Structural features of hydrolysable tannins determine their ability to

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Cite This: J. Agric. Food Chem. 2019, 67, 6798−6808

Structural Features of Hydrolyzable Tannins Determine Their Ability to Form Insoluble Complexes with Bovine Serum Albumin M. T. Engström,*,† J. Arvola,† S. Nenonen,‡ V. T. J. Virtanen,† M. M. Leppä,† P. Tähtinen,† and J. -P. Salminen† †

Natural Chemistry Research Group, Department of Chemistry, University of Turku, FI20014 Turku, Finland Turku PET Centre, Turku University Hospital, Fl20520, Turku, Finland

‡

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S Supporting Information *

ABSTRACT: The ability of 32 purified and characterized hydrolyzable tannins to form insoluble complexes with model protein bovine serum albumin was investigated with a turbidimetric 96-well plate reader method. The results showed a clear relationship between the hydrolyzable tannin structure and the intensity of haze that formed during the tannin−protein complexation. In addition to molecular weight, structural features such as number of galloyl groups, degree of oxidative coupling between the galloyls, positional isomerism, and cyclic vs acyclic glucose core were the major structural features that affected the ability of the monomeric hydrolyzable tannins to form insoluble complexes with bovine serum albumin. While oligomers were superior to monomers in their capability to precipitate the model protein, their activity depended less on the functional groups, but mostly on their size and overall flexibility. These results allowed us to construct an equation that predicted the protein precipitation capacity of the studied hydrolyzable tannins with high accuracy. KEYWORDS: gallic acid derivatives, ellagitannins, gallotannins, protein precipitation capacity, structure−activity patterns

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INTRODUCTION The interactions between proteins and hydrolyzable tannins (HTs) are important in numerous plant related domains, from the most basic functions in plant physiology and ecology to their utilization in agriculture, foods, and medicine.1−8 The interactions occurring between HTs and proteins can be divided into three main categories: covalent or noncovalent,1,5,9−11 soluble or insoluble, and specific or nonspecific.4,12 Both the type of binding and the availability of the HT and protein to interact with each other are affected by the physicochemical conditions such as solvent composition, other ions, and molecules present in the solution as well as ionic strength and temperature. Yet, the key factors that determine the type of interactions are the exact HT and protein structures as well as their concentrations and the pH of the reaction solution.1,4,9,13 The importance of the tannin structure in tannin−protein interactions was underlined already by Haslam,14 and since then, several studies have made an effort to report the structural features that affect the tendency of HTs to form complexes with proteins. Unfortunately, many of the investigations on how the HT structure affects their ability to bind with proteins have suffered from the difficulty of obtaining a library of pure, individual tannin structures to compare against their protein binding activity. Thus, many studies have either utilized semipurified fractions or extracts instead of pure compounds. Alternatively, studies utilizing pure compounds have used only a limited number of compounds thus disabling them to make broader conclusions of, for example, how the protein binding activity of HTs decreases, increases, or stays the same along the various steps or branches of the biosynthetic pathway of HTs. On the other hand, such © 2019 American Chemical Society

broader conclusions may be difficult to obtain also by metaanalysis of previous studies, if they have used different tannins, proteins, and methods to evaluate tannin−protein interactions. Nevertheless, the most obvious structural characteristics of HTs affecting their complex formation with proteins seem to be their size and structural flexibility, degree of galloylation, and oxidative coupling between the galloyls.15−25 In our previous studies, we have shown that the oxidative activities26,27 and the anthelmintic activities of HTs28 can be predicted with equations that take many of the structural details of HTs into account. In addition, the oxidative activity of HTs seemed to be inversely correlated with their protein precipitation capacity (PPC), and the oxidative activity changed logically along the proposed biosynthetic pathway of HTs.29,30 Thus, we hypothesized that the PPC of HTs could also be (1) accurately estimated by their structural characteristics, and that they would (2) depend on the position of HTs in their biosynthetic pathway. Should this be true, it would allow more accurate estimation of the potency of specific HTs, or plant species accumulating specific types of HTs,31 to form complexes with proteins. In the present study, we purified a library of 32 HTs (Figures 1 and 2) representing 11 biosynthetic branches of the HT pathway (Supporting Information (SI) Figure S1) and investigated their ability to form insoluble complexes with BSA using a simple turbidimetric 96-well plate reader method. Our goal was to verify the previously reported patterns between Received: Revised: Accepted: Published: 6798

April 8, 2019 May 27, 2019 May 28, 2019 May 28, 2019 DOI: 10.1021/acs.jafc.9b02188 J. Agric. Food Chem. 2019, 67, 6798−6808

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Journal of Agricultural and Food Chemistry

Figure 1. Structures of the monomeric hydrolyzable tannins studied. HHDP: hexahydroxydiphenoyl, DHHDP: dehydrohexahydroxydiphenoyl, NHTP; nonahydroxytriphenoyl; GG: galloylglucose. (1) monoGG, (2) diGG, (3) triGG, (4) tetraGG, (5) pentaGG, (6) hexaGG, (7) heptaGG, (8) octaGG, (9) tellimagrandin I, (10) tellimagrandin II, (11) pedunculagin, (12) casuarictin, (13) 1,2-digalloyl-4,6-HHDP-glucose, (14) punicalagin, (15) geraniin, (16) chebulagic acid, (17) vescalagin, (18) castalagin, (19) vescavaloninic acid, (20) castavaloninic acid, (21) stachyurin, (22) casuarinin.

