0 METMYOGLOBIN ASSAY (TANNIN)

METMYOGLOBIN ASSAY

The metmyoglobin assay is a rapid method which provides the degree of antioxidant protection possessed by an individual species. Original descriptions of the method can be found in:

1. Halliwell, B. (1987) FASEB J. 1, 358-364

2. Southorn, P.A. (1988) Mayo Clin. Proc. 63, 390-408

3. Miller, N.J., Rice-Evans, C., Davies, M.J., Gopinathan, V., and Milner, A. (1993) Clin. Sci. 84, 407-412

Our application to tannins is described in:

4. Hagerman, A.E.; Riedl, K.M.; Jones, G.A.; Sovik, K.N.; Ritchard, N.T.; Hartzfeld, P.W.; Riechel, T.L. High molecular weight plant polyphenolics (tannins) as biological antioxidants. J. Agric. Food Chem. (1998) in press.

The reagents for the metmyoglobin method can be purchased as a kit from Randox. It is much less expensive to prepare them yourself as follows:

Reagents

PBS Buffer (0.005 M Phosphate, 0.145 M NaCl)

A: Dissolve 0.68 g KH2PO4 and 8.5 g NaCl in 1 L of distilled water

B: Dissolve 1.14 g K2HPO4 x H2O and 8.5 g NaCl in 1 L of distilled water

Mix the two solutions together to give a pH of 7.4 (approximately 770 mL of A and 2 L of B). (Final concentration in assay is 5 mM).

Myoglobin (initial concentration between 1 mg myoglobin/mL PBS to 5 mg/mL). The initial concentration doesn't matter since the solution will be diluted as it runs through the column. The myoglobin is purchased from Sigma (# M-0630).

K₃Fe(CN)₆ (0.24 mg K₃Fe(CN)6 in 1 mL water)

2,2'-Azino-bis(3-ethylbenzthiazoline-6-sulfonic acid (0.8 mg ABTS in 1 mL PBS buffer). The ABTS is purchased form Sigma (# 1888). (Final concentration in the assay is 610 μM).

Trolox Standard (0.1564 g Trolox in 250 mL PBS buffer). The Trolox is purchased from Aldrich (# 23,881-3). If the solid does not dissolve, sonicate gently. (Final concentration in the assay is 2.5 mM).

Tannin Sample (1 mg tannin/mL of water). Make sure the actual concentration used is recorded in the laboratory notebook for the calculations. Prepare 1:10 dilutions.

Hydrogen Peroxide Dilute the 30 % w/v stock solution as follows:

#1 1 part (H₂O₂) plus 9 parts water

#2 1 part (#1) plus 9 parts water

#3 1 part (#2) plus 9 parts water

#4 1.7 parts (#3) plus 8.3 parts PBS buffer

(Final concentration in the assay is 250 μM).

Sephadex G-10-120 column (Height of 20 cm and Diameter of 1 cm). Equilibrate the column with the PBS buffer before using. This column can be prepared and reused indefinately, stored at room temperature.

Metmtoglobin. Mix equal volumes of myoglobin and K₃Fe(CN)₆ solutions. Run sample through column and collect the second fraction where the brown color starts to come off the column. The first fraction is just buffer and the third fraction is yellow containing the K₃Fe(CN)₆. Read the Abs @ 490 nm of the second fraction. Adjust the solution with buffer to give an absorbance reading of 0.147 so that the final concentration of metmyoglobin in the assay will be 6.1 μM). The equations used to calculate the amounts of the various forms of the myoglobin are found in Reference #3.

Procedure

Set up an appropriate number of 0.65 mL microfuge tubes to run each sample in duplicate or triplicate.

Add the 20 μL of the sample (water, standard, or tannin), 250 μL of metmyoglobin and 250 μL of ABTS to the tubes. Vortex the tubes.

Using a microcuvette (1 mL), blank the spectrophotometer at 600 nm with water.

Read the absorbance of each sample and record as Abs1.

