JOURNAL OF BACTERIOLOGY, Feb. 1978, p. 942-9510021-9193/78/0133-0942$02.00/0Copyright © 1978 American Society for Microbiology

Vol. 133, No. 2

Printed in U.S.A.

Induction of a Biopolyester Hydrolase (Cutinase) by LowLevels of Cutin Monomers in Fusarium solani f. sp. pisit

T. S. LIN AND P. E. KOLATTUKUDY*

Department ofAgricultural Chemistry and Program in Biochemistry and Biophysics, Washington StateUniversity, Pullman, Washington 99164

Received for publication 26 August 1977

Cutin hydrolysate induced the production of an extracellular cutinase byglucose-grown Fusarium solani f. sp. pisi. The rate of production depended onthe amount of cutin hydrolysate added up to 80 ,ug/ml, and saturation wasattained at this level. Glucose was found to be a repressor of cutinase production.A radial immunodiffusion assay for cutinase was developed, and the inductionof cutinase by cutin hydrolysate was confirmed by this direct assay. Whencutinase was induced by cutin hydrolysate, exogenous labeled phenylalanine wasincorporated into cutinase, which was shown to be the major (>70%) protein inthe extracellular fluid. Induction of cutinase by cutin hydrolysate was not in-hibited by actinomycin D and was stimulated (=100%) by cordycepin. Additionof cycloheximide with the inducer, or up to 12 h after the addition of the inducer,resulted in a nearly immediate cessation of cutinase production. Deoxyglucose,an inhibitor of proten glycosylation, inhibited the induction of cutinase by cutinhydrolysate. w-Hydroxy fatty acids were more effective in inducing cutinase thanany of the other more polar acids of cutin. Experiments with derivatives andanalogues of o-hydroxy C16 acid indicated that a free hydroxyl group at the w-

position was the most important factor determining the cutinase-inducing activity.n-Aliphatic primary alcohols with 14 or more carbon atoms induced cutinase, andn-C16 was the most effective inducer. These results strongly suggest that themonomers function as the chemical signal which induces the extracellular hydro-lase.

Many microorganisms grow on polymeric ma-terials as their sole source of carbon. The utili-zation of polymers involves excretion of hydro-lytic enzymes by the microorganisms. It is notknown how a polymer, which does not enter thecell, triggers the induction of the hydrolytic en-zymes. One possibility is that microbes excretea basal level of several hydrolytic enzymes, andthe hydrolysis products generated from the pol-ymer enter the cell and induce the synthesis ofthe appropriate hydrolase. In support of such ahypothesis is the observation that cellulase isinduced by cellobiose and derivatives (19). Dur-ing recent years it was found that several phy-topathogenic fungi grow on cutin, the biopolyes-ter made of hydroxy and epoxy fatty acids (7),as the sole source of carbon (4, 5, 16, 22). Undersuch growth conditions, Fusarium solani f. sp.pisi excretes an induced hydrolytic enzyme, cu-tinase (16), which has been purified and char-acterized (17, 18). How the insoluble polymer,cutin, brings about this enzyme induction is un-known. If the hypothesis indicated above holds

t Scientific paper no. 4894, project 2001, of the College ofAgriculture Research Center, Pullman, Wash.

good for the present system, low levels of cutinhydrolysate might induce the synthesis of cuti-nase. In this paper we report that low levels ofthe polymer hydrolysate do, indeed, result inthe induction of cutinase. Such induction, how-ever, occurs only after the depletion of glucosein the medium, thus showing a dual control ofcutinase production. Under such conditions, cu-tinase constitutes the major protein in the extra-cellular fluid.

MATERIALS AND METHODSActinomycin D, cycloheximide, cordycepin, 2-

deoxy-D-glucose, p-nitrophenyl butyrate (PNB), o-dianisidine, glucose oxidase (from Aspergillus niger),and horseradish peroxidase were purchased fromSigma Chemical Co. 16-Hydroxyhexadecanoic acidwas obtained from Aldrich Chemical Co. 10,16-Dihy-droxy C16 acid and 9,10,18-trihydroxy C,8 acid wereisolated from apple cutin and identified by combinedgas-liquid chromatography and mass spectrometry.All other fatty acids and fatty alcohols were purchasedfrom Analabs, Inc. L-Phenyl-[2,3-3H]alanine (16.6Ci/mmol) was obtained from Amersham/Searle.