HTs and their ability to form insoluble complexes with BSA and to complement those with more detailed structure− activity patterns that would arise from our multiple compoundto-compound comparisons. We hoped to be able to (1) create a simple equation for the estimation of the PPC of known HT structures, and to (2) make conclusions on how the PPC of HTs varies between different branches of their proposed biosynthetic pathway. Together with our previous equations, this fundamental knowledge could be used to estimate, for example, how different plant species are ranked in their

bioactivities (oxidative activity, anthelmintic activity, protein precipitation) on the basis of their hydrolyzable tannin composition and content.

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MATERIALS AND METHODS

Chemicals. Technical grade acetone for extraction was purchased from VWR (Haasrode, Belgium). Analytical grade acetone and methanol used in the Sephadex LH-20 fractionation and HPLC grade methanol and acetonitrile used in the preparative and semipreparative purification were from VWR International (Fontenay-Sous-Bois, France). Formic acid and LC-MS Chromasolv acetonitrile for the 6799

DOI: 10.1021/acs.jafc.9b02188 J. Agric. Food Chem. 2019, 67, 6798−6808

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Figure 2. Structures of the oligomeric hydrolyzable tannins studied and the different types of linkages between the monomers. m-DOG: valoneoyl group, m-GOG: dehydrodigalloylgroup, m-GOD: sanguisorboyl group, 2x m-DOG: macrocyclic structure. (23) oenothein B, (24) oenothein A, (25) sanguiin H-6, (26) lambertianin C, (27) gemin A, (28) agrimoniin, (29) salicarinin A, (30) salicarinin B, (31) salicarinin C, (32) salicarinin D. were the same as used in Moilanen et al.31 and white birch (Betula pubescens) leaves the same as in Salminen et al.32 Isolation and Purification of HTs. The extraction and isolation of the HTs followed mainly the methods reported in Baert et al.,33 shortly described in following: Extraction. The plant materials were collected directly into 1 l glass bottles and macerated in acetone at 4 °C for several weeks. After repetitive maceration with additional batches of acetone/water (80/ 20, v/v), the different extraction batches were combined, acetone was evaporated and the remaining aqueous solution was filtered and lyophilized. This kind of a lengthy extraction procedure may cause, for example, oxidation or artifact formation for some of the HTs. This aspect was not relevant in this study, since we focused only on the production of such HTs that we had earlier identified from the plant species used. First Fractionation. The lyophilized extract was dissolved in water, mixed to a slurry of Sephadex LH-20 (in water) material and eluted

UHPLC-ESI-QqQ-MS were obtained from Sigma-Aldrich (Seelze, Germany). Water was purified with a Millipore Synergy water purification system from Merck KGaA (Darmstadt, Germany). Sephadex LH-20 material was obtained from GE Healthcare (Uppsala, Sweden). BSA (purified by heat shock fractionation, pH 7, purity ≥96%; lyophilized powder, 66 kDa) was purchased from Sigma-Aldrich (St. Louis, MO). Plant Material. The plant materials used for the isolation of the studied HTs were collected during the summers 2011−2017 from Southwestern Finland, including willowherb (Epilobium angustifolium) flowers, silverweed (Potentilla anserina) leaves, herb Bennet (Geum urbanum) leaves, English oak (Quercus robur) acorns, purple loosestrife (Lythrum salicaria) leaves and flowers, meadowsweet (Filipendula ulmaria) flowers, raspberry (Rubus idaeus) leaves, and wood cranesbill (Geranium sylvaticum) leaves. Black myrobalan (Terminalia chebula) powder was ordered from Banyan Botanicals (Albuquerque, NM). Sea buckthorn (Hippophae rhamnoides) leaves 6800


