The role of chalcone synthase in the regulation of flavonoid biosynthesis in developing oat primary leaves

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS Vol. 250, No. 2, November 1, pp. 364-3’72,1986

The Role of Chalcone Synthase in the Regulation of Flavonoid Biosynthesis in Developing Oat Primary Leaves’ WOLFGANG AND

KNOGGE,**’ ELMON SCHMELZER,* GOTTFRIED WEISSENBiiCKt

*Max-Planck-Institut fiir Ziichtungsforschung, Abt. Bio~hemi~, D-5000 Cologne 30, and tBotanisches der Universittit, Gyrhofstrasse 15, D-5000 Cologne 41, Federal Republic of Germany Received December

Institut

18,1985, and in revised form May 27,1986

The role of chalcone synthase in the regulation of flavonoid biosynthesis during organogenesis of oat primary leaves has been investigated at the level of enzyme activity and mRNA translation in vitro. Chalcone synthase was purified about 500-fold. The apparent Km values were 1.5 and 6.3 pM for 4-coumaroyl-CoA and malonyl-CoA, respectively. The end products of oat flavonoid biosynthesis, three C-glucosylflavones, did not inhibit the reaction at concentrations as measured up to 60 pM each. Apigenin (4’,5,7trihydroxyflavone), a stable structural analog of the reaction product, 2’,4,4’,6’-tetrahydroxychalcone, was found to be a strong competitive inhibitor of 4-coumaroyl-CoA binding and a strong noncompetitive inhibitor of malonyl-CoA binding. Although apigenin is not supposed to be an intermediate of C-glucosylflavone biosynthesis, this compound might be a valuable tool for future kinetic studies. To date, there is no indication of chalcone synthase regulation by feedback or similar mechanisms which modulate enzyme activity. Mathematical correlation of chalcone synthase activity and flavonoid accumulation during leaf development, however, indicates that chalcone synthase is the rate-limiting enzyme of the pathway. By in vitro translation studies using preparations of total RNA from different leaf stages, we could demonstrate for the first time that the translational activity of chalcone synthase mRNA undergoes marked daily changes. The high values found at the end of the dark phase suggest that light does not exert direct influence on flavonoid biosynthesis but probably functions by controlling the basic diurnal rhythm. 0 1986 Academic Press, Inc.

During the past decade, substantial progress has been made elucidating the biosynthesis of plant secondary phenolics, one major class of which is the flavonoids. Most data on enzymology and regulation of flavonoid formation have been based on work with cultured plant cells (l-3). Parsley cell suspension cultures have been extensively studied. Dark-grown parsley cells respond to uv irradiation by accumulating flavonoids and by inducing the enzymes involved

in the biosynthesis of these compounds (2, 4). Furthermore, the changes in the activities of L-phenylalanine ammonia-lyase (PAL)3 and chalcone synthase (CHS), the key enzymes of plant general phenylpropanoid metabolism and of flavonoid biosynthesis, respectively, have been correlated with transient increases in the transcription rates of their corresponding genes (5, 6). Oat primary leaves provide a system to study flavonoid biosynthesis and its inte-

1 This work was supported in part by a grant from the Deutsche Forschungsgemeinschaft. ‘To whom correspondence should be addressed.

3 Abbreviations used: PAL, L-phenylalanine ammonia-lyase; CHS, chalcone synthase; SDS, sodium dodecyl sulfate.

0003-9861/86 $3.00 Copyright All righta

0 1986 by Academic Press. Inc. of reproduction in any form reserved.