F. solani f. sp. pisi was obtained from Lee Hadwigerof this University and was maintained on potato dex-

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INDUCTION OF CUTINASE BY CUTIN MONOMERS 943

trose agar at 23°C. Suspensions of spores in steriledistilled water were used for inoculation. The numberof spores in the suspension was determined by platingdiluted samples on 2% potato dextrose agar plates(pH 3.5), which were incubated at 230C for 3 days.Usually 107 to 109 spores were added to 100 ml of thegrowth medium contained in Roux Vulture bottles,which were incubated at 230C without shaking. Themineral medium used to grow the fungus contained 2g of (NH4)2SO4, 4 g of KH2PO4, 6 g of Na2HPO4, 0.2 gof MgSO4, 1 mg of FeSO4 7H20, 1 mg of CaCl2, 10,gof H3BO3, 10 ug of MnSO4, 70 ,ug of ZnSO4, 50 itg ofCuSO4, and 10 Mg of MoO3 per liter. The pH wasadjusted to 7.2, and the indicated amounts of glucoseand/or cutin hydrolysate were added.

Routinely, the fungus was grown in the mineralmedium containing 0.1% glucose for 3 days. As soonas glucose was exhausted, an aqueous dispersion ofcutin hydrolysate, the thin-layer chromatographicfractions isolated from the hydrolysate, or other lipidswere added. A concentrated solution (5 to 10 mg/mi)of actinomycin D, cycloheximide, or cordycepin wasadded at the indicated time to give the desired finalconcentration. At indicated time intervals, 1-ml por-tions of the culture medium were withdrawn and cen-trifuged, and the cutinase activity of the supernatantwas measured. The maximal level of cutinase (reachedin 4 days in all cases) was used as a measure ofinducing ability of the compound. To determinewhether production ofsome nonspecific esterase influ-enced these values, in all cases, the radial immunodif-fusion analyses were performed on the extracellularfluid on the 4th day, and the results were identical tothose obtained with PNB hydrolysis assay.

Preparation of aqueous dispersions of cutinhydrolysate and other lipids. Golden Delicious ap-ple cutin was prepared as described previously (24).Cutin powder (60 mesh) was suspended in 95% ethanol(30 ml) containing 10% (wt/vol) KOH, and the suspen-sion was refluxed for 16 h under a nitrogen atmos-phere. The resulting mixture was acidified with con-centrated HCl, and the products were extracted re-peatedly with chloroform. The combined chloroformextract was evaporated to dryness under vacuum. Asuspension of the hydrolysate (200 mg in 20 ml ofwater) was subjected to ultrasonic treatment with aBiosonik III 0.75-inch (about 1.9-cm) probe (about 15x 30 s with cooling after each), and appropriate por-tions of this dispersion were added to the culturemedia. The aqueous dispersions of other lipids wereprepared by ultrasonic treatment (10 x 5 s, with cool-ing in ice bath after each; Biosonik III, intermediateprobe) of 8 mg of 16-hydroxy C16 acid or molar equiv-alents of other compounds placed in 5 ml of water.

Thin-layer chromatography. The alkaline hy-drolysate of cutin was chromatographed on 1-mmlayers of Silica Gel G (20 by 20 cm) activated overnightat 110°C. Plates were developed with diethylether-hexane-methanol-formic acid (40:10:1:2, vol/vol). Thereference strip was spotted with a mixture of authenticsamples of c-hydroxy C16 acid, 10,16-dihydroxy C16acid, and 9,10,18-trihydroxy C18 acid. Components onthe reference strip were visualized by viewing theplates under UV light after spraying the plates witha 0.1% ethanolic solution of 2',7'-dichlorofluorescein.

The fractions corresponding to c-hydroxy C16 acid,10,16-dihydroxy C16 acid, and 9,10,18-trihydroxy C18acid were eluted from the silica gel with a 2:1 mixtureof chloroform and methanol.

Preparation of derivatives. Methyl esters wereprepared by refluxing the acids with 14% BF3 in meth-anol for 4 h; the reaction mixtures were added towater, and the products were extracted with chloro-form. The esters were purified by thin-layer chroma-tography. Acetates of hydroxyalkanes were preparedby treating the compounds with a 2:1 mixture of aceticanhydride and pyridine at room temperature over-night. After addition of water to the reaction mixture,the products were extracted into chloroform, and ex-cess pyridine was removed by extraction of the chlo-roform with acidified water.Glucose and protein determination. Determi-