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Journal of Agricultural and Food Chemistry with water, methanol/water (1:1 v/v), methanol, acetone/water (4:1 v/v), and acetone in a Büchner funnel (⌀ = 240 mm) in vacuo with a filter paper. Column Chromatography. Sephadex LH-20 (in water) was loaded into a glass column (40 × 4.8 cm i.d., Kimble-Chase Kontes Chromaflex) and equilibrated with 1500 mL of ultrapure water at a flow rate of 5 mL min−1. A maximum of 10 g of the samples were dissolved in 15 mL of ultrapure water, filtered (0.2 μm, PTFE) and applied on top of the gel. The eluent profile depended on the HT to be isolated; the solvents used were ultrapure water, aqueous methanol, and aqueous acetone. Obtained fractions were analyzed by UPLC-DAD-MS, concentrated to the water phase and lyophilized. Selected Sephadex fractions were further purified by preparative and semipreparative liquid chromatography (LC). Preparative and Semipreparative Liquid Chromatography. The LC-DAD system consisted of a Waters Delta 600 liquid chromatograph, a Waters 600 Controller, a Waters 2998 Photodiode Array Detector and of a Waters Fraction Collector III. The column (327 × 33 mm) was manually filled with LiChroprep RP-18 (40−63 μm) material (Merck KGaA, Darmstadt, Germany). A binary solvent system with methanol (A) and 1% aqueous formic acid (B) at a constant flow rate of 8 mL min−1 was used and the elution protocol depended on the composition of the fractions; the typical gradient was as follows: 0−5 min, 100% B (isocratic); 5−180 min, 0−40% A in B (linear gradient); 180−220 min, 40−60% A in B (linear gradient); 220−240 min, 60−80% A in B (linear gradient). The final purification of HTs was performed by semipreparative LC with the same LC-DAD system described above. The column was a Gemini C18 column (150 × 21.2 mm, 10 μm, Phenomenex) and the eluents were acetonitrile (A) and 0.1% aqueous formic acid (B) and at a constant flow rate of 8 mL min−1. Different gradients were used for different HTs; for example, a typical gradient for acyclic ETs was as follows: 0−5 min, 2% A in B (isocratic); 5−51 min, 2−32% A in B (linear gradient); 51−55 min, 32−70% A in B (linear gradient). All steps in the preparative and semipreparative purifications were followed by UPLC-DAD-MS. Fractions with as pure HTs as possible were pooled, concentrated to the water-phase and lyophilized. Sample Analysis. After each purification step, the resulting fractions were analyzed by UPLC-MS. Sample analysis was carried out with an Acquity UPLC system (Waters Corporation, Milford, MA) coupled with a Xevo TQ triple quadrupole mass spectrometer (Waters Corporation, Milford, MA). The UPLC system consisted of a sample manager, a binary solvent manager, a column and a diode array detector. The column used was a 100 × 2.1 mm i.d., 1.7 μm, Acquity UPLC BEH Phenyl column (Waters Corporation, Wexford, Ireland). The flow rate of the eluent was 0.5 mL min−1. The elution profile used two solvents, acetonitrile (A) and 0.1% aqueous formic acid (B): 0−0.5 min, 0.1% A in B; 0.5−5.0 min, 0.1−30% A in B (linear gradient); 5.0−5.1 min, 30−90% A in B (linear gradient); 5.1−8.5 min, column wash and stabilization. UV and MS data were collected from 0 to 6 min. Negative ionization mode was used for MS analyses. ESI conditions were: capillary voltage 2.4 kV, desolvation temperature 650 °C, source temperature 150 °C, desolvation and cone gas (N2) 1000 and 100 L/h, respectively and collision gas was argon. All samples were filtered with a syringe filter (4 mm, 0.2 μm PTFE, Thermo Fisher Scientific Inc., Waltham, MA) prior to the UPLC-MS analyses. Structural Characterization of Hydrolyzable Tannins. The structures of the individual HTs are presented in Figures 1 and 2. The stereochemistry of castalagin and vescalagin were recently reinvestigated by computational methods and the nonahydroxytriphenoyl group (NHTP) found to exist in (S, R) configuration.34 Therefore, it is feasible that the NHTP group of vescavaloninic and castavaloninic acids is also in (S, R) configuration. The studied 32 HTs with information on the original plant material, the purity by UPLC at 280 nm, the ESI-MS identification and the original structural identification papers are presented in SI S1 Appendix. One of the compounds, the αβ-anomer of salicarinins A−C35 had not been previously reported. The structure of this novel compound was elucidated by NMR (SI S1 Appendix) and the compound was named salicarinin D.

The NMR experiments were performed for selected HTs with a Bruker Avance-III spectrometer equipped with a Smartprobe (Fällanden, Switzerland) operating at 500.08 MHz for 1H and 125.76 MHz for 13C. Spectra were recorded at 25 °C using acetone-d6 as the solvent. The experiments included standard 1H and 13C NMR spectral measurements, and in addition, gradient-enhanced COSY, 2D-ROESY (with 200 ms mixing time), multiplicity-edited HSQC, HMBC and band-selective CT-HMBC (optimized for 8 Hz longrange JCH coupling constants), and selective 1D-TOCSY (with 200 ms mixing time) experiments. The chemical shifts are reported with respect to the chemical shifts of the solvent signals: δH = 2.05 ppm and δC(Me) = 29.84 ppm. The ECD spectrum for salicarinin D in water at 298 K was measured with a Chirascan circular dichroism spectrometer (Applied Photophysics, Leatherhead, UK) utilizing a 1 mm path-length cuvette. The spectrum was scanned over the range of 200−450 nm, background subtracted and smoothed. Protein Precipitation Capacity. The haze formation of the 32 HTs with BSA was measured with a 96-well plate reader (Multiscan Ascent, Thermo Electron Corporation) at room temperature. HT solutions of 1.0 mM, 0.8 mM, 0.6 mM, 0.4 mM, 0.2 mM, and 0.1 mM were prepared in EtOH/H2O (1/9, v-v) to ensure HT solubility, and 200 μM BSA solution was prepared in pH 5 buffer (0.05 M acetate containing 60 μM ascorbic acid). Thus, the tannin to protein molar ratios (t:p) tested were 5:1, 4:1, 3:1, 2:1, 1:1, and 1:2, respectively. With the method, the haze formation of two compounds were measured on each 96-well plate; first compound on the first four rows and the second compound on the latter four rows. The first of the four rows was always for the control sample, in which 100 μL of HT solution and 100 μL of buffer (without BSA) was added. The three rows below were for the actual haze formation measurements. In these, 100 μL of the HT solution (starting from 1 mM and ending up with 0.1 mM) was pipetted in each well, followed by 100 μL of BSA solution with a 12-channel pipet. In total, six replicates for each concentration were measured. After pipetting the solutions, the plate reader method was started with a 1 min mixing period, after which the absorbance was measured in 30 s intervals for 30 min at 340 and 415 nm (including a 10 s mixing period before each measurement). Both the molar ratios tested and the pH (close to BSA isoelectric point) of the reaction solution were set to favor insoluble complex formation. From the kinetic data, maximal absorbance during the whole measurement time was used in data analysis (see SI Figure S1). Data Analysis. The average insoluble complex formation values (ICFAVER) were calculated from the whole concentration range using the maximal absorbance values from the kinetic data (see SI Figure S1). The initial concentration for the insoluble complex formation (ICFINIT), that is, the x-intercept values were calculated by fitting dose response curves (concentration vs maximal absorbance) in Origin (Origin 2015) using the calculate x-intercept function (see SI Figure S1). For HTs 5−8 and 23−32 the dose was ln transformed to achieve a proper model fit. The regression coefficients for the structural features of the HT monomers predicting insoluble complex formation were obtained by the Ordinary Least Squares estimator as implemented in the lm-function in R-software (version 3.5.2).

