25 min, 70% solvent A; 26.55-32 min, 30% solvent A. The flow rate was 0.56 mL/min. The injection volume was 10 μL and the sample was prepared in 70% (v/v) acetonitrile. Samples were kept at 5 °C prior to injection and the separation temperature was 40 °C. The fluorescence detection wavelengths were: λex = 330 nm and λem = 420 nm for 2-AB; λex = 278 nm and λem = 344 nm for InstantAB. The data collection rate was 20 Hz.N-glycans were separated by UPLC using a Waters Acquity BEH glycan amide column  on a Waters H-Class ACQUITY instrument. Solvent A was 50 mM ammonium formate buffer pH 4.4 and solvent B was acetonitrile. The gradient used was 0-35 min, 25–46% solvent A; 36.5-39.5 min, 100% solvent A; 43.1-55 min, 25% solvent A. The flow rate was 0.4 mL/min. The injection volume was 5 μL and the sample was prepared in 75% (v/v) acetonitrile. Samples were kept at 5 °C prior to injection and the separation temperature was 60 °C. The fluorescence detection wavelengths were λex = 265 nm and λem = 425 nm with a data collection rate of 20 Hz. Waters Empower 3 chromatography workstation software was used for data processing including traditional integration algorithm, no smoothing of the spectra and manual peak picking.RapiFluor-MS-labeled human IgG N-glycans were isolated from recombinant glycoproteins produced in the wild-type and genetically modified Nicotiana benthamiana plants12. Isolation, labeling with anthranilic acid (2-AA) and characterization of these N-glycans were performed as described previously12.Various N-glycans were incubated overnight at 37 °C with 2 U of fucosidase O in 50 mM sodium acetate, pH 5.5 containing 5 mM CaCl2. After incubation, N-glycan products were loaded onto C18 Zip-Tip®  conditioned with 50% ACN + 0.1% TFA and then 0.1% TFA. The C18 resin was washed with 0.1% TFA and digestion products were eluted in 50% ACN + 0.1% TFA mixed with 2,5-dihydroxybenzoic acid (DHB) matrix  so that samples were directly spotted on a 384 well steel polished plate for MALDI analysis and dried at room temperatureThe 2AA-labeled www.glycoworkbench.org). The 2AA label was taken into account as a fixed reducing-end modification and possible glycan composition was set up based on the characterization work previously conducted by Wilbers et al.12 . A deviation of 150 ppm maximum was allowed for assignment of compositions.2AA-labeled undigested and digested glycan samples were analyzed by Matrix Assisted Laser Desorption/Ionisation – Time of flight mass spectrometry (MALDI-TOF-MS) using an UltrafleXtrem® mass spectrometer  equipped with a 1 kHz Smartbeam II laser technology and controlled by the software FlexControl 3.4 Build 119 . All spectra were recorded in the negative-ion reflectron mode using DHB as matrix. Bruker® peptide calibration mix (ref # 8206195) was used for external calibration. Spectra were obtained over a mass window of m/z 700 –3500 with ion suppression below m/z 700 for a minimum of 20,000 shots (2000 Hz) obtained by manual selection of “sweet spots”. The software FlexAnalysis 3.4 Build 76 was used for data processing including smoothing of the spectra , baseline subtraction  and manual peak picking. For our purpose of highlighting fucosidase O specificity, only the major peaks in each spectrum were selected and peaks with a signal to noise ratio inferior to 3 were excluded as well as peak to which no glycan composition could be assigned. Deprotonated masses of the selected peaks were assigned using GlycoPeakfinder® tool of the software GlycoWorkbench 2 (Supplementary Material