364

ROLE

OF CHALCONE

SYNTHASE

gration into organ differentiation and development (7-9). The activities of the two initial enzymes (CHS and chalcone isomerase) and the final two of the pathway (a C-glucosylflavone 0-methyltransferase (10) and a C-glucosylflavone O-arabinosyltransferase) have been followed during organogenesis (7,9,11). CHS, a homodimeric protein (12), catalyzes the formation of the aromatic A ring of the tlavonoid skeleton by sequential condensation of three malonyl-CoA-derived acetate units with 4coumaroyl-CoA yielding 2’,4,4’,6’-tetrahydroxychalcone (Fig. 1). During early stages in the development of oat primary leaves CHS activity has been shown to increase concomitantly with the activities of the three other enzymes of tlavonoid biosynthesis so far investigated. Only for CHS, however, is this initial increase followed by a marked decrease in older leaves giving rise to an activity maximum at the plant stage where the most rapid tlavonoid accumulation was observed (9). In addition, preliminary results indicated an inhibitory effect of the metabolic end products, three C-glucosylflavones (13), on CHS activity in vitro (11). Here we report results which suggest that during leaf organogenesis, CHS is the rate-limiting enzyme of flavonoid formation. CHS biosynthesis may be regulated indirectly by light via diurnal changes of CHS gene transcription or CHS mRNA translational activity.

IN OAT

FLAVONOID

BIOSYNTHESIS

EXPERIMENTAL

PROCEDURES

Chemicals. [2-i4C]Malonyl-CoA (1.67 GBq/mmol) was purchased from New England Nuclear (Dreieich, FRG), L-[%S]methionine (>30 TBq/mmol), and rabbit reticulocyte lysate from Amersham-Buchler (Braunschweig, FRG). I-Coumaroyl-CoA and 2’,4,4’,6’-tetrahydroxychalcone were synthesized according to published methods (14, 15). Flavonoids were either obtained from Roth (Karlsruhe, FRG) or extracted from oat leaves and purified according to published methods (16,17). All phenolics used as substrates or inhibitors in enzyme assays had a degree of purity of >98% as determined by HPLC with detection at 260 nm (9). Buglers. The following buffers were used: (A) 0.1 M potassium phosphate, pH 8.0, containing 4.3 mM 2mercaptoethanol; (B) 0.1 M potassium phosphate, pH 8.0, containing 1.2 mM 2mereaptoethanol; (C) 0.2 M Tris-HCl, pH 8.7, containing 0.5 M NaCl, 10 mM EDTA, 10% (w/v) sucrose, 1% (w/v) iodoacetic acid, 0.5% (w/v) sodium dodecyl sulfate, 0.2% (v/v) 2-mercaptoethanol; (D) 10 mM Tris-HCl, pH 7.8, containing 1 mM EDTA. Plant n&e&l Oat seedlings (Arena sativa L.) were grown under standard conditions in a 13/11-h photoperiod (16). For enzyme assays, 20-40 leaves were harvested, for RNA extraction, 50-100 leaves, and for flavonoid extraction, 20 leaves. Enzyme extraction and assay. Extraction and assay of chalcone synthase were as previously described (9). The assay was carried out in buffer B containing 9.45 pM I-coumaroyl-CoA, 10 pM [2-i4C]malonyl-CoA, and up to 20 ~1 of enzyme solution in a total volume of 0.11 ml (18). Protein was determined by the method of Bradford using the Bio-Rad reagent (19). Since bovine serum albumin was used as standard. data for

OH HOy&

HO

0

II

365

OH

____ ----+ HO

0

III

FIG. 1. Structures of flavonoids mentioned in the text: 2’,4,4’,6’-tetrahydroxychalcone (I), naringenin (4’,5,7-trihydroxyflavanone, II), apigenin (4’,5,7-trihydroxyflavone, III). The structures of the C-glucosylflavones accumulating in oat primary leaves are published elsewhere (9,13).

366

KNOGGE.