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RESULTS AND DISCUSSION

Tannin Selection and Purification. Thirty-two HTs (Figure 1 and 2) were successfully purified (purities between 81% and 100%) from 11 plant species to achieve a good repertoire of compounds representing 11 different branches of the proposed HT pathway (SI Figure S2). The monomers were represented by 22 HTs, from which five were simple galloyl glucoses (1−5) and three gallotannins (6−8). Simple hexahydroxydiphenoyl (HHDP) esters were presented by six ellagitannins (ETs, 9−14), each carrying the characteristic feature of ETs, the HHDP moiety. In addition, 14 was the only HT possessing a gallagyl group. The HHDP group can be oxidized to form a dehydrohexahydroxydiphenoyl group

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Figure 3. Average insoluble complex formation of the whole concentration range (ICFAVER, black columns) and initiation concentration for insoluble complex formation, that is, the x-intercept (ICFINIT, red columns) for the studied hydrolyzable tannins. * ICFINIT value could not be determined. For compound identities, see Figures 1 and 2

formation occurred even at the lowest tested concentration (0.1 mM, t:p = 1:2), absorbance values increasing until reaching or starting to reach the saturation point after which absorbance did not increase when the HT concentration further increased. The saturation point was a consequence of the tested tannin to protein ratios, which were adjusted to favor insoluble complex formation and to achieve insoluble complex formation for all the studied compounds, if possible. Thus, BSA was the limiting reagent in complex formation for the most efficient protein precipitators. Regarding the monomers, 1, 2, and 11 showed no significant haze formation with BSA at the tested concentration range while for the most active monomers, 7 and 8, haze formation occurred even at the lowest concentration tested. To better compare PPCs of the studied HTs, two values were calculated: (1) the average insoluble complex formation for the whole concentration range (ICFAVER) and (2) the initial concentration for the insoluble complex formation (ICFINIT), that is, the x-intercept (Figure 3). Although maximum binding responses and EC50 values (concentration to reach half-maximum binding) are often used in turbidimetry studies, these values did not fit our aims to compare the PPCs using the same HT to protein molar ratios for each HT, since all HTs did not reach the maximum binding responses. Therefore, ICFAVER and ICFINIT values were selected to get an overall picture of the PPCs of the studied HTs at the studied concentration range, taking into account different types of haze formation versus concentration plots and to have one specific value to use in the structure−activity comparisons. The ICFINIT concentration values were of particular interest as they determine both the HT concentration and the HT to BSA ratio required for the initiation of the insoluble complex formation. In general, these two measures correlated well (R2

(DHHDP) as in 15 which can further be rearranged into modified DHHDP as in 16. Six of the monomers were Cglucosidic ETs (17−22). The 10 oligomeric ETs were constructed of either simple HHDP esters or C-glucosidic monomers and were divided into five different oligomer types.36 Compounds 23 and 24 were macrocyclic oligomers, 23 bearing two m-DOG linkages and 24 one m-DOG and one m,m′-D(OG)2 linkage. Sanguisorboyl groups (m-GOD) were found in 25 and 26. Dehydrodigalloyl groups (m-GOG) were found in 27 and 28. Compounds 29− 32 were C-glycosidic oligomers with an intermolecular valoneoyl group (m-DOG). Protein Precipitation Capacity. In the case of BSA and HTs, the kinetic measurements with the plate reader yielded very little information on the reaction kinetics as the haze formation occurred so rapidly that maximal absorbance values were obtained almost immediately after adding the BSA to the HT solution (see SI Figure S1). Therefore, instead of using the raw kinetic measurement data, the maximal absorbance values at each HT to BSA ratio were investigated when looking for structure−activity patterns. The maximal absorbance values of the 32 HTs with BSA varied considerably between the compounds and concentrations as could be seen from the individual concentration versus maximal absorbance plots (SI Figures S3−S6). When comparing the activities of the monomers and the oligomers, it was evident that especially at the higher concentrations the monomers had more compound-to-compound variation in their concentration versus maximal absorbance plots than the oligomers. For the oligomers, most of the compound-to-compound variance was caused by 23 and 26 that had lower and higher absorbance values, respectively, than the other oligomers with more uniform concentration versus absorbance plots. For most of the dimers (25, 27−32) and the two trimers (24 and 26) haze 6802


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Figure 4. Average insoluble complex formation of the whole concentration range (ICFAVER, A) and initiation concentration for insoluble complex formation, that is, the x-intercept (ICFINIT, B) plotted as a function of molecular weight (MW) for the studied hydrolyzable tannins. Gray cubes indicate monogalloylglucose to octagalloylglucose, black cubes other monomers and black dots the oligomeric hydrolyzable tannins. *ICFINIT value could not be determined.