The phenyl ring is inclined to the mean plane of the imidazopyridine unit by 18.2 (1)°. In the crystal, molecules are linked by N—H⋯H hydrogen bonds, forming chains along the c-axis direction. The chains are linked by C—H⋯π interactions, forming slabs parallel to the ac plane. The Hirshfeld surface analysis and fingerprint plots reveal that the crystal structure is dominated by H⋯H (54%) and C⋯H/H⋯C (35.6%) contacts. The crystal studied was refined as an inversion twinThe bicyclic imidazo[1,2- For instance, it is found in several commercialized drugs such as the sedative Zolpidem, the anxiolytics Alpidem, Saridipem and Necopidem, the heart-failure drug Olprinone and the antiulcer drug Zolimidine -2-(phenylethynyl)imidazopyridin-3-amine.Compounds containing the imidazopyridine portion is slightly twisted with a dihedral angle of 3.6 (1)° between the mean planes of the five- and six-membered rings. The dihedral angle between the mean plane of the imidazopyridine moiety  and the phenyl ring (C10–C15) is 18.2 (1)°.In the title compound Fig. 1, the fusc-axis direction by N3—H3A⋯N1i hydrogen bonds  to 1.341 (blue). The red spots on the surface indicate the intermolecular contacts involved in the hydrogen bonds. Fig. 4a illustrates the intermolecular N—H⋯N hydrogen bonding of the title compound with normd mapped on Hirshfeld surface, and the C—H⋯π(ring) contacts are visualized in Fig. 4b. The fingerprint plots are given in Fig. 5b) and C⋯H/H⋯C at 35.6% , followed by N⋯H/H⋯N interactions at 10.2% .The Hirshfield surface analysis imidazopyridine, gave zero hits. A search for N-(tert-butyl)imidazopyridin-3-amines gave six hits (see supporting information). As in the title compound, the ° in the title compound.A search of the Cambridge Structural Database  was added to a mixture of 2-aminopyridine , phenylpropargyl aldehyde  and perchloric acid  in a 50 ml flask at room temperature. The reaction mixture was stirred for 4 h at rt. The crude product was purified by flash chromatography on silica gel (2:8 ethyl acetate/petroleum ether). Colourless crystals were isolated when the solvent was allowed to evaporate .Uiso(H) = 1.5Ueq(C-methyl) and 1.2Ueq(C) for other C-bound H atoms. The NH H atom was located in a difference-Fourier map. Its parameters were adjusted to give N—H = 0.89 Å and it was then refined as riding with Uiso(H) = 1.2Ueq(N). The crystal studied was refined as an inversion twin, with a final BASF value of 0.3 (6).Crystal data, data collection and structure refinement details are summarized in Table 210.1107/S2056989019012751/su5507sup1.cifCrystal structure: contains datablock(s) global, I. DOI: Click here for additional data file.10.1107/S2056989019012751/su5507Isup3.cdxSupporting information file. DOI: 10.1107/S2056989019012751/su5507Isup4.hklStructure factors: contains datablock(s) I. DOI: 10.1107/S2056989019012751/su5507sup5.pdfCSD search. DOI: 1953481CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


The dihedral angles between the pyrrole ring and the two phenyl rings are 6.7 (3) and 54.7 (3)°. In the crystal, a supramolecular ring structure is formed between two azopyrrole and two methanol solvent molecules through four O—H⋯O hydrogen bonds.The azo N=N bond adopts a  17H15N3O2·CH3OH, the azo N=N bond adopts a trans configuration and the pyrrole N and azo group are in an anti orientation. The dihedral angles between the pyrrole ring and the two phenyl rings are 6.7 (3) and 54.7 (3)°. In the crystal, a supramolecular ring structure is formed between two azopyrrole and two methanol solvent molecules through four O—H⋯O hydrogen bonds.In the title azopyrrole compound, C The diazonium salt solution was added dropwise to a solution of N-methylpyrrole  in acetonitrile (25 mL) and three drops of acetic acid. The combined solution was maintained at 273 K for 2 h with stirring. After that, EtOAc (25 mL) and water (25 mL) were added. The organic layer was separated and washed with water (20 mL) and dried with anhydrous