Add 100 μL of the hydrogen peroxide substrate to the tubes. After exactly 3 minutes, read the absorbance and record as Abs2. (Hint: Add the substrate to four tubes at 30 second intervals. This provides enough time to read each sample at exactly three minutes).

Subtract Abs1 from Abs2 for each sample. The typical change in absorbance for the control (water as sample) is 0.296.

Calculations

The amount of putative antioxidant required to supress absorbance of the ABTS radical cation by 50% is compared to the amount of Trolox required for 50% supression in order to compare potency of various antioxidants. A Trolox standard curve is run with each set of samples because there is substantial day-today variability in the assay.

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0 PROTEIN DIGESTIBILITY (TANNIN)

PROTEIN DIGESTIBILITY

Although high levels of dietary tannin can interfere with protein utilization [Salunkhe, D.K.; Chavan, J.K.; Kadam, S.S. 1990. Dietary tannins: Consequences and remedies; CRC Press: Boca Raton, 1990], there is little evidence that tannins consumed in moderate amounts are detrimental to health. There is some evidence that low levels of dietary tannins are beneficial to ruminants [Lees, G.L. 1992. In: Hemingway, R.W. & Laks, P.E. (Editors), Plant Polyphenols. Synthesis, Properties, Significance, Page 915, Plenum Press: New York] and perhaps to humans [e.g., Jankun, J.; Selman, S.H.; Swiercz, R.; Skrzypczak-Jankun, E. Nature 1997, 387, 561], and some mammals have developed mechanisms for accomodating even rather high levels of dietary tannins [McArthur, C.; Hagerman, A.; Robbins, C.T. 1991. In: Palo, R.T. & Robbins, C.T. (Editors), Plant Defenses against Mammalian Herbivory. Page 103, CRC Press: Boca Raton]. Reports of tannin toxicity are generally linked to ingestion of large amounts of tannin or introduction by routes other than oral ingestion . Chemical modification of the tannin, which may occur during food preparation or cooking, may increase or decrease the toxicity of the tannin to certain animals

A major family of proteins secreted by the salivary glands of some animals constitutes the best characterized of the "defense mechanisms" against the possible toxic effects of dietary tannins . The parotid and submandibular savliary glands of some mammals synthesize a group of proteins which are unusually high in proline, the so-called salivary proline-rich proteins (PRPs). The PRPs are characterized by four general regions: a signal peptide, a transition region, a repeat region, and a carboxyl-terminal region [Carlson, D.M.; Zhou, J.; Wright, P.S. Prog. Nucl. Acid Res. Mol. Biol. 1991, 41, 1]. These unusual proteins undergo various post-translational modifications including proteolysis, phosphorylation, and glycosylation. PRPs collectively constitute about 70% of the proteins in human saliva, and several functions for these proteins have been proposed, including calcium binding, inhibition of hydroxylapatite formation, and formation of the dental-acquired pellicle . Recent evidence suggests that a primary role for these proteins may be protection against dietary tannins.

PRPs are constitutive in human saliva, but are induced by treatment with the beta-agonist isoproterenol in parotid and submandibular glands of rats , mice, or hamsters . In rats, dietary tannins induce the same biochemical and morphological changes and polyploidy events in the parotid glands (but not the submandibular glands) as does isoproterenol treatment . When young rats are fed a high tannin diet (2-4% tannin) they lose weight during the first three days, but after induction of the PRPs on the third day of the diet the animals start to gain weight at about the same rate as those on the control diet [Mehanso, H.; Hagerman, A.; Clements, S.; Butler, L. Rogler, J.; Carlson, D.M. Proc. Natl. Acad. Sci. (USA), 1983, 80, 3948]. The logical conclusion is that the PRPs are induced by dietary tannins to "neutralize" the detrimental effects of the tannins. Further evidence for the ability of PRPs to neutralize tannins is provided by observations with hamsters. Dietary tannins do not induce PRP synthesis in hamsters, and tannins have pronounced detrimental effects on hamsters . If weanling hamsters are fed a diet containing 2% tannin the animals fail to gain weight, and increasing the tannin level to 4% causes most of the animals to die within three days.