∼ 0.90) but for some of the compounds differences occurred between the two values. The Effect of Molecular Weight on the Insoluble Complex Formation. The ICFINIT and ICFAVER values revealed several structure−activity patterns between the studied HTs. As reported already by Haslam 197414, to some extent, higher molecular weights and thus, the number of phenolic groups, indicated higher ICFAVER and lower ICFINIT values (Figure 4A,B). For instance, all oligomers but 23 had both higher ICFAVER and lower ICFINIT values than any monomer. Also, for the simple galloyl glucoses and gallotannins (1−8) higher MW resulted in higher ICFAVER and lower ICFINIT values but not in a linear manner (Figure 4A,B). Two series of triplet ETs containing monomer and its dimer and trimer could be compared regarding MW vs protein precipitation: 9, 23 and 24, and 12, 25 and 26. For both triplets good correlations for ICFAVER versus MW and ICFINIT versus MW were obtained (R2 ∼ 0.94/0.91 and R2 ∼ 0.91/0.80 respectively). However, it was evident that for the trimers, the PPC increased less than expected by the corresponding increase from monomers to dimers; this is consistent with the previous results suggesting that correlation between tannin size and protein binding capacity may have an upper limit as the steric hindrance of large tannins may prevent access to binding sites.37,38 Also, as could be seen in the ICFAVER versus MW and ICFINIT versus MW plots (Figure 4A,B) at the MW area 700−1200 Da a wide range of PPCs were measured for HTs with MWs relatively close to each other. Thus, to better explain the reasons for the activity differences of the studied HTs, a more detailed structure versus PPC comparison was required. In these comparisons, both the ICFAVER and the ICFINIT were considered separately. The Effect of the Degree of Galloylation on the Insoluble Complex Formation. In agreement with the results of Kilkowski and Gross,17 comparisons within the galloylglucoses 1−8 indicated that one or two galloyl groups as in 1 and 2 did not yet result in haze formation with BSA even if

the HT concentration was increased up to 4 mM (t:p = 20:1) to obtain ICFINIT values. The first galloyl glucose causing haze formation with BSA at the tested concentration range was 3 with three galloyl groups. The biggest increase in ICFAVER was between 3, 4, and 5 (0.18 → 0.94 → 1.24) and biggest decrease in ICFINIT between 3 and 4 (0.55 → 0.19). Additional galloyl groups forming digalloyl groups, as in gallotannins 6, 7, and 8, did further increase the ICFAVER values (1.25 → 1.40 → 1.53) but not as much as the galloyls attached directly to the glucose core. This might be related to the partial steric hindrance caused by the addition of galloyl groups to the nearly spherical core pentagalloylglucose39 which hinders the positive effect of increased number of galloyl units. Interestingly, the ICFINIT values were less affected after addition of the fourth galloyl group (4 → 5 → 6 → 7 → 8; 0.19 → 0.17 → 0.15 → 0.12 → 0.11), suggesting that the number of tannin molecules required for haze formation to occur did not so much depend on the degree of galloylation. In general, the increase in the number of galloyl groups might increase the ability to form intermolecular complexes leading to more effective haze formation, which could be seen in the increasing ICFAVER values. However, previous studies have suggested different precipitation modes for 5 and the much more polar 7; the latter one would interact with protein by hydrophilic interaction and the former one dominantly by hydrophobic interaction.39 This could explain the differences in the change of ICFAVER between simple galloylglucoses (1− 5) and gallotannins (6−8) and in anyway confirms that the PPC of HTs increase at the same time as the galloylglucose and gallotannin biosynthesis progresses.11,30 The importance of galloyl groups attached directly to the glucose core was emphasized also by the ET pairs 9 versus 10 and 11 versus 12, in which additional galloyl group attached to the hydroxyl group of the anomeric carbon significantly increased the ICFAVER values (0.20 → 1.07 and 0.04 → 0.87) and decreased the ICFINIT values (0.54 → 0.18 and 0.94 → 0.30). Similarly, comparison of the structural isomers 9 and 13 6803