nation of glucose was done by using a coupled glucoseoxidase-peroxidase system (25). The reaction mixturecontaining a portion of the growth medium, 0.0007% o-dianisidine, 6 U of glucose oxidase (A. niger, 200U/mg), and 6 U of horseradish peroxidase (260 pur-purogallin U/mg) in 1.0 ml of 0.083 M phosphatebuffer, pH 6.0, was incubated for 10 min at 23°C. Thecolor was stabilized by adding 0.1 ml of 4 N HCI, andthe mixture was allowed to stand for another 10 minbefore absorbance at 420 nm was read. The concentra-tion was calculated from a calibration curve, using D-glucose as standard. Protein was measured accordingto the method of Lowry et al. (14).Enzyme assays. Cutinase activity was measured

spectrophotometrically, using PNB as the model sub-strate as described before (16). This enzyme was alsoassayed by measuring the amount of radioactivityreleased from cutin biosynthetically labeled with[1-14C]acetate (16).

Immunological assays. Rabbit antibody (againstcutinase I) was prepared and Ouchterlony immunodif-fusion analysis was performed as described earlier(23). Quantitative analysis of cutinase production wasdone by a radial immunodiffusion method. In thismethod, a solution containing 2% Noble agar (DifcoLaboratories), 0.9% NaCl, and 0.01% thimerosal in 100mM Veronal buffer, pH 8.5, was boiled for 10min and cooled to 600C before the rabbit antiserumwas incorporated (4%, vol/vol). The solution wasmixed well and poured into petri dishes. After appli-cation of the antigens, the plates were allowed todevelop for 24 to 48 h. The plates were soaked in0.9% NaCl solution for 2 days and fixed with 7.5%acetic acid. Standard curves were prepared by usingserial dilutions of purified cutinase I isolated fromcutin-grown F. solani f. sp. pisi; a standard curve wasprepared for each plate.

Electrophoresis. Cationic polyacrylamide disc gelelectrophoresis was performed as described by Gabriel(2) with 7.5% polyacrylamide gels (0.5 cm in diameter,6.5 cm long) and 3 mA/tube. The gels were stainedwith 1% amido black in 7% acetic acid for 4 h andthen destained by an Ames model 1801 quick geldestainer.Sodium dodecyl sulfate-polyacrylamide gel electro-

phoresis was performed according to the method ofMaizel (15). The gel system containing 0.1% sodiumdodecyl sulfate consisted of a 12.5% polyacrylamide

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944 LIN AND KOLATTUKUDY

resolving gel (7 by 0.5 cm, pH 8.9) and a 2.5% stackinggel (1.5 by 0.5 cm, pH 7.2). Electrophoresis was per-formed at a constant current of 2 mA/tube over aperiod of 3 h. Protein bands were fixed and stainedby immersion of the gels in a solution of 0.05% Coo-massie brilliant blue in 45% methanol and 9% aceticacid for 24 h. The gels were destained with an aqueoussolution of 7.5% acetic acid and 5% methanol.

Determination of radioactivity. Solutions con-taining 14C or3H were dissolved in 10 ml of ScintiVerse(Fisher) and assayed for radioactivity in a Packardmodel 3255 Tri-Carb liquid scintillation spectrometer.

Following electrophoresis, stained gels werehardened by cooling with powdered dry ice and thencut into 3-mm slices with a razor blade. Gel sliceswere treated with 0.5 ml of 30% hydrogen peroxide at70°C for 2 h in counting vials. After cooling to roomtemperature, the contents were mixed with 10 ml ofScintiVerse and assayed for radioactivity as describedabove.

Incorporation of L-phenyl-[2,3-3Hlalanine intocutinase. F. solani f. sp. pisi cultures were grown inmineral medium (100 ml) containing 0.1% glucose for3 days in Roux bottles. At the end. of this period, 8mg of cutin hydrolysate and 25 ,uCi of labeled phenyl-alanine were added to the medium, and the cultureswere allowed to remain at room temperature for an-other 4 days. The extracellular fluid was collected byfiltration and lyophilized. The residue was dissolvedin 2 ml of distilled water and dialyzed against 0.01 Mphosphate buffer, pH 8.0, overnight. The protein so-lution was loaded on a Sephadex G-100 column (2 by35 cm) and eluted with the same buffer. The fractions(5 ml) were monitored for radioactivity and enzymaticactivity as described before.