MgSO4. The solution was evaporated and the residue was purified by column chromatography on silica (ethyl acetate/petroleum ether = 1:2), which gave the title compound as an orange powder .A 273 K solution of 2-aminophenol 0.272 g (2.5 mmol) and aqueous HCl (2 mL) in water (2 mL) was treated with another 273 K solution of NaNO1H NMR : δ 3.73 , 6.32 , 6.86 , 6.90–6.95 , 6.99 , 7.23–7.31 , 7.65 , 9.95 , 10.43  . Crystals suitable X-ray diffraction analysis were obtained by the slow evaporation of a CHCl3/CH3OH solution of the title compound.Uiso(H) = 1.2Ueq(C) or 1.5Ueq(Cmethyl).Crystal data, data collection and structure refinement details are summarized in Table 210.1107/S2056989018007764/ff2153sup1.cifCrystal structure: contains datablock(s) I. DOI: 10.1107/S2056989018007764/ff2153Isup2.hklStructure factors: contains datablock(s) I. DOI: Click here for additional data file.10.1107/S2056989018007764/ff2153Isup3.cmlSupporting information file. DOI: 1845021CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


III atom in the solvated title complex adopts a distorted octahedral geometry coordinated by two C,N-chelating 2,6-dimethyl-3-(pyridin-2-yl)pyridin-4-yl ligands and one O,O′-chelating 2,2,6,6-tetramethylheptane-3,5-dionate ligand. The title compound shows bright blue–green emission in solution at room temperature.The Ir 12H11N2)2(C11H19O2)]·CH3CO2C2H5, the IrIII ion adopts a distorted octahedral coordination environment resulting from its coordination by two C,N-chelating 2,6-dimethyl-3-(pyridin-2-yl)pyridin-4-yl ligands and one O,O′-chelating 2,2,6,6-tetramethylheptane-3,5-dionate ligand. The C,N-chelating ligands are perpendicular to each other [dihedral angle between the least-squares planes = 87.86 (5)°] and are arranged in a cis-C,C′ and trans-N,N′ fashion. In the crystal, pairwise C—H⋯π interactions between inversion-related IrIII complexes are present, forming a dimeric structure. The title complex shows bright bluish–green emission with good quantum efficiency in solution at room temperature.In the solvated title compound, [Ir(C Compared with alkoxy substituents, the methyl group has been regarded as essentially the same substituent because of its electron-donating nature. However, an IrIII complex based on methyl-substituted bipyridine as a main ligand emits strong green phosphorescence emission at room temperature  complexes have recently attracted much attention because of their applications in organic light-emitting diodes (OLEDs) pyridin-4-yl ligands and a 2,2,6,6-tetramethylheptane-3,5-dionate ligand. The IrIII atom has a distorted octahedral coordination sphere defined by two C,N-chelating ligands and one O,O′-chelating ligand. The C,N-chelating ligands, which are almost perpendicular to each other [dihedral angle between the least-squares planes = 87.86 (5)°], are arranged in cis-C,C′ and trans-N,N′ fashions. These arrangements are similar to those in [Ir(ppy)2(acac)] ] pyridin-4-yl-κC4]iridium(III) propane-1,3-dionato-κ2O,O′]iridium(III) bisiridium(III) , leading to the formation of a dimeric structure. The IrIII complex molecules and the ethyl acetate solvent molecules are also connected by a C—H⋯π interactions Ir(acetylacetonate) (617 K). This may be due to the methyl substituents of the main bipyridine ligand.As shown in Fig. 4κ2N,C4)iridium(III)  transition based on a previous report  of the title complex was estimated to be 2.52 eV using the emission spectrum at 77 K. The quantum efficiency (ΦPL) of the title complex was estimated using FIrpic, bis(picolinato)iridium(III), as a standard (ΦPL = 0.6) to be 0.4. The high thermal stability and good quantum efficiency of the title complex makes it a potentially useful triplet emitter for applications in OLEDs.The title compound displays