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the gallagyl group markedly increased the ICFAVER (0.03 → 1.30) and lowered the ICFINIT (0.94 → 0.30) when compared against the 4,6-HHDP bearing analogue 11. Interestingly, 14 (1084 Da) formed insoluble complexes with BSA much efficiently than 4, 5, or 6 (788−1092 Da), suggesting a relatively high importance for the large-sized, but nonflexible gallagyl group in comparison to the smaller and flexible galloyls. It is also possible that the relatively less water-soluble nature of the ellagic acid moiety in the gallagyl group of 14 increased the hydrophobic interaction of 14 with BSA, thus forcing the complex out of water solution. This finding is in line with the general trend of increasing water-solubility of HTs−often co-occurring with the progressing biosynthesis of especially ETs−affecting their PPC negatively.16,29,30 The Effect of the Cyclic versus Acyclic Glucose Core. The HTs used in the study could be divided in two categories whether the HT structure is based on cyclic (1−16, 23−28) or acyclic glucose (17−22, 29−32). Regarding monomers, previous studies have reported the beneficial effect of cyclic glucose core over acyclic glucose core in regard to noncovalent complexation.21−23 Our results confirmed the findings for monomeric hydrolyzable tannins; the comparison of 21 and 22 with 12 suggested that C-glycosidic ETs resulted in lower insoluble complex formation than the corresponding glucopyranose-based ETs (ICFAVER 0.45 and 0.60 vs 0.87; ICFINIT 0.41 and 0.36 vs 0.30). In all three HTs, the phenolic moieties consisted of two HHDP groups and one galloyl group. For the NHTP-containing ETs such direct comparisons could obviously not be made since NHTPs cannot be found in others but C-glycosidic ETs. However, comparison of acyclic 17 and 18 with cyclic 2, 11, 9, and 13 showed that the combination of HHDP + NHTP in the acyclic glucose core resulted in more efficient complex formation than G+G, HHDP + HHDP, or HHDP + G + G in the glucopyranose ring, if the critical O1 position was ungalloylated; this again highlighted the importance of MW within the monomers. Regarding oligomers, the oligomerization seemed to conceal the importance of the structural characteristics of the participating monomers and the activities of the studied dimers, cyclic or acyclic, were of the same magnitude. This is likely to be caused by the increased size and overall flexibility of the HT molecules.16,21,23,24 The only outlier was 23 due to its rigid macrocyclic structure that was earlier shown to have even weaker protein affinity than its monomeric counterparts.24 The orientation of the hydroxyl group at the C1 position of the C-glycosidic ETs, such as vescalagin and castalagin, has been shown to affect both their physicochemical properties as well as bioactivities.27,28,41−47 In the present study, the orientation of the hydroxyl group at the anomeric carbon affected both the ICFAVER and the ICFINIT values as could be seen when comparing monomer pairs 17 versus 18, 19 versus 20, 21 versus 22 and the dimers 29−32. In the monomers, αorientation resulted in more efficient complex formation in all three pairs, the effect being greatest between 17 and 18 (ICFAVER 0.35 → 0.86, ICFINIT 0.51 → 0.27) and lower between 19 and 20 (ICFAVER 0.50 → 0.71, ICFINIT 0.50 → 0.37) and between 21 and 22 (ICFAVER 0.45 → 0.60, ICFINIT 0.41 → 0.36). In the dimers, the differences were subtle but seemed to favor β-orientation as the order with increasing ICFAVER and decreasing ICFINIT was ββ, βα, αα, αβ (29, 30, 31 and 32, respectively).

further confirmed the importance of galloyl group at the O1position for obtaining a more efficient protein precipitation; the ICFAVER of 13 was more than twice as high as for 9 (0.53 vs 0.20) and the ICFINIT of 13 was more than one-third lower than for 9 (0.35 vs 0.54). The Effect of the Oxidative Coupling on the Insoluble Complex Formation. The characteristic functional unit of the HTs in the competing biosynthetic branch with gallotannins, that is, the HHDP unit of ellagitannins, is formed by oxidative coupling of two galloyl groups. Previous studies have systematically reported the negative effect of oxidative coupling for the protein affinities of HTs.13,21,23,40 Indeed, the negative effect of this modification to the ICFAVER and ICFINIT values was revealed by the comparison of following pairs; 4 versus 13, 5 versus 10, 10 versus 12 and 9 versus 11. In addition, from the oligomers, 27 versus 28 could be compared in a similar manner. In each monomer pair, the ICFAVER decreased (0.94 → 0.53, 1.24 → 1.07, 1.07 → 0.87, and 0.20 → 0.04) and ICFINIT increased (0.19 → 0.35, 0.17 → 0.18, 0.18 → 0.30, and 0.54 → 0.94) due to the formation of the HHDP group from the two galloyls. However, in the oligomeric pair 27 and 28, the latter with two galloyls coupled to form an HHDP group in comparison to the former was slightly more active regarding ICFAVER (1.73 → 1.79), whereas in the ICFINIT, no significant difference was observed (0.086 → 0.085). This further confirmed that the oligomerization partially fades the importance of different structural units that certainly affect the PPCs of monomeric HTs with BSA. Although the oxidative coupling of two galloyls to form an HHDP decreased the PPC of HTs, HHDP groups cannot be considered as signs of inactivity. Structures comprising two HHDP groups, as in 11, did not result in significant activity but neither did 2 with two galloyl groups. However, when comparing the mono-, di-, tri-, and tetragalloylglucoses (1, 2, 3, and 4) to HTs with additional HHDP groups (12, 9, 13, and 10), HTs with additional HHDP groups were more active from both ICFAVER and ICFINIT point of view. Also, the combination of two HHDP groups and one galloyl group (12) resulted in higher ICFAVER and lower ICFINIT values than two galloyls and one HHDP (9 and 13). Similarly, three galloyl groups and one HHDP group in the structure yielded better activities than four galloyl groups (10 vs 4). So, again the size of the HTs (molecular masses around 332, 484, 636, 788, and 940 Da) is more decisive factor for the protein precipitation than any single difference in the esterification pattern (e.g., two galloyls vs HHDP). However, two differences instead of one may already turn the favor to the compound in the smaller group, for example, trigalloylglucose (636 Da, i.e., smaller with three galloyls) being more active than pedunculagin (784 Da, i.e., larger with no galloyls). In addition to the comparisons above with several compound pairs, our selection of HTs enabled to differentiate the PPC between the DHHDP group and the modified DHHDP group (15 versus 16), and HHDP and the gallagyl group (11 versus 14). As these structural modifications were compared only by one HT pair each, the results should be treated with a certain caution in comparison to the other structural modifications compared above. The DHHDPcontaining 15 was more active than modified DHHDP containing 16 in both ICFAVER and ICFINIT (0.75 → 0.40, 0.28 → 0.52), again suggesting that the biosynthetically more advanced monomeric ETs might have lowered PPC. The addition of ellagic acid in between two galloyls in 14 to form 6804