RESULTS AND DISCUSSIONInduction of cutinase by cutin hydroly-

sate. To test whether cutin hydrolysis productsinduce cutinase, F. solani f. sp. pisi was grownin a medium containing 0.1% glucose as thecarbon source and 0.001% chemically preparedhydrolysate of cutin. Growth of the fungus didnot appear to be affected by the presence ofcutin hydrolysate. The extracellular fluid wasexamined for cutinase activity using a modelsubstrate, PNB, which is known to be a goodsubstrate for purified cutinase from F. solani f.sp. pisi (17, 18). Measurable amounts of PNBhydrolase activity could not be detected in themedium for 2 days. By the third day, PNBhydrolase activity could be detected in the me-dium, and the extracellular hydrolase activityincreased almost linearly for the next 3 days;no significant further increase could be detectedthereafter (Fig. 1). Even though an equally vig-orous growth of the fungus occurred in the glu-cose medium containing no hydrolysate, littlePNB hydrolase activity was detected in the ex-tracellular fluid. That the production of PNBhydrolase reflected production of cutinase wassuggested by the observation that the extracel-

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0 2 3 4 5 6 7

DAYS AFTER INNOCULATIONFIG. 1. Time course ofproduction of extracellular

cutinase by F. solani f. sp. pisi. The mineral medium(100 ml each) contained 0.1% glucose and the indi-cated amounts (in milligrams) of cutin hydrolysateat the time of inoculation (zero time) with 108 spores.Cutinase activity of the extracellular fluid was mea-sured with PNB. The inset shows the effect of theamount ofcutin hydrolysate on the rate ofproductionof cutinase (0- - -0) and on the total amount ofextracellular cutinase produced O@-). The slopeofthe linearportion (in arbitrary units) ofeach curveis designated as the rate of cutinase production.

lular fluid of the hydrolysate-containing culturecatalyzed hydrolysis of labeled cutin, whereasthe control cultures did not.The effect of the amount of cutin hydrolysate

added to the medium on the production of ex-tracellular cutinase was also determined (Fig.1). Since the amount of glucose added mightaffect the rate of cutinase production, equalquantities of glucose (0.1%) were used in allcultures. With this level of glucose, PNB hydro-lase production began by day 3 at all levels ofcutin hydrolysate used. However, the rate ofproduction of extracellular cutinase increasedlinearly with increasing amounts of hydrolysateup to about 80 ,ug/ml; further increase in theamount of hydrolysate resulted in no furtherincrease. The total amount of hydrolase pro-duced by the fungus also increased with increas-ing amounts of hydrolysate up to 80 ,ug/ml, andfurther increase in the concentration of cutinhydrolysate did not result in any further increasein the total amount of extracellular hydrolaseactivity. Thus, the rate of production and thetotal amount of hydrolase produced dependedon the concentration of cutin hydrolysate, anda saturation pattern of response was observedwith respect to the amount of hydrolysateadded.

Identification of the induced enzyme as

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INDUCTION OF CUTINASE BY CUTIN MONOMERS 945

cutinase. Even though cutinase is known tocatalyze hydrolysis of PNB by a mechanismsimilar, if not identical, to that involved in cutinhydrolysis (18), PNB hydrolase activity couldnot be taken as conclusive evidence for the in-duction of cutinase by cutin hydrolysate. There-fore, a time course of production of cutinase wasdetermined by a direct assay using hydrolysisof "C-labeled cutin. With such a technique, itwas found that cutinase began to appear in themedium by day 3; the hydrolase activity in-creased rapidly during the next 3 days, and,thereafter, no further increase was observed.This time course was identical to that observedwith PNB as a model substrate.

If the cutinase induced by the low levels ofcutin hydrolysate was immunologically similarto the cutinase generated during the growth ofthe fungus on cutin, it might be possible tomeasure the enzyme in the medium directly byimmunological techniques. To test this possi-bility, Ouchterlony double-diffusion analyseswere performed with the antiserum preparedagainst the major cutinase isozyme isolated fromthe extracellular fluid of cutin-grown F. solanif. sp. pisi (23). The cutinase, induced by cutinhydrolysate, strongly cross-reacted with anticu-tinase I, showing that the two enzymes wereimmunologically quite similar (Fig. 2). However,the precipitant lines formed by the two proteinsdid not appear to fuse completely; a spur was

often observed. Therefore, it appears that thetwo enzymes may not be immunologically iden-tical. In any case, the very strong similaritybetween the two proteins allowed us to developa radial immunodiffusion assay for the extracel-lular cutinase induced by cutin hydrolysate. Dif-fusion of cutinase I into the agar containing theantiserum against cutinase I resulted in immu-noprecipitant rings, which had areas propor-

tional to the concentration of the antigen ap-plied. This linear relationship (Fig. 2) was usedto determine the amount of cutinase releasedby the cells grown in glucose in the presence oflow levels of cutin hydrolysate. With such an

assay, the time course of production of extracel-lular immunologically cross-reacting protein wasdetermined. Such a protein appeared only afterabout 2.5 days; a rapid accumulation of thisprotein occurred during the next 3 days, and no

further change was observed during the subse-quent period. Thus, the results obtained withthe PNB hydrolase assay, with the labeled cutinhydrolysis assay, as well as with the radial im-munodiffusion assay, are in excellent agreement.The extracellular fluid of F. solani f. sp. pisi

grown in powdered cutin contained phenolic ma-terials which were most probably released fromcutin. These phenolic materials were associatedwith cutinase and thus complicated the proce-dure used for the purification of the enzyme(17). On the other hand, the extracellular fluidof the fungus grown in cutin hydrolysate was

essentially colorless, and the radial immunodif-fusion assays showed that at least 50% of theextracellular protein in the medium was cuti-nase. Electrophoresis of the concentrated extra-cellular fluid confirmed that the major proteinin the extracellular fluid had a mobility identicalto that of cutinase previously purified from cu-

tin-grown fungus (Fig. 3). Sodium dodecyl sul-fate electrophoresis also showed that the majorprotein in the extracellular fluid had a molecularweight quite similar to that of cutinase isolatedfrom the cutin-grown fungus (17).Thses findings suggested that the induction

of cutinase by cutin hydrolysate might be usedin studying the biosynthesis of cutinase. To testthis possibility, cutin hydrolysate and [3H]-phenylalanine were added after a 3-day growth

0.4

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0 50 00 150 200 250

pg/ml CUTINASE IFIG. 2. Ouchterlony double-diffusion showing cross-reactivity of cutinase induced by cutin hydrolysate

against the antiserum prepared against cutinase I isolated from cutin-grown F. solani f. sp. pisi (left). Centerwell, rabbit immunoglobulin G; top and bottom outer wells, cutinase I; left and right, cutinase induced bycutin hydrolysate. Both cutinase preparations were at 150 pg/mtl (Right) A typical standard curve obtainedfrom a radial immunodiffusion plate using cutin-grown cutinase I as reference protein. The area representsthe area of the precipitin ring minus the area of the center well.

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946 LIN AND KOLATTUKUDY

period in glucose. After 4 days of further incu-bation, the extracellular fluid was concentratedand dialyzed. Sephadex G-100 gel filtration (Fig.4, top), showed that the major radioactive com-ponent was coincident with the cutinase activity,strongly suggesting that the major protein (cu-tinase) had the bulk of the 3H found in theextracellular proteins. Electrophoresis of the cu-

tinase fraction showed that virtually all of theradioactivity of this fraction was contained incutinase (Fig. 4, bottom). Thus, synthesis ofcutinase was demonstrated, and it appears thatthis system might be quite suitable for biosyn-thetic studies on cutinase.Repression of cutinase synthesis by glu-

cose. It was previously observed that glucosein the medium inhibited the production of cuti-nase by F. solani, f. sp. pisi induced by cutin(16). To determine whether the 3-day lag in theproduction of cutinase by the fungus grown inglucose in the presence of cutin hydrolysate wasdue to a similar repression by glucose, the timecourse of depletion of glucose in the mediumwas determined. About 24 h after inoculation,the level of glucose began to decrease rapidly,and by day 3 measurable amounts of glucosecould not be found in the medium. When cutinhydrolysate was added after depletion of glu-cose, PNB hydrolase activity began to appearin the medium without the lag observed in Fig.1. Thus, the lag in the production of the extra-

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FIG. 3. Polyacrylamide disc gel electrophoresis ofpurified cutinase I (30 j,g) from cutin-grown cells(right), the crude cutinase induced by cutin hydroly-sate (middle; 45 jig ofprotein from the crude extra-cellular fluid), and sodium dodecyl sulfate-polyacryl-amide gel electrophoresis of the crude cutinase (50,ig) induced by the hydrolysate (left).

0 2 4 6MOBILITY (cm)

FIG. 4. Sephadex G-100 gel filtration of the extra-cellular fluid from F. solani f. sp. pisi, induced bycutin hydrolysate to generate cutinase in thepresenceof tritiated phenylalanine (top); polyacrylamide discgel electrophoresis of the cutinase fraction obtainedfrom the gel filtration step (bottom). The bar graphindicates distribution of radioactivity in the gel.

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INDUCTION OF CUTINASE BY CUTIN MONOMERS

cellular hydrolase most probably represents thetime required to deplete the supply of glucose,which probably causes repression of synthesisof the hydrolase. This conclusion is consistentwith observation that, at all levels of cutin hy-drolysate added to the medium, the lag periodfor the appearance of extracellular cutinase wasconstant (Fig. 1).To test whether the lag in the production of

cutinase by the cells grown in glucose in thepresence of cutin hydrolysate truly representsrepression by glucose as suggested above, cutinhydrolysate was added with additional glucoseto cultures immediately after depletion of glu-cose originally present in the medium. As notedabove, the 0.1% glucose originally used in themedium was depleted by day 3, and at that timethe optimal amount of hydrolysate (8 mg/100ml) was added with different levels of additionalglucose. Without any addition of glucose, thehydrolysate caused a rapid increase in extracel-lular cutinase, which leveled off in about 3 days.With additional amounts of glucose, the produc-tion of cutinase was delayed (Fig. 5). Increasingamounts of glucose added with the cutin hy-drolysate caused increasing delays in the ap-pearance of extracellular cutinase. However, therate of appearance of cutinase and the totalamount of extracellular cutinase produced werenot significantly altered by the addition of glu-cose. Growth of the fungus was relatively con-stant in all cultures. For example, the final dryweight of the mycelia was 0.5 g per 100 ml incultures which received no additional glucose,whereas 0.55 g per 100 ml was obtained fromthe cultures receiving 0.5% glucose. Thus, it ap-pears that the delay was probably caused bythe repression by glucose, and that the increas-ing delay observed with increasing glucose con-

centration might simply represent the timetaken for depletion of this nutrient. In fact,measurement of glucose in the medium indi-cated that, in all cases, the rapid rise in extra-cellular cutinase occurred immediately after de-pletion of glucose (Fig. 5). Therefore, it appearsquite clear that glucose is a repressor of thesynthesis of cutinase; once the repressor is de-pleted, cutin hydrolysate induces the synthesisof cutinase. Thus, these results strongly suggestthat cutinase synthesis is under dual control.Effect of actinomycin D, cordycepin, cy-

cloheximide, and deoxyglucose on cutinaseproduction. To determine whether the induc-tion of cutinase by cutin hydrolysate involvestranscription sensitive to actinomycin D, thischemical was added to the medium after deple-tion of glucose, and the time course of produc-tion of cutinase was followed. When actinomycinD was added simultaneously with the inducer(cutin hydrolysate) or any time after the addi-

tion of the inducer, no inhibition of productionof cutinase was observed (Table 1). However,some inhibition of cutinase production was ob-served when actinomycin was added after thedepletion of glucose, but 24 or 48 h prior to theaddition of the inducer. Thus, occurrence of anactinomycin D-sensitive step after the additionof the inducer could not be demonstrated underthe present experimental conditions.Cordycepin is known to be an inhibitor of

synthesis of poly(A)-containing mRNA and thuscould possibly inhibit cutinase production (6).To test this possibility, cordycepin was addedto the medium after glucose depletion, simulta-neously with the inducer or a few hours after

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DAYS AFTER HYDROLYSATE ADDITIONFIG. 5. Effect of addition of glucose with the in-

ducer on the appearance of extracellular cutinase.The cultures were grown in mineral medium contain-ing 0.1% glucose for 3 days, after which 8 mg ofhydrolysate was added to each flask (100 ml) withglucose at concentrations indicated (zero time). Thedashed lines represent the glucose level deterrninedby glucose oxidase-peroxidase method. The solidlines represent the cutinase activity (assayed by PNBhydrolysis).

TABLE 1. Effect of actinomycin D on the inductionof cutinase by cutin hydrolysate in F. solani f. sp.

pisia

Time of addition of acti- Cutinase activity at time indi-nomycin D __ cated

(days) 8 h 16 h 24 h 48 h

2 Days prior to inducer 0.70 2.1 2.90 3.051 Day prior to inducer 0.90 2.30 3.10 3.15Simultaneously with in-ducer .............. 1.30 2.80 3.40 3.60

1 Day after inducer .... 1.30 2.80 3.60 3.852 Days after inducer 1.65 3.30 3.95 4.05No actinomycin ....... 1.35 2.75 3.50 3.65

a Actinomycin D (5 ,ug/ml) was added at the indi-cated number of days after the addition of the inducer(8 mg of cutin hydrolysate), and the extracellularcutinase activity was measured at the time intervalsafter the addition of the inducer. Enzyme activity isexpressed as change in absorbancy at 405 nm perminute per milliliter.

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948 LIN AND KOLATTUKUDY

the addition of the inducer (cutin hydrolysate),and the production of cutinase was measured.Addition of cordycepin with the inducer stimu-lated the production of cutinase, and cordycepincaused a doubling in the total amount of cutinasegenerated (Fig. 6). When the addition of cordy-cepin was delayed for 4 h after the addition ofthe inducer, the stimulation of cutinase produc-tion was only about one-half of that observedwhen cordycepin was added with the inducer.An 8-h delay in the addition of cordycepin afterthe addition of the inducer resulted in no meas-urable effect. Even though a molecular levelexplanation for such observations cannot be pro-vided, it appears possible that inhibition of syn-thesis of poly(A)-containing mRNA resulted inan enhancement of competing syntheses involv-ing other RNAs, including those involved in thesynthesis of cutinase. The recent report thatcordycepin stimulates in vitro translation of sev-eral types of mRNA in both the wheat germsystem and reticulocyte lysate (10) provides aprobable explanation for the stimulation of cu-tinase synthesis observed in the present study.

Since exogenous labeled amino acids were in-corporated into cutinase when the glucose-de-pleted cultures were treated with cutin hydrol-ysate, it appeared that protein synthesis, whichoccurred after the addition of the inducer, wasinovlved in the appearance of extracellular cu-tinase. To further test this possibility, cyclohex-imide was added at various time intervals afterthe addition of inducer to glucose-depleted me-dia, and the subsequent appearance of extracel-lular cutinase was followed (Fig. 7). Addition ofcycloheximide simultaneously with the inducerresulted in complete inhibition of appearance ofcutinase in the medium. Similarly, addition ofcycloheximide up to 12 h after the addition ofthe inducer virtually prevented any further in-

5 CORDYCEPIN

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0 12 24 36 48INDUCTION PERIOD(Hr)

FIG. 6. Effect of cordycepin on the production ofextracellular cutinase. Cordycepin (33 ,ug/ml, finalconcentration) was added at the times indicated afterthe addition of hydrolysate (8 mg/100 ml).

cn' 12-Hr

< 34~~~~~~~~4H

0-Hr0 8 16 24

INDUCTION PERIOD (Hr)FIG. 7. Effect of cycloheximide on the production

of extracellular cutinase by F. solani f. sp. pisi. Zerotime refers to the time of addition of hydrolysate;after the number of hours shown on each line, cyclo-heximide (6 pig/ml) was added and the production ofcutinase was monitored.

crease in the level of extracellular cutinase.About 12 h after the addition of cycloheximide,some ability to recover from the effects of thisinhibitor appeared. In any case, it is quite clearthat a cycloheximide-sensitive process (proteinsynthesis) is directly involved in the productionof extracellular cutinase induced by cutin hy-drolysate.

Cutinase is known to be a glycoprotein (11,12), and deoxyglucose is known to inhibit gly-cosylation of proteins and thus inhibit excretionof proteins (1, 3, 9, 13, 20). Therefore, the effectof 2-deoxyglucose on the induced production ofextracellular cutinase by F. solani f. sp. pisi wasdetermined. Since glucose is known to be a re-pressor of cutinase production, there is a possi-bility that 2-deoxyglucose might also have asimilar effect. Therefore, the effect of glucosewas compared with that of equal quantities of2-deoxyglucose. Deoxyglucose inhibited the pro-duction of extracellular cutinase when it wasadded together with the inducer, and the inhib-itory effect of deoxyglucose was much more se-vere than that observed with equal amounts ofglucose (Fig. 8). For example, after 2 days ofinduction, the amount of extracellular cutinasefound in cultures containing 0.1% glucose wasnearly the same as that found in controls,whereas in cultures containing 0.1% deoxyglu-cose, less than half as much cutinase was found.Similarly, in 2 days the amount of cutinase inthe deoxyglucose-containing medium was onlyone-fifth of that found in the glucose-containingmedium when 0.2% of each sugar was used. Inall cases the level of extracellular cutinasetended to recover, and it approached controlvalues in about 5 days. It is clear that deoxyglu-cose is the more potent inhibitor of the produc-tion of extracellular cutinase. However, definiteconclusions concerning the mechanism of inhi-

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INDUCTION OF CUTINASE BY CUTIN MONOMERS

°E 23, 4 5W~~~~~~

E ,,

2 3 4 5INDUCTION PERIOD (Days)

FIG. 8. Effect of deoxyglucose on the productionofextracellular cutinase. When glucose was depleted,cutin hydrolysate was added (zero time) without anyother addition (control), or with the indicatedamounts ofglucose (Gl) or 2-deoxy-D-glucose (DOG).