SCHMELZER,

the purified protein were corrected according to the manufacturer’s instruction manual. Purification of chalcone synthase. Six-day-old oat primary leaves (300 g) were homogenized in liquid nitrogen. The resulting powder was extracted for 40 min with 1.2 liters of buffer A in the presence of 30 g of Polyclar AT and 30 g of Dowex AG 1X2 (phosphate form). After filtration through miracloth and centrifugation (30 min, 15,OOOg) the supernatant was treated with another 30 g of Dowex. Removal of the ion-exchange resin was achieved by filtration through miracloth. The purification on DEAE-cellulose and hydroxyapatite was essentially as described by Kreuzaler and Hahlbrock (20). The fractions containing the highest CHS activity were pooled and concentrated by ultrafiltration to a final volume of 2.2 ml. After bovine serum albumin was added to yield a protein concentration of 3 mg/ml, 50-~1 aliquots were stored at -20°C. Enzyme kinetics. Apparent Michaelis constants were calculated from Hanes-Woolf plots by linear regression. For product inhibition studies, inhibitor concentrations up to 60 pM were used in the standard assay. A more detailed inhibition analysis was performed with different apigenin concentrations, varying the concentration of one substrate at a constant concentration (10 pM) of the other substrate. Inhibition constants were calculated by linear regression from secondary plots of slopes (K,) and intercepts (&), respectively (21, 22). Extraction of jZawonoi& Flavonoid products were extracted from the leaves with 80% (v/v) methanol. Quantitative determination was performed after HPLC-assisted separation of the compounds as previously described (9). Extraction of total RNA. For the extraction of total RNA from oat primary leaves a published method (23,24) was modified. Leaves (3 g) were homogenized in liquid nitrogen. The resulting powder was allowed to thaw for 2 min in 4 ml of buffer C under continuous grinding. Tissue debris was removed by centrifugation (2 min, 12,000g). The supernatant was successively extracted twice with 1 vol each of phenol (saturated with buffer D, 0.1% (w/v) S-hydroxyquinoline)/chloroform (4% (v/v) isoamyl alcohol) l/l and with 1 vol of chloroform (4% (v/v) isoamyl alcohol). The RNA was precipitated with 2.5 vol of ethanol in the presence of 0.3 M Na acetate (-2O”C, overnight). After centrifugation (20 min, 12,OOOg)the pellet was washed with 3 M Na acetate, pH 5.2. After a second alcohol precipitation, the pellet was dried under reduced pressure and dissolved in a minimum amount of sterile water. RNA concentration was determined spectrophotometrically. Cell-free protein synthesis and immunopreeipitation Portions (50 pg) of total RNA were translated in vitro in the presence of [35S]methionine using a cell-free lysate from rabbit reticulocytes (24,25). Translational

AND

WEISSENBGCK

efficiency was determined by measuring the trichloroacetic acid-insoluble radioactivity in 47~1 aliquots. Equal amounts of total translation products (100,000 cpm) were submitted to immunoprecipitation (26), using an antiserum raised against parsley CHS (27). The precipitates were dissolved in 30 pl of sample buffer (28). Those volumes of the solutions which reflected the RNA from half a leaf were subjected to sodium dodecyl sulfate (SDS)-polyacrylamide (lo15%)gel electrophoresis. Synthesized CHS was quantitated after autoradiography (29) by scanning the films with a laser densitometer. RESULTS

Enzyme PurQication

The procedure developed for the purification of CHS from parsley cell cultures (20) was also applicable to the purification of CHS from oat primary leaves. On native polyacrylamide gels the resulting enzyme preparation yielded only one band which showed CHS activity as measured using homogenized gel slices in the standard assay (result not shown). An additional faint band demonstrated the presence of a minor impurity. The specific activity was 153 pkat/kg (Table I). The enzyme preparation was free of chalcone isomerase activity. Under appropriate assay conditions (30), therefore, the assay yielded more than 80% 2’,4,4’,6’-tetrahydroxychalcone. In addition, minor amounts of 4’,5,7-trihydroxyflavaa compound being none (naringenin), formed in the assay by spontaneous cyclization of the chalcone at a pH >4 (Fig. l), and a compound of unknown chemical nature (cf. (30)) were found. Maximum activity was obtained in 0.1 M potassium phosphate, pH 8.0 (1.2 mM 2-mercaptoethanol). Enzyme Kinetics

Apparent Michaelis constants were 1.5 for 4-coumaroyl-CoA in the presence of 29.2 PM malonyl-CoA and 6.3 PM for malonyl-CoA with 12.6 PM 4-coumaroyl-CoA. In addition, the influence of reaction products and metabolic end products on CHS reaction rate was analyzed. As previously reported for the CHS isolated from parsley cell cultures (20) and tulip anthers (31), CHS from oat was inhibited in vitro by coenzyme A, a product of the reaction, as PM