Article

Journal of Agricultural and Food Chemistry The orientation of the hydroxyl group at the C1 position of the C-glycosidic ETs did have an impact also on the effect of other structural modifications. The results showed that the addition of gallic acid to 17 with β-hydroxyl at C1 to form 19 increased the ICFAVER (0.35 → 0.50) and slightly decreased the ICFINIT (0.51 → 0.50). However, in the second pair with α-hydroxyl at anomeric carbon (18 and 20) the additional gallic acid decreased the ICFAVER (0.86 → 0.71) and increased the ICFINIT (0.27 → 0.37). The effect of oxidative coupling could be compared also in the C-glycosidic HT pairs 21 versus 17 and 22 versus 18, in which galloyl and HHDP group of the former are oxidatively coupled to form the NHTP group of the latter. In the first pair with β-hydroxyl at C1, the formation of NHTP group decreased the PPC as evidenced by lowered ICFAVER (0.45 → 0.35) and increased ICFINIT (0.41 → 0.51). However, again in the second pair with α-hydroxyl at C1, the effect was the opposite as the formation of the NHTP increased the PPC: ICFAVER (0.60 → 0.86), ICFINIT (0.36 → 0.27). These examples suggest a more specific interaction for some of the studied ETs and BSA that cannot be interpreted by the more general patterns observed across all the HTs. Equation for the Estimation of the Insoluble Complex Formation of Hydrolyzable Tannins with Bovine Serum Albumin. The comparisons above enabled to conclude what kind of an effect the structural differences of the studied HTs have on the formation of insoluble complexes with BSA. Based on these conclusions, it was possible to create an equation to estimate the PPC for the studied HTs. As ICFINIT values were not obtained for all HTs, ICFAVER values were used for this purpose. The coefficients were empirically adjusted in Microsoft Excel to produce activity estimates that would best correlate with the measured values. First, all the major structural features were taken into account and for monomers the final equation was

Figure 5. Correlation between the measured and the calculated average insoluble complex formation of the whole concentration range (ICFAVER) for the studied hydrolyzable tannins. Black color indicates monomeric structures and red color represents oligomeric structures.

To confirm the conclusions and the correctness of the adjusted coefficients, we utilized the regression function in Rsoftware to get coefficients for each structural characteristic of the monomers. This approach confirmed all the assumptions made by compound to compound comparisons and the coefficients matched very well with the ones obtained empirically (R ∼ 0.96). Thus, it seems that the ability of monomeric HTs to form insoluble complexes with BSA can be accurately estimated based on the HT structure. Distribution of the Protein Precipitation Capacity along the Biosynthetic Pathway of HTs. Regarding the distribution of the PPC along the biosynthetic pathway of HTs, the results indicated that the progression of the galloylglucose and gallotannin biosynthesis yields HTs with higher PPCs. On the contrary, each oxidative step along the suggested biosynthetic pathway of ETs reduced the PPC as did the opening of the cyclic glucose core to yield C-glycosidic ETs. Also, on the proposed biosynthetic pathway of HTs, all steps leading to oligomeric HTs increased the PPC. Interestingly, regarding monomers an opposite trend has been observed for the oxidative activities of galloylglucoses and ETs; the progression of galloyl glucose biosynthesis decreased the oxidative activity and the progression of ET biosynthesis increased the oxidative activity.11,13,26,27,48 Altogether, as suggested previously, it seems that the PPC and oxidative activities of monomeric HTs are negatively correlated; HTs efficient in forming insoluble complexes with proteins have lower oxidative activities and vice versa. However, regarding the C-glycosidic oligomers, even HTs with high oxidative activity (29−32) had high PPCs. This brings an interesting insight for why some plant species are synthesizing these biosynthetically costly HTs: the gained double defense via both good oxidative activity and good PPC might be worth the costs. The same can be said of the C-glycosidic monomers 19 and 20, having the valoneic acid moiety, since this specific structural feature significantly increase both the oxidative and protein-binding activities of the C-glycosidic monomers (this study and Moilanen and Salminen27). However, one must bear in mind that BSA is not the best possible model for the defense