bright bluish–green emission in solution at room temperature, as shown in Fig. 52 atmosphere using standard Schlenk techniques. All solvents were freshly distilled over appropriate drying reagents prior to use. All starting materials were commercially purchased and used without further purification. The 1H NMR spectrum was recorded on a Bruker Avance 400 MHz spectrometer. The thermogravimetric spectrum was recorded on a Perkin–Elmer TGA-7 under a nitrogen environment at a heating rate of 10 K min−1 over a range of 298–973 K. The IrIII dimer, [(Me2pypy)2Ir(μ-Cl)]2, and the title compound were synthesized according to previous reports 2Ir(μ-Cl)]2, , sodium carbonate , and 2,2,6,6-tetramethylheptane-3,5-dione  were dissolved in THF/MeOH . The reaction mixture was stirred overnight at ambient temperature. All volatile components were removed under reduced pressure. The mixture was poured into EtOAc (50 ml), and then washed with water (3 × 50 ml) to remove excess sodium carbonate. Silica gel column purification with EtOAc and hexane gave a yellow powder in 60% yield. Yellow plates were recrystallized from ethyl acetate/hexane solution at low temperature. 1H NMR : 8.41, 8.08 , 7.80 , 7.12 , 6.02 , 5.47, 2.66 , 2.22 , 0.68 .All experiments were performed under a dry Nd(C—H) = 0.95 Å, Uiso(H) = 1.2Ueq(C) for Csp2 H atoms, and 0.98 Å, Uiso(H) = 1.5Ueq(C) for methyl protons.Crystal data, data collection and structure refinement details are summarized in Table 310.1107/S2056989018011076/hb7762sup1.cifCrystal structure: contains datablock(s) I, New_Global_Publ_Block. DOI: 10.1107/S2056989018011076/hb7762Isup2.hklStructure factors: contains datablock(s) I. DOI: 1859964CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information:


The title compound is composed of a bis(hydrazoniminato)copper(II) complex and the basic hydrazone imine shows a molecular framework with layers each constructed from one of the two components arranged in alternating order. Layer formation is supported by C—H⋯O interactions. 24H24N8O6)]·C11H11N3O4, each of the crystal components forms undulating layers which stack alternately along the b-axis direction. Molecules of the CuII complex are connected via C—H⋯O hydrogen bonds involving the nitro and keto oxygen atoms, thus forming supramolecular networks. Molecules of the arylhydrazone component are linked by C—H⋯O interactions into zigzag strands showing no interstrand association.In the title 1:1 co-crystal, [Cu(C In the presence of a diamine and NiII, related bis(hydrazonoimine) complexes are formed displaying an unusual behavior of the effective magnetic moment at low temperature hydrazine-1-ylidene]-4-oxopentane-2-iminato}copper(II) and 3-[2-(3-nitrophenyl)hydrazine-1-ylidene]pentane-2,4-dione in a 1:1 stoichiometric composition (the title compound), whose crystal structure is reported on herein.Hydrazone imines derived from β-diketones and aryldiazonium salts using a Japp–Klingemann route  with one molecule of the CuII complex and one molecule of the arylhydrazone in the asymmetric unit. A perspective view is shown in Fig. 1et al., 2018The title co-crystal possesses orthorhombic symmetry hydrazono-4-oxo-5,5,6,6,7,7,8,8-octafluorooctane-2-iminato]nickel(II) hydrazine-1-ylidene]pentane-2,4-dione and bis(ethylenediamine)copper(II) chloride in Uiso(H) = 1.5Ueq(C) for methyl and Uiso(H) = 1.2Ueq(C) for other H atoms.Crystal data, data collection and structure refinement details are summarized in Table 210.1107/S2056989019005838/yk2120sup1.cifCrystal structure: contains datablock(s) I. DOI: 10.1107/S2056989019005838/yk2120Isup2.hklStructure factors: contains datablock(s) I. DOI: 1912679CCDC reference:  crystallographic information; 3D view; checkCIF reportAdditional supporting information: