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Biosynthesis of gallotannins: β-Glucogallin-dependent formation of 1,2,3,4,6-pentagalloylglucose by enzymatic galloylation of 1,2,3,6-tetragalloylglucose
ARCHIVES
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 273, No. 1, August 15, pp. 58-63,1989
Biosynthesis of Gallotannins: ,&Glucogallin-Dependent Formation of 1,2,3,4,6-Pentagalloylglucose by Enzymatic Galloylation of 1,2,3,6-Tetragalloylglucose’ JESSICA
CAMMANN,*
KLAUS DENZEL,* GERHARD GEORG G. GROSS*,’
SCHILLING,t
AND
*Univw&Tt Urn, Abteilung Allgemeine Botanik, Albert-Einstein-AUee 11, D-7900 Ulm, Federal Republic of Germany, and ~Unive-rsit& Heidelberg, Organ&h-Chemisches Institut, Im Neuenheimer Feld 270, O-6900 Heidelberg, Federal Republic of Germany Received February
13.1989, and in revised form April
17,1989
An acyltransferase was detected in young leaves of pedunculate oak (Quercus robur) that catalyzed the formation of 1,2,3,4,6-penta-0-galloyl-fl-D-glucose, the common precursor of gallotannins and the related ellagitannins. This enzyme depended on @glucogallin (l-O-gallOyl-B-D-glUCOSC) as acyl donor; 1,2,3,6-tetra-O-galloyl-@D-glucose was specifically required as acceptor molecule, whereas no reaction occurred with the 1,2,4,6isomer of this substrate. The partially purified enzyme (1M, 260,000) was stable between pH 5.0 and 6.5; highest activities were observed at pH 6.3 and 40%. Km values of 2:3 and 1.0 mM, respectively, were determined for the substrates /?-glucogallin and tetragalloylglucose. In accordance with stoichiometric studies, the systematic name “B-glucogallin: 1,2,3,6-tetra-0-galloylglucose 4-0-galloyltransferase” is proposed for this new enzyme. 0 1989 Academic Press. Inc.
Hydrolyzable plant tannins are polyphenolic natural products which are characterized by a central polyol moiety (mostly @-D-glucopyranose) whose hydroxyl groups are esterified with gallic acid (3,4,5trihydroxybenzoic acid). In the case of the gallotannins proper, further galloyl residues are attached to this core structure via meta-depside linkages. The related ellagitannins, in contrast, are the result of dehydrogenative dimerization reactions between spatially adjacent galloyl residues, thus forming the hexahydroxy-2,2’-diphenoyl residues characteristic of such compounds. Innumerable variations of these fundamental structural principles have been realized in the plant kingdom, and the well-documented detailed knowledge
of the chemical configuration and natural distribution of hydrolyzable tannins prompted considerations on their biosynthesis [reviewed, e.g., in (l)]. It was proposed on this basis that both gallotannins and ellagitannins are formed via 12t ,3,4 ,6penta-0-galloyl+D-glucose as the common and central precursor. Recent enzymatic studies in our laboratory have shown that the biogenetic pathway leading to this pivotal ester is initiated by the formation of P-glucogallin (l-O-galloyl-~-Dglucose) from free gallic acid and UDPglucose under the catalysis of a specific glucosyltransferase (2-4). Further experiments revealed that the group-transfer potential of this monogalloyl ester is sufficiently high to permit various subsequent transacylation reactions (5). One of these was an exchange reaction of still unknown physiological significance that could be demonstrated only by incubating p-glucogallin together with labeled glucose (6,7). Of greater relevance was the detection of
1 This work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. ’ To whom correspondence should be addressed. 0003-9861/89 $3.00 Copyright All rights
0 1989 hy Academic Press. Inc. of reproduction in any form reserved.
58
BIOSYNTHESIS
OF PENTAGALLOYLGLUCOSE
different galloyltransferases that catalyzed the formation of 1,6-digalloylglucose (8) and 1,2,6-trigalloylglucose (9) in the expected manner, i.e., by utilizing ,&glucogallin as a specific acyl donor. In continuation of these studies we were able to isolate an acyltransferase that specifically converts 1,2,3,6-tetragalloylglucose to 1,2,3,4,6-pentalgalloylglucose and that again utilizes @glucogallin as galloyl donor. The partial purification and some important properties of this new enzyme are reported in this communication. MATERIALS
AND METHODS
Chemicals. @-Glueogallin was synthesized from 2,3,4,6-tetra-o-acetyl+D-glucose and tri-O-acetylgalloylchloride (2); 1,2,3,4,6-penta-0-galloyl-&D-glucase was prepared via tri-0-acetylgalloylchloride and B-D-glucose (5), ,&-[U-‘4C-gZucosy~Glucogallin was synthesized enzymatically (7). Reference samples of 1,2,3,6- and 1,2,4,6-tetra-o-galloyl-/3-D-glucose were generous gifts of Professor G. Nonaka (Kyushu University, Fukuoka, Japan) and Professor E. Haslam (University of Sheffield, England), respectively. Greater quantities of 1,2,3,6-tetragalloylglucose required as substrate in this investigation were isolated from commercially available tannin (Carl Roth KG, Karlsruhe, FRG). This crude product was prepurified in three 10-g portions by column chromatography on Sephadex LH-20 (Pharmacia, Freiburg, FRG, 30 X 4 cm i.d.) in ethanol (10-12). After being washed with 1 liter solvent, the desired ester was eluted with further 2.5 liters ethanol in about 40% purity as determined by RP-HPLC (cf. Analytical methods). The residue remaining after vacuum evaporation at 40°C was dissolved in methanol (1 g/10 ml), filtered through l-pm membrane filters (RC-60, Schleicher & Schuell, Dassel, FRG), and subjected in 200-mg portions to preparative HPLC on LiChrosorb RP-18 (Merck, Darmstadt, FRG, particle size 5 pm, column 30 X 2 cm i.d.; solvent 18% acetonitrile in 0.05% acetic acid, flow rate 1’7ml/min). After rechromatography in 16% acetonitrile (other conditions as above), 240 mg of pure (>98%) 1,2,3,6-tetra-Ogalloyl-p-D-glucase was obtained as shown by analytical HPLC (see Analytical methods) and ‘H NMR spectroscopy in DMS03 [a 6.99, 6.91,6.89,6.80 (s, each 2 H, galloyl H2,6), 6.10 (d, H-l,3&.12 = 8.4 Hz), 5.50 (t, H-3), 5.27 (t, H-2), -4.40 (m, H-6,6’), -4,05 (m, H-5), 3.80 (t, H-4)]. Enzyme preparatim Unless stated otherwise, all operations were done at 0-4”C, and all buffers were
’ Abbreviations aq, aqueous.
used: DMSO, dimethyl sulfoxide;
59
supplemented with 5 mM 2-mercaptoethanol. Protein concentrations were determined turbidimetrically after precipitation with trichloroacetic acid (13), using bovine serum albumin as standard. Leaves (80 g) from 3- to 5-month-old Quercus ro&r (pedunculate oak) plantlets grown in the greenhouse were washed, frozen in liquid nitrogen, and homogenized in a precooled ultracentrifugal mill (Retsch KG, Haan, FRG). The frozen powder was immediately mixed with 80 g (wet weight) prewashed insoluble polyvinylpyrrolidone (polyclar AT, Sigma, Munich, FRG) and stirred for 30 min with a mixture of 150 ml 0.1 M borate buffer, pH 7.5, and 100 ml 1 M Tris-HCl buffer, pH 8.0. During the initial phases of the extraction, the pH of the homogenate had to be readjusted occasionally with Tris buffer. (Smaller amounts of leaf material were conveniently homogenized in a chilled mortar with quartz sand under analogous conditions.) The homogenate was squeezed through four layers of muslin and centrifuged (20 min, 35,OOOg). The supernatant of the crude extract was fractionated with solid ammonium sulfate. The 3560% precipitate was redissolved in a minimum of 50 mM TrisHCl, pH 7.5, and desalted by gel filtration on Sephadex G-25 (PD-10 columns; Pharmacia). This solution was adsorbed on a DEAE 52-cellulose column (Serva, Heidelberg, FRG, 4 X 1.6 cm i.d.) equilibrated in the same buffer. After being washed with 0.15 M KC1 in buffer, the enzyme was eluted with 0.3 M KCI; the most active fractions were combined, desalted, and stored at 0-4OC for l-2 weeks without significant losses of activity. In a typical experiment the resulting enzyme preparation was 22-fold purified in 37% yield and contained 7.6 mg protein with a specific activity of 96 pkat/mg. Enzyme assay. Standard assay mixtures (volume 50~1) containing 10 rmol potassium phosphate buffer, pH 6.5, 50 nmol 1,2,3,6-tetragalloylglucose, 300 nmol /3-glucogallin, and suitable amounts of protein (ca. 0.05-0.3 mg) were incubated at 30°C for 30-120 min (corresponding blanks were stopped at 0 min). After the enzyme reaction was terminated with 20 ~1 1 N HCl, the mixture was centrifuged and the supernatant carefully removed. Surprisingly, this fraction contained most of the unreacted @glucogallin, but only traces of both residual tetragalloylglucose, and produced pentagalloylglucose. These latter compounds were, due to their extreme tanning capacity (14,15), found tightly associated with the denatured protein of the pellet from which they could be released in 95-98% yield by ultrasonicating in 50% aq methanol (2 X 70 ~1) and centrifugation. The amount of enzymatically formed pentagalloylglucose in the combined three supernatants (volume 210 ~1)was determined by analytical HPLC as described under Analytical methods. Reaction-product identification Scaled-up assay mixtures (volume 150 ml) containing 50 mg (63 pmol) tetragalloylglucose, 100 mg (300 pmol) p-glucogallin,
60
CAMMANN
ET AL.
B
A
1
0
I
1
10
1
1
20
1
I
LI
I
30
0
10
I
I
I
20
I
30
Retention Time (min) FIG. 1. HPLC analysis of 4-0-galloyltransferase assays. (A) Control with heat-denatured enzyme; (B) complete assay mixture. (1) @-Glucogallin; (2) gallic acid; (3) 1,2,3,6-tetragalloylglucose; (4) 1,2,3,4,6-pentagalloylglucose. Aliquots of deproteinized standard assay mixtures were chromatographed on RP-18 columns under the conditions employed in stoichiometric studies (cf. Materials and Methods).
30 mmol phosphate buffer, pH 6.5, and 30 mg protein were incubated at 30°C for 60 min. After precipitation of protein with 20 ml 1 N HCl and centrifugation (10 min, 2O,OOOg), the supernatant was collected, the pellet was washed by ultrasonication with 50% aq methanol (5 X 20 ml) and centrifugation. The combined six supernatants were concentrated by rotary evaporation (40°C) and lyophilized. The remaining solid was dissolved in 5% aq methanol (20 mg/ml), filtered (1 pm pore size), and purified by preparative HPLC (Merck LiChrosorb RP-185 pm, column 30 X 2 cm i.d.; solvent 40% methanol in 0.05% acetic acid; flow rate 12 ml/min). After rechromatography under the same conditions, the pure reaction product was concentrated in vacua, lyophilized (yield 5 mg), and analyzed by cochromatography with authentic references (HPLC conditions under Analytical methods). Definitive proof of the enzyme product as 1,2,3,4,6penta-o-galloyl-@-D-glucopyranose was obtained by ‘H NMR spectroscopy in DMSO: d 6.98,6.92,6.86,6.82, 6.78 (s, each 2 H, galloyl H-2,6), 6.35 (d, H-l, aJnl,z = 8.1 Hz), 5.92 (t, H-3), -5.44 (m, H-2, H-4), -4.55 (m, H-5), -4.32 (m, H-6,6’). These values were identical with those obtained from chemically prepared reference material. Determination of the reaction stoichiomxtrg. Standard enzyme assays, modified by addition of @-[U-14CgZucosyl]glucogallin (2 kBq, 120 nmol), were incubated and worked up as described above (cf. Enzyme assay).
Aliquots of the combined supernatants were analyzed by RP-HPLC under conditions allowing the separation of all substrates and products involved in the reaction, including relevant by-products (Merck Liloo-
602 .% X .g
60-
0 0) .g m
40-
if
20-
01 3
I
I
1
5
I 7
,
\ 0, 9
PH
FIG. 2. Effects of pH on activity and stability of 40-galloyltransferase. The activity of the enzyme was measured under standard assay conditions at the indicated pH values. To test the stability, the enzyme was exposed for 30 min at 30°C to the pH values given, with subsequent determination of the residual activity at pH 6.5 (-) pH optimum; (---) stability. Buffers: (A) Sodium acetate; (0) potassium phosphate, (0) Tris-HCl.
BIOSYNTHESIS
61
OF PENTAGALLGYLGLUCOSE
30
A
Ho oP'o
20
P: /
v ,/
0.05
8
0
a
.rp'*
10 LOO
0.5 l/S
0l 0
-0
5
Substrate
concentration,
1
2
S (mM)
FIG. 3. Substrate saturation curves for /3-glucogallin (A) and 1,2,3,6-tetragalloylglucose (B).
Chrosorb RP-18,5 pm, column 180 X 3 mm i.d.; solvent A methanol; solvent B 0.05% HaPO,; gradient O-7 min = 7% A,7-20 min = ‘7-50% A,20-25 min = 50-90% A; flow rate 0.8 ml/min). Quantification was done by uv photometry at 280 nm (aromatic compounds) or by radioactivity measurements after fractionation of the HPLC eluates (glucose, &glucogallin). The resulting data were corrected against blanks with heat-denatured enzyme. Analytical methods. Determination of enzymatitally formed pentagalloylglucose, as well as estimations of the progress in the purification of the substrate tetragalloylglucose, were carried out by analytical HPLC on Merck LiChrosorb RP-18 columns (180 X 3 mm i.d., particle size 5 pm). The solvent systems used were modifications of that given in (12), consisting of acetonitrile (A) and 0.05% H3P04 (B). In routine enzyme activity assays, an efficient separation of tetragalloylglucose (retention time Rt = 10.2 min) and pentagalloylglucose (Rt = 11.70 min) was achieved by a linear gradient of lo-80% A within 22.5 min. The purification of tetragalloylglucose from crude tannin was monitored by a linear gradient of O40% A within 20 min, resulting in a Rt value of 13.5 min. A flow rate of 1.0 ml/min was maintained in all these experiments. Quantitative data were obtained by uv photometry at 280 nm, in combination with a computing integrator (Merck-Hitachi D-2000) referring to external standard solutions of the investigated compounds. Galloylglucoses to be analyzed by ‘H NMR spectroscopy were dried in DUCUO (5 mm Hg) at 85°C over PzOsand KOH pellets for 2 days and dissolved in DMSO-&. Spectra were recorded with a Bruker AM 200 at 200 MHz, using tetramethylsilane as internal standard. Assignment of glucose proton sequences was achieved by spin-decoupling experiments.
RESULTS AND DISCUSSION
Enzyme pur$cat&m. 4-O-Galloyltransferase was extracted from leaves of young oak plants grown in the greenhouse which were still devoid of the viscous mucilage characteristic of older leaves that had severely hampered some previous investigations (3,5). To ensure a continuous supply of suitable plant material, the stems of older plantlets were cut to enforce the subsequent development of new shoots bearing fresh leaves of the required age. (Unavoidable gaps could be bridged by using leaves stored at -20°C.) The crude extracts obtained from this plant material were partially purified by ammonium sulfate precipitation and anion-exchange chromatography (cf. Material and Methods). The resulting enzyme preparations were sufficiently pure, except in the stoichiometry experiments where the effects of a contaminating esterase had to be considered. General properties of the galloyltransferase. The synthesis of pentagalloylglucose was strictly dependent on the presence of both /3-glucogallin and 1,2,3,6-tetragalloylglucose, and no reaction occurred in samples with heat-denatured enzyme (Fig. 1). When isomeric 1,2,4,6-tetragalloylglucose was assayed as a potential substrate, only negligible activity (1.1%) was found. Under standard assay conditions, a roughly linear progress of the enzymatic reaction
62
CAMMANN
ET AL.
TABLE I STOICHIOMETRYOFTHE~-O-GALLO~RANSFERASE-CATALYZEDFORMATIONOFPENTAGALLOYLGLUCOSE"
Component assayed
Gross reaction rate (nmol)
Corrected reaction rate* (nmol)
Molar ratio”
P-Glucogallin, consumed 1,2,3,6-Tetragalloylglucose, consumed 12 t I3I476-Pentagalloylglucose, formed Glucose, formed Gallic acid, formed
28.4 8.7 8.9 28.4 20.0
8.4 8.7 8.9 8.4 0
1.0 1.0 1.0 1.0 -
a Experimental details are described under Materials and Methods. bIt is assumed that gallic acid is produced by hydrolysis of the substrate /3-glucogallin, liberating concomitantly an equal amount of glucose. ‘Values refer to the reaction rate determined for tetragalloylglucose.
was observed with respect to time for ca. ‘70 min, gradually declining thereafter with a maximum at about 100 min. In contrast, a nonlinear relationship to the amount of added protein occurred; it is assumed that this anomaly is caused by the pronounced tanning (i.e., protein-binding and precipitating) capacities of tetra- and pentagalloylglucose in the assay mixtures (14,X). The effect of pH on the stability of the enzyme and on the velocity of the reaction is depicted in Fig. 2. The optimal pH was 6.3; maximal stability was found between pH 5.0 and 6.5. The temperature optimum of the reaction was at ea. 40°C. The rather unusual observation that the enzyme exhibited some activity even at 0°C parallels previous observations with related enzymes from oak (8) and sumac (9). Between 25 and 35%, a Q10value of 2.0 and an activation energy of 28.8 kJ/mol were calculated.
OH
The enzyme showed normal MichelisMenten kinetics with both substrates (Fig. 3). Replots of these data according to Lineweaver-Burk gave Km values of 2.3 and 1.0 IIIM for B-glucogallin and tetragalloylglucase, respectively. From gel-filtration experiments with a calibrated Sephadex G-200 column (16), an apparent molecular weight of 260,000 was estimated for the enzyme. This comparatively high value is consistent with those determined for other galloyltransferases from oak (6,8), but in clear contrast to the A& 75,000 for a related trigalloylglucosesynthesizing enzyme (9) from sumac leaves (unpublished data). Reaction-product identification. The enzyme reaction product was tentatively identified as 1,2,3,4,6-penta-Q-galloyl-@Dglucopyranose by cochromatography with authentic references (cf. Fig. 1). This conclusion was unequivocally confirmed by ‘H
+DG
FIG. 4. Reaction equation for the formation of pentagalloylglucose by galloylation of tetragalloylglucose. j3G, &Glucogallin, Glc, glucose.
BIOSYNTHESIS
63
OF PENTAGALLOYLGLUCOSE
NMR spectroscopy of material that had been isolated and purified from scaled-up assay mixtures, including D20 exchange, spin-decoupling experiments, and comparison with data from pure compounds (for details, see Materials and Methods). The recorded spectrum showed five singlets (6 6.78-6.89) corresponding to the five galloyl residues of the product, and the signals for all of the glucose protons were shifted downfield in response to acylation. P-Configuration of the product was proven by a coupling constant 3JH-1,2 of 8 Hz; other coupling constants for H-l to H-6,6’ (J ca. 10 Hz) showed that the glucose moiety had the expected normal 4C1-chair eonfiguration. Stoichicvmetrg Most likely, 1 mol of /3glucogallin serves as galloyl donor for the acylation of 1 mol of tetragalloylglucose under the concomitant production of 1 mol each of pentagalloylglucose and free glucose. Experimental proof was complicated, however, by the contamination of the enzyme preparation with an esterase activity that effectively hydrolyzed the substrate P-glucogallin. Assuming that gallic acid found in the assay mixtures was exclusively formed by this process, the amount of this by-product could serve to correct the determined concentrations of 8-glucogallin and glucose. Under this premise, and as summarized in Table I, it is evident that the observed turnover of substrates and products was fully consistent with the expected reaction depicted in Fig. 4. According to this equation, the systematic name “P-glucogallin: 1,2,3,6-tetra-O-galloyl$-Dglucose 4Ggalloyltransferase” (EC 2.3.1.-) is proposed for this new enzyme. In conclusion, earlier postulates (5,8) on the role of P-glucogallin as the principal acyl donor in the pathway from gallic acid to pentagalloylglucose have been confirmed by this investigation. Moreover, the precise structure of all the intermediates along this sequence is now known, a fact that must be attributed mainly to recent enzyme studies [cf. (17)]. Only one step, i.e., the supposed acylation of 1,2,6-trigalloyl-
glucose to 1,2,3,6-tetragalloylglucose, has not yet been fully characterized by this means; however, analyses of the galloylglucoses formed by oak tissues in tivo (18) or in vitro (8) are fully consistent with this proposal. ACKNOWLEDGMENTS We thank Miss U. Semler for expert technical assistance, and Professors G. Nonaka (Fukuoka) and E. Haslam (Sheffield) for providing reference substances. REFERENCES 1. HASLAM, E. (1982) Fartschr. Chem. Org. N&u&. 41,1-46. 2. GROSS,G. G. (1982) FEBSLett. 14?3,67-70. 22,2179-2182. 3. GROSS,G. G. (1983) Phytochemistry 4. WEISEMANN, S., DENZEL, K., SCHELING, G., AND GROSS,G. G. (1988) Bioorg. Chem l&29-37. 5. GROSS,G. G. (1983) Z. Nuturfwsck C38,519-523. 6. GROSS, G. G., SCHMIDT, S. W., AND DENZEL, K. (1986) J. Plant Physid 126,1’73-179. 7. DENZEL, K., WEISEMANN, S., AND GROSS, G. G. (1988) J. Plant Physiol. 133,113-115. 8. SCHMIDT, S. W., DENZEL, K., SCHILLING,G., AND GROSS, G. G. (1987) Z. Nuturforsch
C42,87-92.
9. DENZEL, K., SCHILLING, G., AND GROSS, G. G. (1988) PZuntu 176,135-137.
10. NISHIZAWA, M., YAMAGISHI, T., NONAKA, G., AND NISHIOKA, I. (1980) Chem Phcwm. BulL 28, 2850-2852. 11. NISHIZAWA, M.,YAMAGISHI, T., NONAKA, G., AND NISHIOKA, I. (1983) J. Chem. Sot. Perkin Trans.
1,961-965. 12. HADDOCK, E. A., GUPTA, R. K., AL-SHAFI, S. M. K., AND HASLAM, E. (1982) .J. Chem 5’~. Perkin
Trans. 1,2515-2524.
13. KUNITZ, M. (1952) J. Gen Physid 35,423-450. 14. HASLAM, E., AND LILLEY, T. H. (1985) in The Biochemistry of Plant Phenol& (Van Sumere, C. F., and Lea, P. J., Eds.), Annu. Proc. Phytothem. Sot. Eur.,Vol.25,pp. 237-256, Clarendon Press, Oxford. 15. OZAWA, T., LILLEY, T. H., AND HASLAM, E. (1987) Phytochxmistry
26,2937-2942.
16. ANDREWS, P. (1965) B&hem, .I 96,595-606. 17. GROSS, G. G. (1989) in Plant Cell Wall Polymers: Biogenesis and Biodegradation (Lewis, N. G., and Paice, M. G., Eds.), ACS Symp. Ser. Vol. 399, Washington, DC, in press. 18. KRAJCI, I., AND GROSS,G. G. (1987) Phytochemistry 26,141-143.
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 273, No. 1, August 15, pp. 58-63,1989
Biosynthesis of Gallotannins: ,&Glucogallin-Dependent Formation of 1,2,3,4,6-Pentagalloylglucose by Enzymatic Galloylation of 1,2,3,6-Tetragalloylglucose’ JESSICA
CAMMANN,*
KLAUS DENZEL,* GERHARD GEORG G. GROSS*,’
SCHILLING,t
AND
*Univw&Tt Urn, Abteilung Allgemeine Botanik, Albert-Einstein-AUee 11, D-7900 Ulm, Federal Republic of Germany, and ~Unive-rsit& Heidelberg, Organ&h-Chemisches Institut, Im Neuenheimer Feld 270, O-6900 Heidelberg, Federal Republic of Germany Received February
13.1989, and in revised form April
17,1989
An acyltransferase was detected in young leaves of pedunculate oak (Quercus robur) that catalyzed the formation of 1,2,3,4,6-penta-0-galloyl-fl-D-glucose, the common precursor of gallotannins and the related ellagitannins. This enzyme depended on @glucogallin (l-O-gallOyl-B-D-glUCOSC) as acyl donor; 1,2,3,6-tetra-O-galloyl-@D-glucose was specifically required as acceptor molecule, whereas no reaction occurred with the 1,2,4,6isomer of this substrate. The partially purified enzyme (1M, 260,000) was stable between pH 5.0 and 6.5; highest activities were observed at pH 6.3 and 40%. Km values of 2:3 and 1.0 mM, respectively, were determined for the substrates /?-glucogallin and tetragalloylglucose. In accordance with stoichiometric studies, the systematic name “B-glucogallin: 1,2,3,6-tetra-0-galloylglucose 4-0-galloyltransferase” is proposed for this new enzyme. 0 1989 Academic Press. Inc.
Hydrolyzable plant tannins are polyphenolic natural products which are characterized by a central polyol moiety (mostly @-D-glucopyranose) whose hydroxyl groups are esterified with gallic acid (3,4,5trihydroxybenzoic acid). In the case of the gallotannins proper, further galloyl residues are attached to this core structure via meta-depside linkages. The related ellagitannins, in contrast, are the result of dehydrogenative dimerization reactions between spatially adjacent galloyl residues, thus forming the hexahydroxy-2,2’-diphenoyl residues characteristic of such compounds. Innumerable variations of these fundamental structural principles have been realized in the plant kingdom, and the well-documented detailed knowledge
of the chemical configuration and natural distribution of hydrolyzable tannins prompted considerations on their biosynthesis [reviewed, e.g., in (l)]. It was proposed on this basis that both gallotannins and ellagitannins are formed via 12t ,3,4 ,6penta-0-galloyl+D-glucose as the common and central precursor. Recent enzymatic studies in our laboratory have shown that the biogenetic pathway leading to this pivotal ester is initiated by the formation of P-glucogallin (l-O-galloyl-~-Dglucose) from free gallic acid and UDPglucose under the catalysis of a specific glucosyltransferase (2-4). Further experiments revealed that the group-transfer potential of this monogalloyl ester is sufficiently high to permit various subsequent transacylation reactions (5). One of these was an exchange reaction of still unknown physiological significance that could be demonstrated only by incubating p-glucogallin together with labeled glucose (6,7). Of greater relevance was the detection of
1 This work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. ’ To whom correspondence should be addressed. 0003-9861/89 $3.00 Copyright All rights
0 1989 hy Academic Press. Inc. of reproduction in any form reserved.
58
BIOSYNTHESIS
OF PENTAGALLOYLGLUCOSE
different galloyltransferases that catalyzed the formation of 1,6-digalloylglucose (8) and 1,2,6-trigalloylglucose (9) in the expected manner, i.e., by utilizing ,&glucogallin as a specific acyl donor. In continuation of these studies we were able to isolate an acyltransferase that specifically converts 1,2,3,6-tetragalloylglucose to 1,2,3,4,6-pentalgalloylglucose and that again utilizes @glucogallin as galloyl donor. The partial purification and some important properties of this new enzyme are reported in this communication. MATERIALS
AND METHODS
Chemicals. @-Glueogallin was synthesized from 2,3,4,6-tetra-o-acetyl+D-glucose and tri-O-acetylgalloylchloride (2); 1,2,3,4,6-penta-0-galloyl-&D-glucase was prepared via tri-0-acetylgalloylchloride and B-D-glucose (5), ,&-[U-‘4C-gZucosy~Glucogallin was synthesized enzymatically (7). Reference samples of 1,2,3,6- and 1,2,4,6-tetra-o-galloyl-/3-D-glucose were generous gifts of Professor G. Nonaka (Kyushu University, Fukuoka, Japan) and Professor E. Haslam (University of Sheffield, England), respectively. Greater quantities of 1,2,3,6-tetragalloylglucose required as substrate in this investigation were isolated from commercially available tannin (Carl Roth KG, Karlsruhe, FRG). This crude product was prepurified in three 10-g portions by column chromatography on Sephadex LH-20 (Pharmacia, Freiburg, FRG, 30 X 4 cm i.d.) in ethanol (10-12). After being washed with 1 liter solvent, the desired ester was eluted with further 2.5 liters ethanol in about 40% purity as determined by RP-HPLC (cf. Analytical methods). The residue remaining after vacuum evaporation at 40°C was dissolved in methanol (1 g/10 ml), filtered through l-pm membrane filters (RC-60, Schleicher & Schuell, Dassel, FRG), and subjected in 200-mg portions to preparative HPLC on LiChrosorb RP-18 (Merck, Darmstadt, FRG, particle size 5 pm, column 30 X 2 cm i.d.; solvent 18% acetonitrile in 0.05% acetic acid, flow rate 1’7ml/min). After rechromatography in 16% acetonitrile (other conditions as above), 240 mg of pure (>98%) 1,2,3,6-tetra-Ogalloyl-p-D-glucase was obtained as shown by analytical HPLC (see Analytical methods) and ‘H NMR spectroscopy in DMS03 [a 6.99, 6.91,6.89,6.80 (s, each 2 H, galloyl H2,6), 6.10 (d, H-l,3&.12 = 8.4 Hz), 5.50 (t, H-3), 5.27 (t, H-2), -4.40 (m, H-6,6’), -4,05 (m, H-5), 3.80 (t, H-4)]. Enzyme preparatim Unless stated otherwise, all operations were done at 0-4”C, and all buffers were
’ Abbreviations aq, aqueous.
used: DMSO, dimethyl sulfoxide;
59
supplemented with 5 mM 2-mercaptoethanol. Protein concentrations were determined turbidimetrically after precipitation with trichloroacetic acid (13), using bovine serum albumin as standard. Leaves (80 g) from 3- to 5-month-old Quercus ro&r (pedunculate oak) plantlets grown in the greenhouse were washed, frozen in liquid nitrogen, and homogenized in a precooled ultracentrifugal mill (Retsch KG, Haan, FRG). The frozen powder was immediately mixed with 80 g (wet weight) prewashed insoluble polyvinylpyrrolidone (polyclar AT, Sigma, Munich, FRG) and stirred for 30 min with a mixture of 150 ml 0.1 M borate buffer, pH 7.5, and 100 ml 1 M Tris-HCl buffer, pH 8.0. During the initial phases of the extraction, the pH of the homogenate had to be readjusted occasionally with Tris buffer. (Smaller amounts of leaf material were conveniently homogenized in a chilled mortar with quartz sand under analogous conditions.) The homogenate was squeezed through four layers of muslin and centrifuged (20 min, 35,OOOg). The supernatant of the crude extract was fractionated with solid ammonium sulfate. The 3560% precipitate was redissolved in a minimum of 50 mM TrisHCl, pH 7.5, and desalted by gel filtration on Sephadex G-25 (PD-10 columns; Pharmacia). This solution was adsorbed on a DEAE 52-cellulose column (Serva, Heidelberg, FRG, 4 X 1.6 cm i.d.) equilibrated in the same buffer. After being washed with 0.15 M KC1 in buffer, the enzyme was eluted with 0.3 M KCI; the most active fractions were combined, desalted, and stored at 0-4OC for l-2 weeks without significant losses of activity. In a typical experiment the resulting enzyme preparation was 22-fold purified in 37% yield and contained 7.6 mg protein with a specific activity of 96 pkat/mg. Enzyme assay. Standard assay mixtures (volume 50~1) containing 10 rmol potassium phosphate buffer, pH 6.5, 50 nmol 1,2,3,6-tetragalloylglucose, 300 nmol /3-glucogallin, and suitable amounts of protein (ca. 0.05-0.3 mg) were incubated at 30°C for 30-120 min (corresponding blanks were stopped at 0 min). After the enzyme reaction was terminated with 20 ~1 1 N HCl, the mixture was centrifuged and the supernatant carefully removed. Surprisingly, this fraction contained most of the unreacted @glucogallin, but only traces of both residual tetragalloylglucose, and produced pentagalloylglucose. These latter compounds were, due to their extreme tanning capacity (14,15), found tightly associated with the denatured protein of the pellet from which they could be released in 95-98% yield by ultrasonicating in 50% aq methanol (2 X 70 ~1) and centrifugation. The amount of enzymatically formed pentagalloylglucose in the combined three supernatants (volume 210 ~1)was determined by analytical HPLC as described under Analytical methods. Reaction-product identification Scaled-up assay mixtures (volume 150 ml) containing 50 mg (63 pmol) tetragalloylglucose, 100 mg (300 pmol) p-glucogallin,
60
CAMMANN
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B
A
1
0
I
1
10
1
1
20
1
I
LI
I
30
0
10
I
I
I
20
I
30
Retention Time (min) FIG. 1. HPLC analysis of 4-0-galloyltransferase assays. (A) Control with heat-denatured enzyme; (B) complete assay mixture. (1) @-Glucogallin; (2) gallic acid; (3) 1,2,3,6-tetragalloylglucose; (4) 1,2,3,4,6-pentagalloylglucose. Aliquots of deproteinized standard assay mixtures were chromatographed on RP-18 columns under the conditions employed in stoichiometric studies (cf. Materials and Methods).
30 mmol phosphate buffer, pH 6.5, and 30 mg protein were incubated at 30°C for 60 min. After precipitation of protein with 20 ml 1 N HCl and centrifugation (10 min, 2O,OOOg), the supernatant was collected, the pellet was washed by ultrasonication with 50% aq methanol (5 X 20 ml) and centrifugation. The combined six supernatants were concentrated by rotary evaporation (40°C) and lyophilized. The remaining solid was dissolved in 5% aq methanol (20 mg/ml), filtered (1 pm pore size), and purified by preparative HPLC (Merck LiChrosorb RP-185 pm, column 30 X 2 cm i.d.; solvent 40% methanol in 0.05% acetic acid; flow rate 12 ml/min). After rechromatography under the same conditions, the pure reaction product was concentrated in vacua, lyophilized (yield 5 mg), and analyzed by cochromatography with authentic references (HPLC conditions under Analytical methods). Definitive proof of the enzyme product as 1,2,3,4,6penta-o-galloyl-@-D-glucopyranose was obtained by ‘H NMR spectroscopy in DMSO: d 6.98,6.92,6.86,6.82, 6.78 (s, each 2 H, galloyl H-2,6), 6.35 (d, H-l, aJnl,z = 8.1 Hz), 5.92 (t, H-3), -5.44 (m, H-2, H-4), -4.55 (m, H-5), -4.32 (m, H-6,6’). These values were identical with those obtained from chemically prepared reference material. Determination of the reaction stoichiomxtrg. Standard enzyme assays, modified by addition of @-[U-14CgZucosyl]glucogallin (2 kBq, 120 nmol), were incubated and worked up as described above (cf. Enzyme assay).
Aliquots of the combined supernatants were analyzed by RP-HPLC under conditions allowing the separation of all substrates and products involved in the reaction, including relevant by-products (Merck Liloo-
602 .% X .g
60-
0 0) .g m
40-
if
20-
01 3
I
I
1
5
I 7
,
\ 0, 9
PH
FIG. 2. Effects of pH on activity and stability of 40-galloyltransferase. The activity of the enzyme was measured under standard assay conditions at the indicated pH values. To test the stability, the enzyme was exposed for 30 min at 30°C to the pH values given, with subsequent determination of the residual activity at pH 6.5 (-) pH optimum; (---) stability. Buffers: (A) Sodium acetate; (0) potassium phosphate, (0) Tris-HCl.
BIOSYNTHESIS
61
OF PENTAGALLGYLGLUCOSE
30
A
Ho oP'o
20
P: /
v ,/
0.05
8
0
a
.rp'*
10 LOO
0.5 l/S
0l 0
-0
5
Substrate
concentration,
1
2
S (mM)
FIG. 3. Substrate saturation curves for /3-glucogallin (A) and 1,2,3,6-tetragalloylglucose (B).
Chrosorb RP-18,5 pm, column 180 X 3 mm i.d.; solvent A methanol; solvent B 0.05% HaPO,; gradient O-7 min = 7% A,7-20 min = ‘7-50% A,20-25 min = 50-90% A; flow rate 0.8 ml/min). Quantification was done by uv photometry at 280 nm (aromatic compounds) or by radioactivity measurements after fractionation of the HPLC eluates (glucose, &glucogallin). The resulting data were corrected against blanks with heat-denatured enzyme. Analytical methods. Determination of enzymatitally formed pentagalloylglucose, as well as estimations of the progress in the purification of the substrate tetragalloylglucose, were carried out by analytical HPLC on Merck LiChrosorb RP-18 columns (180 X 3 mm i.d., particle size 5 pm). The solvent systems used were modifications of that given in (12), consisting of acetonitrile (A) and 0.05% H3P04 (B). In routine enzyme activity assays, an efficient separation of tetragalloylglucose (retention time Rt = 10.2 min) and pentagalloylglucose (Rt = 11.70 min) was achieved by a linear gradient of lo-80% A within 22.5 min. The purification of tetragalloylglucose from crude tannin was monitored by a linear gradient of O40% A within 20 min, resulting in a Rt value of 13.5 min. A flow rate of 1.0 ml/min was maintained in all these experiments. Quantitative data were obtained by uv photometry at 280 nm, in combination with a computing integrator (Merck-Hitachi D-2000) referring to external standard solutions of the investigated compounds. Galloylglucoses to be analyzed by ‘H NMR spectroscopy were dried in DUCUO (5 mm Hg) at 85°C over PzOsand KOH pellets for 2 days and dissolved in DMSO-&. Spectra were recorded with a Bruker AM 200 at 200 MHz, using tetramethylsilane as internal standard. Assignment of glucose proton sequences was achieved by spin-decoupling experiments.
RESULTS AND DISCUSSION
Enzyme pur$cat&m. 4-O-Galloyltransferase was extracted from leaves of young oak plants grown in the greenhouse which were still devoid of the viscous mucilage characteristic of older leaves that had severely hampered some previous investigations (3,5). To ensure a continuous supply of suitable plant material, the stems of older plantlets were cut to enforce the subsequent development of new shoots bearing fresh leaves of the required age. (Unavoidable gaps could be bridged by using leaves stored at -20°C.) The crude extracts obtained from this plant material were partially purified by ammonium sulfate precipitation and anion-exchange chromatography (cf. Material and Methods). The resulting enzyme preparations were sufficiently pure, except in the stoichiometry experiments where the effects of a contaminating esterase had to be considered. General properties of the galloyltransferase. The synthesis of pentagalloylglucose was strictly dependent on the presence of both /3-glucogallin and 1,2,3,6-tetragalloylglucose, and no reaction occurred in samples with heat-denatured enzyme (Fig. 1). When isomeric 1,2,4,6-tetragalloylglucose was assayed as a potential substrate, only negligible activity (1.1%) was found. Under standard assay conditions, a roughly linear progress of the enzymatic reaction
62
CAMMANN
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TABLE I STOICHIOMETRYOFTHE~-O-GALLO~RANSFERASE-CATALYZEDFORMATIONOFPENTAGALLOYLGLUCOSE"
Component assayed
Gross reaction rate (nmol)
Corrected reaction rate* (nmol)
Molar ratio”
P-Glucogallin, consumed 1,2,3,6-Tetragalloylglucose, consumed 12 t I3I476-Pentagalloylglucose, formed Glucose, formed Gallic acid, formed
28.4 8.7 8.9 28.4 20.0
8.4 8.7 8.9 8.4 0
1.0 1.0 1.0 1.0 -
a Experimental details are described under Materials and Methods. bIt is assumed that gallic acid is produced by hydrolysis of the substrate /3-glucogallin, liberating concomitantly an equal amount of glucose. ‘Values refer to the reaction rate determined for tetragalloylglucose.
was observed with respect to time for ca. ‘70 min, gradually declining thereafter with a maximum at about 100 min. In contrast, a nonlinear relationship to the amount of added protein occurred; it is assumed that this anomaly is caused by the pronounced tanning (i.e., protein-binding and precipitating) capacities of tetra- and pentagalloylglucose in the assay mixtures (14,X). The effect of pH on the stability of the enzyme and on the velocity of the reaction is depicted in Fig. 2. The optimal pH was 6.3; maximal stability was found between pH 5.0 and 6.5. The temperature optimum of the reaction was at ea. 40°C. The rather unusual observation that the enzyme exhibited some activity even at 0°C parallels previous observations with related enzymes from oak (8) and sumac (9). Between 25 and 35%, a Q10value of 2.0 and an activation energy of 28.8 kJ/mol were calculated.
OH
The enzyme showed normal MichelisMenten kinetics with both substrates (Fig. 3). Replots of these data according to Lineweaver-Burk gave Km values of 2.3 and 1.0 IIIM for B-glucogallin and tetragalloylglucase, respectively. From gel-filtration experiments with a calibrated Sephadex G-200 column (16), an apparent molecular weight of 260,000 was estimated for the enzyme. This comparatively high value is consistent with those determined for other galloyltransferases from oak (6,8), but in clear contrast to the A& 75,000 for a related trigalloylglucosesynthesizing enzyme (9) from sumac leaves (unpublished data). Reaction-product identification. The enzyme reaction product was tentatively identified as 1,2,3,4,6-penta-Q-galloyl-@Dglucopyranose by cochromatography with authentic references (cf. Fig. 1). This conclusion was unequivocally confirmed by ‘H
+DG
FIG. 4. Reaction equation for the formation of pentagalloylglucose by galloylation of tetragalloylglucose. j3G, &Glucogallin, Glc, glucose.
BIOSYNTHESIS
63
OF PENTAGALLOYLGLUCOSE
NMR spectroscopy of material that had been isolated and purified from scaled-up assay mixtures, including D20 exchange, spin-decoupling experiments, and comparison with data from pure compounds (for details, see Materials and Methods). The recorded spectrum showed five singlets (6 6.78-6.89) corresponding to the five galloyl residues of the product, and the signals for all of the glucose protons were shifted downfield in response to acylation. P-Configuration of the product was proven by a coupling constant 3JH-1,2 of 8 Hz; other coupling constants for H-l to H-6,6’ (J ca. 10 Hz) showed that the glucose moiety had the expected normal 4C1-chair eonfiguration. Stoichicvmetrg Most likely, 1 mol of /3glucogallin serves as galloyl donor for the acylation of 1 mol of tetragalloylglucose under the concomitant production of 1 mol each of pentagalloylglucose and free glucose. Experimental proof was complicated, however, by the contamination of the enzyme preparation with an esterase activity that effectively hydrolyzed the substrate P-glucogallin. Assuming that gallic acid found in the assay mixtures was exclusively formed by this process, the amount of this by-product could serve to correct the determined concentrations of 8-glucogallin and glucose. Under this premise, and as summarized in Table I, it is evident that the observed turnover of substrates and products was fully consistent with the expected reaction depicted in Fig. 4. According to this equation, the systematic name “P-glucogallin: 1,2,3,6-tetra-O-galloyl$-Dglucose 4Ggalloyltransferase” (EC 2.3.1.-) is proposed for this new enzyme. In conclusion, earlier postulates (5,8) on the role of P-glucogallin as the principal acyl donor in the pathway from gallic acid to pentagalloylglucose have been confirmed by this investigation. Moreover, the precise structure of all the intermediates along this sequence is now known, a fact that must be attributed mainly to recent enzyme studies [cf. (17)]. Only one step, i.e., the supposed acylation of 1,2,6-trigalloyl-
glucose to 1,2,3,6-tetragalloylglucose, has not yet been fully characterized by this means; however, analyses of the galloylglucoses formed by oak tissues in tivo (18) or in vitro (8) are fully consistent with this proposal. ACKNOWLEDGMENTS We thank Miss U. Semler for expert technical assistance, and Professors G. Nonaka (Fukuoka) and E. Haslam (Sheffield) for providing reference substances. REFERENCES 1. HASLAM, E. (1982) Fartschr. Chem. Org. N&u&. 41,1-46. 2. GROSS,G. G. (1982) FEBSLett. 14?3,67-70. 22,2179-2182. 3. GROSS,G. G. (1983) Phytochemistry 4. WEISEMANN, S., DENZEL, K., SCHELING, G., AND GROSS,G. G. (1988) Bioorg. Chem l&29-37. 5. GROSS,G. G. (1983) Z. Nuturfwsck C38,519-523. 6. GROSS, G. G., SCHMIDT, S. W., AND DENZEL, K. (1986) J. Plant Physid 126,1’73-179. 7. DENZEL, K., WEISEMANN, S., AND GROSS, G. G. (1988) J. Plant Physiol. 133,113-115. 8. SCHMIDT, S. W., DENZEL, K., SCHILLING,G., AND GROSS, G. G. (1987) Z. Nuturforsch
C42,87-92.
9. DENZEL, K., SCHILLING, G., AND GROSS, G. G. (1988) PZuntu 176,135-137.
10. NISHIZAWA, M., YAMAGISHI, T., NONAKA, G., AND NISHIOKA, I. (1980) Chem Phcwm. BulL 28, 2850-2852. 11. NISHIZAWA, M.,YAMAGISHI, T., NONAKA, G., AND NISHIOKA, I. (1983) J. Chem. Sot. Perkin Trans.
1,961-965. 12. HADDOCK, E. A., GUPTA, R. K., AL-SHAFI, S. M. K., AND HASLAM, E. (1982) .J. Chem 5’~. Perkin
Trans. 1,2515-2524.
13. KUNITZ, M. (1952) J. Gen Physid 35,423-450. 14. HASLAM, E., AND LILLEY, T. H. (1985) in The Biochemistry of Plant Phenol& (Van Sumere, C. F., and Lea, P. J., Eds.), Annu. Proc. Phytothem. Sot. Eur.,Vol.25,pp. 237-256, Clarendon Press, Oxford. 15. OZAWA, T., LILLEY, T. H., AND HASLAM, E. (1987) Phytochxmistry
26,2937-2942.
16. ANDREWS, P. (1965) B&hem, .I 96,595-606. 17. GROSS, G. G. (1989) in Plant Cell Wall Polymers: Biogenesis and Biodegradation (Lewis, N. G., and Paice, M. G., Eds.), ACS Symp. Ser. Vol. 399, Washington, DC, in press. 18. KRAJCI, I., AND GROSS,G. G. (1987) Phytochemistry 26,141-143.