ICFAVER = 0.0015A + 0.15B + 0.4C + 0.55D − 0.15E − 0.6F (1)

where the letters present the following structural features: (A) how much smaller is the molecule to the “threshold” molecule pentagalloylglucose, that is, (X Da−940 Da; not calculated for HTs with MW larger than 940 Da); (B) the total number of biosynthetically original galloyl groups (i.e., HHDP, DHHDP = 2, NHTP = 3 etc.), or gallic acid attached to the 4,6-HHDP; (C) galloyl group attached to the O1, or OH-group with αorientation in C-glucosidic ETs (i.e., value 1 or 0); (D) the number of gallagyl groups; (E) the number of HHDP, DHHDP, or NHTP groups, or acyclic glucose cores; (F) the number of modified DHHDP groups. The equation did result in good correlation between the measured and calculated ICFAVER values for the monomers (R2 ∼ 0.95), but not for the oligomers (R2 ∼ 0.50), since we above-noted that oligomerization as such hinders the importance of the minor structural features. For this reason, another, more simplified, equation was created for oligomers: ICFAVER = 1.18A − 0.1B − 0.5C

(2)

where: (A) the total number of galloyl groups (calculated as for the monomers); (B) the number of macrocyclic linkages; (C) whether the ET is trimer or larger (i.e., value 1 or 0). When utilizing these two equations, a correlation with R2 ∼ 0.98 was obtained between measured and calculated ICFAVER values for the tested HTs (Figure 5). 6805



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related protein interaction that involves, for example, salivary proteins. Altogether, to understand the basis for the bioactivity of a given plant species or organ, it is not good enough to know which branches of the HT pathway are utilized for the production of its main HTs, but it is crucial to know the smaller structural differences within the single branches as well. Importantly, it must be noted that the results from the present study are valid for the formation of insoluble HT− protein complexes with globular proteins such as BSA. For other types of proteins, the PPC orders of the HTs might be slightly different. For example, in our study on the anthelmintic activities of HTs,28 most of the results were in agreement with present results but, for example, the progression of biosynthesis of pentagalloylglucose to gallotannins decreased the activity, whereas in the present study, the opposite was observed. Also, when comparing the present results to the results of Okuda et al.15 with HTs and hemoglobin, the main results are again convergent, but when comparing the effect of structural details of ETs, differences in results were observed. For example, the effect of the orientation of the hydroxyl group attached to the anomeric carbon in 21 and 22 was opposite for the complexation with hemoglobin and BSA, and the oligomerization did not increase the complexation as much for hemoglobin as it did for BSA. To conclude, the results obtained in the present study indicated a clear relationship between structural features of HTs and their ability to form insoluble complexes with the model protein BSA. We now know how the structure of the HTs affects their PPC and what molar ratios are required for initiation of haze formation between different types of HTs and BSA. However, this study did not yet yield information on the stability of the formed insoluble complexes nor the effect of pH in the complex formation. These aspects would be the obvious next steps to investigate to better understand the role of tannins in, for example, protecting dietary proteins from ruminal degradation and further liberation of proteins in the intestine of ruminants as pH and the nature of tannin−protein interactions change. Altogether, our current results form a good platform for choosing a more limited array of HTs into more detailed mechanistic studies of tannin−protein complexation.

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Article

AUTHOR INFORMATION

Corresponding Author

*Phone: +358 29 450 3168; e-mail: mtengs@utu.fi. ORCID

M. T. Engström: 0000-0003-4123-6039 M. M. Leppä: 0000-0002-7376-8462 J. -P. Salminen: 0000-0002-2912-7094 Funding

This work was supported by the Academy of Finland (Grant no. 298177 to J.-P.S. and grant no 310549 to Maarit Karonen). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We acknowledge the help of Eerik Piirtola, Susanna Räikkönen, Maria Hokkanen, Terhi Sundman, Sanjib Saha, Anu Tuominen, Jussi Suvanto, and Elina Puljujärvi during the purification process of some of the ellagitannins. Maarit Karonen and the LipidET project kindly provided one of the purified ellagitannins. Juuso Laitila calculated the regression coefficients with R. Naomi Engström is acknowledged for fruitful discussions.

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ABBREVIATIONS USED BSA, bovine serum albumin; DHHDP, dehydrohexahydroxydiphenoyl; ESI, electrospray ionization; ET, ellagitannin; G, galloyl group; HHDP, hexahydroxydiphenoyl; HT, hydrolyzable tannin; ICFAVER, average insoluble complex formation; ICFINIT, initiation concentration for insoluble complex formation; MW, molecular weight; NHTP, nonahydroxytriphenoyl; PPC, protein precipitation capacity; TQ, triple quadrupole; t:p, tannin to protein molar ratio; UPLC, ultrahigh performance chromatography.

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REFERENCES

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.9b02188. The studied 32 hydrolyzable tannins with information on the original plant material, the purity by UPLC at 280 nm and the ESI-MS identification. ECD and NMR interpretation and structural characterization of the new hydrolyzable tannin, salicarinin D (Appendix S1). A simple flowchart to illustrate how the initial haze formation data acquired was utilized to obtain the average insoluble complex formation values (ICFAVER) and the initial concentration for the insoluble complex formation (ICFINIT), that is, the x-intercept values (Figure S1). A simplified scheme of the current view of hydrolyzable tannin biosynthetic pathway (Figure S2). Haze formation vs concentration plots for the studied hydrolyzable tannins (Figures S3−S6). Structure and numbering of atoms for salicarinin D (Figure S7) (PDF) 6806


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Structural features of hydrolysable tannins determine their ability to

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