Salivary tannin-binding proteins have been found in some wild herbivorous mammals which consume tannin-containing plants. For example, mule deer saliva contains a protein which has high affinity for tannin ; mule deer are generalist herbivores and can accommodate tannin in their diets [Hagerman, A.E.; Robbins, C.T.; Weerasuriya, Y.; Wilson, T.C.; McArthur, C. J. Range Manag. 1992, 45, 57]. Herbivores such as sheep, which do not produce salivary tannin-binding proteins, prefer to consume tannin-free forages [Austin, P.J.; Suchar, L.A.; Robbins, C.T.; Hagerman, A.E. J. Chem. Ecol. 1989, 15, 1335] and are unable to accommodate dietary tannin [Hagerman, A.E.; Robbins, C.T.; Weerasuriya, Y.; Wilson, T.C.; McArthur, C. J. Range Manag. 1992, 45, 57]. The affinity of salivary tannin-binding proteins for specific types of tannin may be related to the feeding preferences of herbivores. Moose and beaver are specialist herbivores, and have very selective tannin-binding proteins . The tannin-binding proteins of generalist herbivores such as mule deer and bear show little selectivity for binding specific tannins [Hagerman, A. E.; Robbins, C.T. Can. J. Zool. 1993, 71, 628]. Although the amino acid sequences have not been reported for the salivary tannin-binding proteins from any wild mammals, the protein found in mule deer saliva is proline-rich.

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0 TANNINS AS ANTIOXIDANTS

TANNINS AS ANTIOXIDANTS

Although dietary tannins are often perceived as detremental because of their potential to affect protein digestibility or metal ion availability, it is also possible that tannins are beneficial. It is likely, based on our knowledge of tannin chemistry, that tannins are potential biological antioxidants.

Antioxidants are widely believed to be an important line of defense against oxidative damage, which has been implicated in a range of diseases including cancer, cardiovascular disease, arthritis, and aging (Kehrer, J.P. Crit. Rev. Toxicol. 1993, 23, 21). Biological antioxidants are generally divided into three groups: enzymes, such as superoxide dismutase; inhibitors of radical formation, such as Fenton reaction inhibitors; and free radical quenching agents, such as alpha-tocopherol (Vitmain E). Phenolics are good candidates as antioxidants because of their favorable redox potentials and the relative stability of the aryloxy radical (Simic, M.G. and Jovanovic, S.V. 1994. In: Ho, C-T., Osawa, T., Huand, M-T., & Rosen, R.T. (Editors), Food Phytochemicals for Cancer Prevention II. Teas, Spices and Herbs. Page 20, American Chemical Society: Washington DC).

Many low molecular weight, naturally occurring phenolics scavenge radicals as effectively as the antioxidant vitamins E and A when tested in vitro (Rice-Evans, C.A.; Miller, N.J.; Paganga, G. Free Rad. Biol. Med. 1996, 20, 933). Our recent work suggests that free or protein-complexed condensed and hydrolyzable tannins are more effective than small phenolics

Some low molecular weight phenolics are pro-oxidants in Fenton-driven systems, apparently because the phenolic is able to redox cycle the metal ion required for radical formation (Aruoma, O.I.; Murcia, A.M.;

Butler, J.; Halliwell, B. J. Agric. Food Chem. 1993, 41, 1880). The tannins we have tested do not act as pro-oxidants in Fenton systems, and in fact react very rapidly to quench the hydroxyl radical.

We have used the deoxyribose method and the metmyoglobin (Randox) method to characterize the antioxidant capabilities of the tannins (Hagerman, A.E.; Riedl, K.M.; Jones, G.A.; Sovik, K.N.; Ritchard, N.T.; Hartzfeld, P.W.; Riechel, T.L. J. Agric. Food. Chem. 1998, 46, 1887-1892). We have also

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0 DEOXYRIBOSE ASSAY (TANNIN)

DEOXYRIBOSE ASSAY

The deoxyribose assay is used to determine the reactivity of tannins toward hydroxyl radicals. The method sounds simple and straightforward, but it is technically difficult to get good results, and interpretation of the data is ambiguous especially for phenolics (which can participate not only as hydroxyl radical scavengers, but also as "pro-oxidants" and as metal ion chelators). I do not recommend this method for general use. Instead, the metmyoglobin (Randox) method is simple and yields better results.

Results we have obtained for tannins are described in Hagerman, A.E.; Riedl, K.M.; Jones, G.A.; Sovik, K.N.; Ritchard, N.T.; Hartzfeld, P.W.; Riechel, T.L. J. Agric. Food. Chem. 1998, 46, 1887-1892.

Original descriptions can be found in:

1. Aruoma, O. (1994) Methods in Enzymology. 233, 57-66

2. Halliwell, B., Gutteridge, J.M.C., and Aruoma, O. (1987) Analytical Biochemistry. 165, 215-219.

Reagents

Deaerated water. Heat an appropriate amount of distilled water at a hard boil for 5 to 10 minutes and allow to come to room temperature. Alternatively, bubble nitrogen through the water for approximately 10 minutes.

33.6 mM Deoxyribose. (450.7 mg deoxyribose in 100 mL of distilled water). Allow the deoxyribose solid to come to room temperature before weighing. Refrigerate the solution. The 2-Deoxy-D-Ribose is purchased from Sigma (D-2751). (Final concentration in the assay is 2.8 mM).

0.1 M FeCl3/0.1 M HCl (16.221 g of solid ferric chloride in 50 mL of 2N HCl and bring final volume to 1000 mL with deaerated water). Store indefinitely.

300 μM FeCl3 (150 μL of 0.1 M solution to a final volume of 50 mL of deaerated water). Prepare

immediately before use. (Final concentration in the assay is 25 μM).

1.2 mM EDTA (35.1 mg disodium salt in 100 mL water). Store indefinitely. (Final concentration in the assay is 100 μM).

120 mM Phosphate Buffer (1.6330 g potassium phosphate monobasic in 100 mL water). Adjust pH to 7.4 with 25% KOH. Store indefinitely. (Final concentration in the assay is 10 mM).

1.2 mM Ascorbate (21.1 mg L-ascorbic acid in 100 mL deaerated water). Prepare immediately before use. (Final concentration in the assay is 100 μM).

50 mM sodium hydroxide (200 mg sodium hydroxide dissolved in 100 mL distilled water). Stable for one week.

33.6 mM Hydrogen Peroxide (335 μL stock 30 % w/v solution to 100 mL distilled water). Prepare immediately before use. (Final concentration in the assay is 2.8 mM).

1 % w/v TBA (0.25 g thiobarbituric acid in 25 mL of 50 mM sodium hydroxide). Stir for approximately 1 hour and prepare daily.

2.8 % w/v TCA (1.4 mL 100% w/v trichloroacetic acid solution diluted with 50 mL water). Prepare daily.

Tannin (approximately 10 mg tannin in 1 mL deaerated water). Make sure to record the actual concentration of tannin used in a laboratory book. If not all the tannin dissolves, vortex the solution and sonicate for about 1 minute. Remove the tube from the sonicator, vortex, and centrifuge. Transfer the solution to a new tube. Repeat the centrifugation as needed.

A 37 C water bath and a boiling water bath will be needed in this experiment.

Method

Start by preparing a spreadsheet of all the reagent and sample amounts. Include blanks for each set of samples and the various reagent conditions.

Obtain and label an appropriate number of 15 mL polypropylene screw-top clinical centrifuge tubes for each set of samples to be tested in triplicate.

Add 35 μL each of the buffer and the deoxyribose solutionm to the 15 mL tubes. Add the appropriate amount of water listed on the spreadsheet. Vortex the tubes.

Add 35 μL of H₂O₂ and vortex again.

Add 35 μL each of the EDTA and FeC_l3 solutions. Vortex the tubes.

Finally, add the tannin and 35 μL of the ascorbate. Vortex and cap the tubes.

The final volume of the mixture in each tube should be 420 μL. Centrifuge @ 3200 rpm (~ 1900 g-force) for 1 minute to ensure that the entire sample is at the bottom of the tube. Vortex each tube lightly.

Place the tubes in the 37 C water bath for 1 hour.

Allow the tubes to cool for 5 minutes. Vortex and shake the tubes. Centrifuge @ 3200 rpm for 1 minute.

Add 350 μL each of the TBA and the TCA solutions, in that order. The volume in each tube is 1120 μL..

Loosely cap and vortex the tubes. Place them in the boiling water bath for exactly 20 minutes.

Carefully remove the tubes with a pair of tongs to avoid splashing the boiling water and cool for 20 minutes. The pink color will form.

Vortex and shake the tubes. Centrifuge @ 3200 rpm for 1 minute.

Add 1120 μL 1-butanol and shake gently to mix the two layers. Centrifuge @ 3200 rpm for 6 minutes to separate the layers.

Using a solvent-resistant (glass or quartz) microcuvette, blank the spectrophotometer at 532 nm with 1-butanol. Take at least 900 μL of the upper layer of each sample and record the absorbance.

Subtract the blank absorbance from each appropriate sample set reading.

The typical absorbance for the control (no tannin) is 1.02 (although this seems to vary substantially).

Modifications

Several modifications were made to the Aruoma method in Reference #1.

The volume of the overall reaction was reduced by 35%. This allows less tannin to be used in each sample.

The temperature of the first water bath needs to be exactly 37 C. The second water bath was originally described as an 80 C bath, but a boiling water bath (~ 100 C) gives better reproducibility.

When the ascorbate and/or the EDTA are omitted, water is used to replace the missing volume. The volume before the TBA and the TCA are added needs to be 420 μL.

The order of addition of the reagents given in the Aruoma method is changed for our experiments. In the

Aruoma method, the order of addition is FeCl₃, EDTA, buffer, water, H₂O₂, tannin, dOR, and Asc.

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0 TANNINS AS METAL ION CHELATORS

TANNINS AS METAL ION CHELATORS

Phenolics can affect the biological availability or activity of metal ions by chelating the metal (McDonald, M.; Mila, I.; Scalbert, A. J. Agric. Food Chem. 1996, 44, 599). Chelation requires appropriate patterns of substitution and a pH above the pKa of the phenolic group. Bacterial siderophores with multiple phenolic groups and very high affinities for essential metals such as iron have been characterized (Harris, W.R.; Carrano, C.J.; Cooper, S.R.; Sofen, S.R.; Avdeef, A.E.; McArdle, J.V.; Raymond, K.N. J. Am. Chem. Soc. 1979, 101, 6097). The similarity between siderophore ortho-dihydroxy substitution pattern and the substitution patterns on condensed and hydrolyzable tannins suggests that tannins may also have very high affinities for metals.

Phenolic-metal ion complexes are often colored, and it has been suggested that characteristic colors can be used to identify specific arrangements of phenolic groups (Mole, S.; Waterman, P.G., Oecologia 1987, 72, 137-147). However, these methods have not been adequately tested and are not recommended.

It is widely believed that tannin-chelated metal ions are not bioavailable. For example, consumption of large quantities of tea or other tannin-rich foods is sometimes associated with deficiency diseases such as anemia (Baynes, R.D. and Bothwell, T.H. Ann Rev. Nutr. 1990 10, 133). In many ecosystems, the slow decomposition of tannin-rich leaves has been attributed in part to the low levels of biologically available metal ions (Vituosek, P.M.; Turner, D.R.; Parton, W.J. and Sanford, R.L. Ecology 1994, 75, 418). The populations of microfauna essential to leaf decomposition and soil formation are unable to grow when metals are unavailable.

Metal ion chelation can alter the redox potential of the metal, or prevent its participation in redox reactions. Thus metal ion chelators can be inhibitors or enhancers of Fenton-driven oxidative reactions.

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0 BIOLOGICAL ACTIVITIES OF TANNINS

BIOLOGICAL ACTIVITIES OF TANNINS

Tannins have diverse effects on biological systems because they are potential metal ion chelators, protein precipitating agents, and biological antioxidants. Because tannins can play such varied biological roles, and because of the enormous structural variation among tannins, it has been difficult to develop models which allow accurate prediction of the effects of tannins in any system. An important goal of future work on the biological activities of tannins is the development of structure/activity relationships so that biological activities can be predicted.

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0 HYDROLYZABLE TANNIN STRUCTURAL CHEMISTRY

HYDROLYZABLE TANNIN STRUCTURAL CHEMISTRY

Hydrolyzable tannins are derivatives of gallic acid (3, 4, 5-trihydroxyl benzoic acid). Gallic acid is esterified to a core polyol, and the galloyl groups may be further esterified or oxidatively crosslinked to yield more complex hydrolysable tannins.

Early work on hydrolyzable tannins included Haslam’s significant elucidations of the structures of the simple gallotannins (Haslam, E. Plant polyphenols. Vegetable tannins revisited, ed.; Cambridge University Press: Cambridge, U. K., 1989). More recently, Okuda et al.(Okuda, T.; Yoshida, T.; Hatano, T. Hydrolyzable tannins and related polyphenols. Progress in the Chemistry of Organic Natural Products 1995, 66, 1-117) have been particularly active in characterization and classification of complex hydrolyzable tannins. Feldman’s synthetic work (Feldman KS, Lawlor MD, and Sahasrabudhe K Ellagitannin chemistry. Evolution of a threecomponent coupling strategy for the synthesis of the dimeric ellagitannin coriariin A and a dimeric gallotannin analogue. 2000; 8011-9) has provided useful insights into likely biosynthetic routes for the complex hydrolyzable tannins. A limited survey of structures and their relationships is provided here.

Gallotannins.

The simplest hydrolyzable tannins, the gallotannins, are simple polygalloyl esters of glucose.

The prototypical gallotannin is pentagalloyl glucose (β-1,2,3,4,6-Pentagalloyl-O-DGlucopyranose). Pentagalloyl glucose, or PGG, has five identical ester linkages that involve aliphatic hydroxyl groups of the core sugar. The alpha anomer is not common in nature.

Like all of the gallotannins, PGG has many isomers. The molecular weights of all the isomers of PGG are the same (940 g/mol), but chemical properties such as susceptibility to hydrolysis and chromatographic behavior; and biochemical properties such as ability to precipitate protein; are structure-dependent. 

The polygalloyl ester chains found in gallotannins are formed by either meta- or para-depside bonds, involving a phenolic hydroxyl rather than an aliphatic hydroxyl group. The depside bond is more easily hydrolyzed than an aliphatic ester bond. Methanolysis in weak acid in methanol breaks depside but not ester bonds. Thus the core polyol with its esterified galloyl groups can be produced from complex mixtures of polygalloyl esters by methanolysis with acetate buffer. Strong mineral acid, heat and methanol can be used to methanolzye both despide and ester bonds yielding the core polyol and methyl gallate. Hydrolysis with strong acid converts gallotannins to gallic acid and the core polyol.

Simple gallotannins with up to 12 esterifed galloyl groups and a core glucose are routinely found in tannins from sumac or oak galls. Commercial tannic acid is comprised of mixtures of gallotannins from sumac (Rhus semialata) galls (Chinese gallotannin); Aleppo oak (Quercus infectoria) galls (Turkish gallotannin); or sumac (R. coriaria, R. typhina) leaves (sumac gallotannin). Although commercial sources provide a nominal molecular weight for tannic acid (1294 g/mol), the preparations are heterogeneous and variable mixtures of galloyl esters. Tannic acid is not an appropriate standard for any tannin analysis because of its poorly defined composition. PGG can be prepared from some commercial tannic acids by methanolysis in acetate buffer. For the preparation to be successful, the tannic acid must have PGG as its core ester, most likely in preparations of Chinese or sumac gallotannin. Turkish gallotannin is comprised of esters of 1,2,3,6-tetragalloyl glucose; or 1,3,4,6-tetragalloyl glucose.

 

Although for many gallotannins glucose is the alcohol, other polyols including glucitol; hammamelose; shikimic acid; quinic acid; and quercitol; have been reported as constitutents of gallotannins from a few species. For example, aceritannin is found in leaves of several specie 

of maple (Acer), and hamamelitannin is found in bark of witch hazel (Hamamelis virginiana), oak (Quercus rubra), and several chestnut species (Castanea sp.).

Ellagitannins.

Oxidative coupling of galloyl groups converts gallotannins to the related ellagitannins. The simple ellagitannins are esters of hexahydroxydiphenic acid (HHDP). HHDP spontaneously lactonizes to ellagic acid in aqueous solution 

Intramolecular carbon-carbon coupling to form HHDP is most common between C-4/C-6 (e.g. eugeniin); and C-2/C-3 (e.g. casuarictin, also has C-4/C-6), as would be expected for polygalloyl glucoses in the more stable 4C₁ conformation. However, in a few plants intramolecular coupling 

occurs at C-3/C-6 (e.g. corilagin), C-2/C-4 (e.g. geraniin, also has C-3/C-6), or C-1/C-6 (e.g. davidiin), suggesting the polygalloyl glucose starting material was in the less stable₁C₄ 

conformation. Geraniin is further characterized by partial oxidation of the C-2/C-4 HHDP to dehydro-HHDP, and in aqueous solution several forms of dehydro-HHDP can be detected in geraniin by nmr.

In some plants including oak and chestnut the ellagitannins are further elaborated via ring opening. Thus after conversion of casuarictin to pedunculagin, the pyranose ring of the glucose opens and the family of compounds including casuariin, casuarinin, castalagin, and castlin; stachyurin, vescalagin and vescalin forms. 

The ellagitannins can undergo intermolecular oxidative coupling with other hydrolysable tannins to yield dimers. For example, in several euforbs (e.g. Euphorbia watanabei) geraniin oxidatively condenses with PGG to yield various euphrobins, characterized by the valoneoyl group.

Oenothein B, Woodfordin C, Cuphiin D1 and Eugeniflorin D1 are macrocyclic dimers linked by two valoneoyl groups, and the nobotanins are macrocyclic trimers.

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0 CONDENSED TANNIN STRUCTURAL CHEMISTRY

CONDENSED TANNIN STRUCTURAL CHEMISTRY

Proanthocyanidins (condensed tannins) are polymeric flavanoids. The flavanoids are a diverse group of metabolites based on a heterocyclic ring system derived from phenylalanine (B) and polyketide biosynthesis (A). Although the biosynthetic pathways for flavanoid synthesis are 

well understood, the steps leading to condensation and polymerization have not been elucidated.

The flavanoid skeleton, the standard letters to identify the rings, and the numbering system are shown here.

The most widely studied condensed tannins are based on the flavan-3-ols (-)-epicatechin and (+)-catechin. 

 Flavan-3-ols

Addition of a third phenolic group on the B ring yields epigallocatechin and gallocatechin. Much less common are flavan-3-ols with only a single phenolic group on the B ring, para to C-2 (epiafzelechin, afzelechin with stereochemistry corresponding to epicatechin, catechin respectively).

The best characterized condensed tannins are linked via a carbon-carbon bond between C8 of the terminal unit and C4 of the extender. The four common modes of coupling are illustrated by the dimers isolated by Haslam, and originally named B-1, B-2, B-3 and B-4. The more complete names specify the position and stereochemistry of the interflavan bond completely. In addition to these dimers, related dimers linked by C6 of the terminal unit and C4 of the extender have been isolated. 

Further polymerization can yield the linear 4,8 polymers such as the Sorghum procyanidin. Linear polymers based on 4,6 dimers; and branched dimers containing both 4,6 and 4,8 linkages are less common. 

Although the term condensed tannins is still widely used to describe these flavonoid-based polyphenolics, the chemically more descriptive term “proanthocyanidin” (NOT hydrolysis) in hot alcohols, e.g.via acid butanol chemistry. 

Anthocyanidins

The products of the acid butanol reaction are an unmodified terminal unit, and the colored anthocyanidins produced by the extender units. Catechin- and epicatechin-based polymers produce cyanidin, and thus are known as procyanidins. Gallocatechin and epigallocatechinbased polymers yield delphinidin, and the rare mono-substituted flavan-3-ol based polymers yield pelargonidin.

An important group of condensed tannins are 5-deoxy-flavan-3-ols polymers. Branching is common in these tannins, because of the reactivity of the 5-deoxy A ring. Profisetinidins and 

prorobinetinidins comprise the major tannins found in quebracho and acacia tannin preparations.

Acid butanol reaction yields the 5-deoxy anthocyanidins fisetinidn and robinetinidin. The acid butanol reaction can be carried out with a nucleophilic trapping agent to produce the terminal unit plus derivitized extender units. These can usually be separated and quantitated by HPLC to give composition and average molecular weight estimates for the parent tannin. Thiols such as toluene-α-thiol are often used in this reaction, but phloroglucinol is more convenient. The efficiency of the reaction with branched condensed tannins, especially the 5-deoxy-flavanolbased tannins, has not been established.

Another type of linkage that has been described but not studied extensively involves oxidative CO coupling between the flavonoid rings to yield A2 and related proanthocyanidins.

The flavan-3,4-diols, or luecoanthocyanidins, are sometimes confused with proanthocyanidins.

The flavan-3,4-diols are monomeric flavonoids that yield the anthocyanidins upon treatment with heat and acid. They thus have reactive chemistry similar to that of the condensed tannins, but they do not interact with protein to form precipitable complexes. 

The flavan-4-ols are also luecoanthocyanidins, but are unique in their lability. They yield the anthocyanidins upon treatment with alcoholic acid at room temperature. 

Stafford has suggested that pro-3-deoxyanthocyanidins might exist in a few plants. Evidence to date is limited to spectroscopy and some chemical conversions that are consistent with the chemistry shown here. (Stafford, H.A. Flavonoid Metabolism; CRC Press: Boca Raton, FL, 1990, pages 65-83).

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0 WHAT IS A TANNIN

WHAT IS A TANNIN?

Plants accumulate a wide variety of "secondary" compounds, including alkaloids, terpenes and phenolics. Although these compounds apparently do not function in "primary" metabolism such as biosynthesis, biodegradation and other energy conversions of intermediary metabolism, they do have diverse biological activities ranging from toxicity to hormonal mimicry, and may play a role in protecting plants from herbivory and disease.

Phenolic metabolism in plants is complex, and yields a wide array of compounds ranging from the familiar flower pigments (anthocyanidins) to the complex phenolics of the plant cell wall (lignin). However, the group of phenolic compounds known as tannins is clearly distinguished from other plant secondary phenolics in their chemical reactivities & biological activities.

Tradition use of tannins as agents for converting animal hides to leather ("tanning") is one manifestation of the most obvious activity of the tannins: their ability to interact with and precipitate proteins, including the proteins found in animal skin. The term "tannin" comes from the ancient Celtic word for oak, a typical source for tannins for leather making.

Bate-Smith defined tannins as "water-soluble phenolic compounds having molecular weights between 500 and 3000...[giving] the usual phenolic reactions...[and having] special properties such as the ability to precipiate alkaloids, gelatin and other proteins".

Haslam has more recently substituted the term "polyphenol" for "tannin", in an attempt to emphasize the multiplicity of phenolic groups characteristic of these compounds. He notes that molecular weights as high as 20,000 have been reported, and that tannins complex not only with proteins and alkaloids but also with certain polysaccharides. I prefer to use the term tannin, which emphasizes the character which sets tannins apart from all other phenolics: the ability to precipitate proteins.

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