ROLE

OF CHALCONE

SYNTHASE

IN OAT TABLE

Crude extract after second Dowex treatment 50-80% (NH,), SO1 fractionation, Sephadex G-25 DEAE-cellulose Hydroxyapatite, ultrafiltration

Protein (mg)

CHS activity (pkat)

367

BIOSYNTHESIS

I

PARTIAL PURIFICATION Purification step

FLAVONOID

OF CHS

Specific activity (rkatkz)

Purification (fold)

Recovery (%I

1290

381

0.30

1.0

100

735

295

185

100

0.40 0.54

1.3 1.8

77 26

0.62

95

well as by naringenin. The three end products of oat flavonoid biosynthesis, three Cglucosylflavones (13), did not affect the reaction rate of the purified enzyme at concentrations as measured up to 60 PM each. This is in contrast to preliminary experiments using a crude enzyme extract (11). The natural product of the CHS reaction, 2’,4,4’,6’-tetrahydroxychalcone, cannot be used in product inhibition studies because of its high instability under assay conditions. Therefore, it was replaced by 4’,5,7trihydroxyflavone (apigenin), a stable structural analog of the chalcone (Fig. 1). Fifty percent inhibition was achieved with 35 PM coenzyme A, 85 pM naringenin, and 4.7 PM apigenin, respectively. In a more detailed analysis using apigenin, competitive inhibition was obtained by varying the concentration of 4-coumaroyl-CoA (ki, = 0.4 PM) and noncompetitive inhibition by

02 l/[Lc-coA,

153

511

25

varying the concentration of malonyl-CoA (Kii = 6.8 PM, Ki, = 6.3 FM). The inhibition constants were calculated from linear secondary plots as shown in Fig. 2. Product Accumulation

and CHS Activity

The results on flavonoid biosynthesis in developing oat primary leaves (7,9) clearly demonstrated that the CHS activity profile differs from all other enzymes of this pathway so far investigated. Chalcone isomerase, the flavonoid O-methyltransferase (lo), and the tlavonoid O-arabinosyltransferase all follow a hyperbolic time course during leaf development. In contrast, CHS activity reaches a maximum in the plant stage which shows the most rapid flavonoid accumulation (9, 11). When the activity of the rate-limiting enzyme of a pathway is not regulated, e.g.,

05 WI-’

02 ‘Hal

- CoAl

05 b”‘-’

FIG. 2. Inhibition of CHS reaction by apigenin (a) with 4-coumaroyl-CoA concentrations varied at 10 PM malonyl-CoA, (b) with malonyl-CoA concentrations varied at 10 pM I-coumaroyl-CoA. Inhibitor concentrations were 0 (0), 2 (O), 4 (A), and 6 pM (A). The insets show the secondary plots used to calculate the inhibition constants. 4C-CoA, I-coumaroyl-CoA; I, apigenin.

368

KNOGGE,

SCHMELZER,

by feedback inhibition, and degradation of products is negligible, product accumulation can be described by the integration of the enzyme activity curve using the equation P(t) = $ E(T)& (l-3). Under the growth conditions applied, the flavonoids are not metabolized during the development of oat primary leaves (32). Furthermore, there is no indication for a regulation of CHS by the metabolic end products (see above). Therefore, an attempt was made to mathematically correlate flavonoid accumulation and CHS activity (Fig. 3). All experimentally derived data on flavonoid accumulation lay on the theoretical curve. This result suggests that CHS is the ratelimiting enzyme of flavonoid biosynthesis in developing oat primary leaves. PAL is the only enzyme showing an activity profile during leaf growth which is similar to that of CHS (7,9). An analogous mathematical correlation of PAL activity

5

6 Plant

7 Age

8

9

10

[dl

FIG. 3. Mathematical correlation of relative CHS activity and flavonoid accumulation. Enzyme activity (0) was measured during leaf development. The obtained curve for relative CHS activity (-) was integrated to yield the product accumulation curve (---). All experimentally derived data for flavonoid accumulation (0) lay on the calculated curve. The absolute CHS activity as measured in vitro accounted for 65% of the actual amount of flavonoids accumulating in oat primary leaves. Accordingly, the calculated curve had to be corrected.

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WEISSENBGCK

and llavonoid accumulation, however, has two major difficulties. First, PAL is an enzyme involved in general phenylpropanoid metabolism, the products of which are precursors of different classes of phenolic compounds in oat leaves, such as lignin and hydroxycinnamic acid esters (9). Second, PAL is found in all tissues in contrast to CHS, which is located in the leaf mesophyll (9). Since flavonoid biosynthesis is assumed to take place in the leaf mesophyll, only the PAL activity profile in this tissue was integrated. There was, however, no coincidence between the calculated curve and the measured flavonoid accumulation (data not shown).

Isolation of RNA and Translation in Vitro In uv-irradiated parsley cell suspension cultures, the timing of flavonoid accumulation was found to be determined by the kinetics of CHS gene transcription (6). As a first step in the investigation of CHS biosynthesis in oat primary leaves, it was attempted to quantitate the CHS mRNA during leaf development. Total RNA extracted during organogenesis yielded 30-70 pg/leaf (1 mg/g fresh weight of 4-day-old leaves, about 0.4 mg/g fresh weight of old leaves). In young leaves (4-6 days) the amount of RNA was 50-65s higher at the end of the light phase as compared to that at the end of the dark phase. In older plants this difference became less pronounced. Since a parsley-derived CHS cDNA clone (5) did not hybridize to oat CHS mRNA in Northern and dot blot experiments, the amount of CHS mRNA extracted from the leaves could not be demonstrated in a direct approach. An antiserum raised against parsley CHS (27), however, inhibited CHS activity in crude enzyme extracts from oat an antiserum raised leaves, whereas against parsley PAL did not, demonstrating the specificity of the inhibition (data not shown). Relative CHS mRNA amounts were, therefore, quantitated indirectly after translation in vitro and immunoprecipitation. Equal amounts of total RNA were subjected to cell-free translation. Total

ROLE

OF CHALCONE

1

2

SYNTHASE

345

IN OAT

6

7

8

FLAVONOID

9

10

11

BIOSYNTHESIS

12

369

13

FIG. 4. Autoradiogram of an SDS-polyacrylamide gel after separation of in vitro translation products as immunoprecipitated using anti-(parsley CHS) serum. Lanes: 1 and 13, protein standards (200,000, 92,500, 69,000, 46,000, 30,000); 2, immunoprecipitate obtained when total RNA from uvinduced parsley cell suspension cultures was used as an external control for the efficiency of in vitro translation and immunoprecipitation; 3-11, immunoprecipitates after translation in vitro using total RNA from 4-, 4.5-, 5-, 5.5-, 6-, 6.5-, ‘i’-, 8-, and lo-day-old oat primary leaves; 12, control immunoprecipitate of an assay performed in the absence of RNA. The bands in lanes 7-9 migrating slower than CHS show nonspecifically immunoprecipitated protein. Since these bands sometimes were seen in controls like lane 12 the precipitated protein must be of lysate origin.

RNA from uv-induced parsley cells was used as an external control for the efficiency of in vitro translation and immunoprecipitation. The RNA isolated from younger oat leaves translated better as compared to older leaves. Equal amounts of total translation products were, therefore, used in the immunoprecipitation step. Finally, the amount of precipitate reflecting the RNA from half a leaf was applied to SDS-polyacrylamide gel electrophoresis. Figure 4 shows an autoradiogram of a gel after separation of the in vitro translation products precipitated with anti-CHS serum. Figure 5 shows the time course of relative translational activity of CHS mRNA as determined by densitometric scanning of the autoradiograms (cf. Fig. 4). High activity was already present at the end of the dark phase and subsequently decreased during illumination.

DISCUSSION

CHS catalyzes the first step in the flavonoid biosynthesis of plants. This key metabolic position makes the enzyme an attractive target for regulatory mechanisms controlling the pathway. Work on the purified enzyme from parsley and carrot cell suspension cultures (20,33), as well as on the partially purified enzyme from tulip anthers (31), carrot cell cultures (34), and now oat primary leaves, however, does not yield any information for possible regulation of CHS at the level of enzyme activity. It can be expected that the products of the reaction inhibit the enzyme. This holds true for coenzyme A (cf. (21, 31)). Naringenin, usually applied as a racemate, exerts an inhibitory effect on the reaction possibly because of its partial structural similarity to the chalcone product. Re-

370

KNOGGE,

SCHMELZER,

FIG. 5. Relative CHS mRNA translational activity during leaf development. The points represent the mean of at least two translations in vitro using total RNA from two independent experimental series. Shaded areas indicate the 11-h dark phases of the photoperiod.

cently, with partially purified CHS from cultured carrot cells, an uncompetitive inhibition pattern has been described for naringenin at varying concentrations of 4coumaroyl-CoA (34). It is not known, however, whether the naturally occurring (-)(2s) enantiomer of narigenin, which is the product of the subsequent enzyme in flavonoid biosynthesis, chalcone isomerase (35, 36), can act as an inhibitor in vivo. 2’,4,4’,6’-Tetrahydroxychalcone, the product of the CHS reaction, cannot be used in kinetic studies because it is cyclized spontaneously to naringenin under the optimal assay conditions. We, therefore, chose apigenin as a stable structural analog which differs from the chalcone only by the closed heterocycle (Fig. 1). The observed inhibition patterns (Fig. 2) clearly demonstrate that apigenin competes with 4-coumaroylCoA for the same binding site. Apigenin is a metabolic intermediate in the biosynthesis of 0-glycosylflavones (37, 38), but probably not of C-glucosylflavones (39-41) which accumulate in oat leaves. The inhibition, therefore, is not interpreted as being relevant in vivo. Apigenin, however, may be a valuable tool in future studies on CHS kinetics or possibly as a ligand in affinity chromatography. Similar to the CHS from carrot cells (34), the noninhibition of oat CHS by the oat llavones in vitro apparently excludes end product inhibition in vivo.

AND

WEISSENBiiCK

From studies with a Petunia hybrida mutant altered in CHS gene expression (42), as well as from studies with white and red flowers of the wild carrot (43), it is known that petals containing little or no anthocyanins also have little or no CHS activity. The activities of other enzymes involved in the pathway, however, were less reduced as compared to colored petals. This has been interpreted as an indication for a possible role of CHS as the rate-limiting enzyme. For developing oat primary leaves, mathematical correlation of CHS activity and flavonoid accumulation resulted in the coincidence of experimentally obtained and calculated curves for product formation (Fig. 3). This very close coincidence tends to exclude regulatory mechanisms modulating CHS activity in vivo. Furthermore, this relationship can only be expected for the rate-limiting enzyme of a biosynthetic pathway. Under the plant growth conditions applied, therefore, the amount of flavonoids synthesized appears to be determined exclusively by the CHS activity present in the leaves. For dark-grown parsley cell suspension cultures it could be demonstrated that the biosynthesis of llavonoids was induced by uv irradiation. The timing of product formation was determined by the kinetics of CHS gene transcription (6). In developing oat primary leaves regulation of flavonoid biosynthesis appears to be more complex. Oat leaves are known to accumulate flavonoids when grown in continuous darkness and to further increase the amount of flavonoids upon illumination (7). When grown in a light-dark rhythm, however, CHS activity was found to increase and subsequently decrease showing a maximum in 5.6-day-old leaves (10) (Fig. 3); i.e., enzyme activity did not appear to respond to illumination at the beginning of the light phase of the photoperiod. Surprisingly, CHS mRNA translational activity showed drastic daily changes with high values at the end of the dark phase, and a subsequent decrease during the light phase. This pattern was repeated during the 3-day period in which significant translation of CHS

ROLE

OF

CHALCONE

SYNTHASE

mRNA was measured in vitro (Fig. 5). Preliminary results indicate that a rhythm can be measured for CHS activity, too, when leaves are analyzed in shorter time intervals. The amplitude of the curve is lower, however, and the peak positions are delayed as compared with translational activity (G. WeissenbBck, unpublished). From the results presented, we conclude that light did not exert a direct influence on flavonoid biosynthesis in oat primary leaves under the growth conditions applied. This becomes particularly obvious because of the marked increases of CHS mRNA translational activity during the dark phases of the photoperiod. Light acts, however, indirectly in controlling the basic daily rhythm and, thus, the developmental program of the leaf as a whole. Recently, it was shown that kinetin stimulated flavonoid accumulation in buckwheat cotyledons in the dark (44), whereas gibberellic acid inhibited anthocyanin biosynthesis at the CHS level in carrot cell suspension cultures (45). The control of plant hormone concentrations, therefore, may represent a possible mechanism of light action in the regulation of flavonoid biosynthesis. In this connection it may be of interest how CHS activity and CHS mRNA translational activity in developing oat primary leaves respond to variations in the photoperiod. Their time courses in plants grown in continuous darkness are currently being investigated. The answer to the question whether the diurnal rhythm is controlled post-transcriptionally, e.g., by changes in the specific translational activity of CHS mRNA, or, alternatively, by corresponding changes in CHS gene activity, has to await the development of an oat CHS cDNA clone. ACKNOWLEDGMENTS We thank Monika Walther for technical assistance, Prof. Dr. K. Hahlbrock for providing anti-(parsley CHS) serum, and Maya Spies for typing the manuscript. REFERENCES 1. HAHLBROCK, K., AND GRISEBACH, H. (1979) Rev. Plant PhysioL 30,105-130.

Annu.

IN

OAT

FLAVONOID

371

BIOSYNTHESIS

2. HAHLBROCK, K., SCHRC~DER, J., AND VIEREGGE, J. (1980) in Plant Cell Cultures II (Viechter, A., ed.), pp. 39-60, Springer, Berlin. 3. HAHLBROCK, K. (1981) in The Biochemistry of Plants (Stumpf, P. K., and Conn, E. E., eds.), Vol. 7, pp. 425-456, Academic Press, New York. 4. HELLER, W., EGIN-BUHLER, B., GARDINER, S. E., KNOBLOCH, K.-H., MATERN, U., EBEL, J., AND HAHLBROCK, K. (1979) Plant Physiol. 64, 371373. 5. KREUZALER, F., RAGG, H., FAUTZ, E., KUHN, D. N., AND HAHLBROCK, K. (1983) Proc. NatL Acud Sci

USA 80,2591-2593. 6. CHAPPELL, J., AND HAHLBROCK, K. (1984) Nature (London) 311,76-78. WEISSENB~CK, G. (1975) 2. PJanzenphysiol. 74, 226-254. WEISSENB~CK, G., AND SACHS, G. (1977) Planta 137, 49-52. KNOGGE, W., AND WEISSENB~CK, G. (1986) Planta 167,196-205. 10. KNOGGE, W., AND WEISSENB~CK, G. (1984) Eur. J. B&hem. 140,113-118. 11. FUISTING, K., AND WEISSENB~CK, G. (1980) Z. Na-

turfmsch.

35c, 973-977.

12. KREUZALER, F., RAGG, H., HELLER, W., TESCH, R., WITT, I., HAMMER, D., AND HAHLBROCK, K. (1979) Eur. J. B&hem. 99,89-96. 13. CHOPIN, J., DELLAMONICA, G., BOUILLANT, M. L., BESSET, A., POPOVICI, G., AND WEISSENB~CK, G. 16,2041-2043. (1977) Phytochemistry 14. STOCKIGT, J., AND ZENK, M. H. (1975) 2. Naturforsch. 3Oc, 352-358. 15. MOUSTAFA, E., AND WONG, E. (1967) Phytochem-

istry 6, 625-632. 16. WEISSENB~CK,

G.,

PjZanzenphysioL

AND

EFFERTZ,

B.

(1974)

Z.

74.298-326.

17. POPOVICI, G., WEISSENB~CK, G., BOUILLANT, M. L., DELLAMONICA, G., AND CHOPIN, J. (1977) Z. PjZanzenphysioL 85, 103-115. 18. SCHRBDER, J., HELLER, W., AND HAHLBROCK, K. (1979) Plant Sci. IAt. 14,281-286. 19. BRADFORD, M. M. (1976) Anal Biochem. 72, 248254. F., AND HAHLBROCK, K. (1975) Eur. 20. KREUZALER,

J. Biochem. 56,205-213. 21. CLELAND, W. W. (1963) B&him. Biophya Acta 67, 173-196. 22. CLELAND, W. W. (1970) in The Enzymes (Boyer, P. D., ed.), 3rd ed., Vol. 2, pp. l-65, Academic Press, New York. 23. LANGRIDGE, P., PINTO-T• RO, J. A., AND FEIX, G. (1982) Mol. Gen Genet. 187,432-438. 24. KUHN, D. N., CHAPPELL, J., BOUDET, A., AND HAHLBROCK, K. (1984) Proc. Natl. Acad. Sci USA 81,1102-1106. 25. PELHAM, H. R. B., AND JACKSON, R. J. (1976) Eur.

J. B&hem.

67,247-256.

372

KNOGGE,

SCHMELZER,

26. MACCECCHINI, M. -M., RUDIN, Y., BLOBEL, G., AND SCHATZ, G. (1979) Proc Natl. Acad Sci USA 76,343-347. 27. SCHR~DER, J., KREUZALER, F., SCHHFER, E., AND HAHLBROCK, K. (1979) J. Biol. Chem 254, 5765. 28. KING, J., AND LAEMMLI, U. K. (1971) J. h4oL Biol. 64,465-477. 29. BONNER, W. M., AND LASKEY, R. A. (1974) Eur. J. B&hem. 46,83-88. 30. HELLER, W., AND HAHLBROCK, K. (1980) Arch. Biochem Biophys. 200,617-619. 31. S~I-FELD, R., KEHFCEL,B., AND WIERMANN, R. (1978) Z. Naturforsch 33c, 841-846. 32. PROKSCH, M., STRACK, D., AND WEISSENB~CK, G. (1981) 2. Naturforch. 36c, 222-233. 33. OZEKI, Y., SAKANO, K., KOMAMINE, A., TANAKA, Y., NOGUCHI, H., SANKAWA, U., AND SUZUKI, T. (1985) J. Biochxm. (Tokyo) 98,9-17. 34. HINDERER, W., AND SEITZ, H. U. (1985) Arch. B&hem. Biophys. 240,265-272. 35. WONG, E., AND MOUSTAFA, E. (1966) Tetrahedron L&t. 26,3021-3022.

AND

WEISSENBijCK

36. BOLAND, M. J., AND WONG, E. (1975) Eur. J. Biocbm. 50,383-389. 37. SIJITER, A., POIJLTON,J., AND GRISEBACH, H. (1975) Arch. Biochem. Biophys. 170,547-556. 38. STOTZ, G., AND FORKMANN, G. (1981) 2. Naturforsch. 36c, 737-741. 39. WALLACE, J. W., MARBY, T. J., AND ALSTON, R. E. (1969) Phytochemistvy 8,93-99. 40. WALLACE, J. W., AND MARBY, T. J. (1970) Phytochemistry 9,2133-2135. 41. WALLACE, J. W., AND GRISEBACH, H. B&him Biophys. Acta 304,837~841.

(1973)

42. MOL, J. N. M., SCHRAM, A. W., DE VLAMING, P., GERATS, A. G. M., KREUZALER, F., HAHLBROCK, K., REIF, H. J., AND VELTKAMP, E. (1983) Mol. Gen Genet. 192,424-429. 43. HINDERER, W., NOB, W., AND SEITZ, H. U. (1983) Phytochemistry 22.2417-2420. 44. MARGNA, U., AND VAINJXRV, T. (1983) Z. Naturfbrsch 38c, 711-718. 45. HINDERER, W., PETERSEN, M., AND SEITZ, H. U. (1984) Planta 160,544-549.