bition of induction cannot be drawn. 2-Deoxy-glucose might have inhibited glycosylation andtherefore excretion of cutinase, or it might haveacted as a repressor of synthesis of cutinase,just as glucose. The probable metabolic inert-ness of deoxyglucose might have made it appearas a more potent inhibitor than glucose.Cutinase-inducing ability of cutin mono-

mers and analogs. Induction of cutinase pro-duction by the hydrolysate of cutin provided anopportunity to determine the nature of the com-ponents responsible for induction. Thin-layerchromatographic fractionation of the hydroly-sate allowed us to determine the relative abilityof the various fractions to induce cutinase pro-duction. Among the individual fractions, the w-hydroxy acid fraction (which was a mixture ofw-hydroxypalmitic acid, o-hydroxystearic acid,w-hydroxyoleic acid, and w-hydroxylinoleic acid)was the most effective for the induction of cuti-nase (Table 2), whereas the more polar 9,10,18-trihydroxy C18 acid fraction was the least effec-tive. No individual fraction was as effective asthe total hydrolysate in inducing cutinase. How-ever, mixtures ofindividual fractions were nearlyas effective as the total hydrolysate, and there-fore it is clear that the aliphatic componentscontained in these isolated fractions were, in-deed, responsible for the induction broughtabout by the total hydrolysate.To determine which functional groups in the

monomers might be important to the ability toinduce cutinase, 16-hydroxypalmitic acid waschosen as the reference monomer. Esterificationof the carboxyl group did not substantially alterits ability to induce cutinase, whereas acetyla-tion of the hydroxyl group resulted in a sharploss in activity. Modification of both functionalgroups did not decrease the activity much fur-ther than that observed with the acetylated acid

(Table 2, experiment II). Replacement of thehydroxyl group by a hydrogen or by anothercarboxyl group virtually eliminated the cutinase-inducing activity. Therefore, the hydroxyl groupappears to be important to the cutinase-inducingactivity. Neither 2-hydroxypalmitic acid nor9,10-dihydroxystearic acid was as effective as16-hydroxypalmitic acid in inducing cutinase(Table 2, experiment III). Therefore, the posi-tion of the hydroxyl group in the aliphatic chainis critical to the cutinase-inducing activity. Eventhough metabolic interconversions of the deriv-

TABLE 2. Induction of cutinase by cutin monomersand analogs in F. solani f. sp. piSia

Induction ofInducer cutinase

(%)Exp I

Cutin hydrolysate (8 mg) ..........w-Hydroxy acid fraction (8 mg) .....10,16-Dihydroxy C16 acid fraction (8mg) ...........................

9,10,18-Trihydroxy C,8 acid fraction(8 mg) ............ .. .....

w-Hydroxy acid (3 mg) + 10,16-dihy-droxy C16 acid (3 mg) + 9,10,18-trih-ydroxy C18 acid (2 mg) ...........

w-Hydroxy C16 acid (8 mg) .........Exp II

16-Hydroxy C16 acid ...............16-Hydroxy C16 acid methyl ester16-Acetoxy C16 acid ..............16-Acetoxy C16 acid methyl ester ...

Exp III16-Hydroxy C16 acid ...............2-Hydroxy C16 acid ................9,10-Dihydroxy C18 acid ...........C16 acid ..........................C16 dioic acid .....................C16 alcohol .......................

Exp IVC8 alcohol ......................C0o alcohol ....................C12 alcohol ............. .....

C14 alcohol .... .. ... .... .....

C16 alcohol .............. .....

C,8 alcohol ............

C20 alcohol ....... ....

C22 alcohol ..................C24 alcohol ...............C26 alcohol ......................C28 alcohol ....................

10061

53

17

9143

100945851

100311298

110

0

0

0

53100472947353518

a In all cases, after the glucose in the medium wasexhausted, 8 mg or the indicated amount of lipid and,in experiments II, III, and IV, 8 mg of 16-hydroxy C16acid or molar equivalents of the other compoundswere added. In each experiment, the value obtainedwith a selected compound is taken as 100, and thoseobtained with other compounds are expressed as per-centages.

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950 LIN AND KOLATTUKUDY

atives could significantly affect their cutinase-inducing activities, the results presented heresuggest that the hydroxyl group at the w-carbonis the most important factor determining thecutinase-inducing activity of the monomer; thecarboxyl group does not appear to be significantfor this purpose. In fact, hexadecanol, whichrepresents replacement of the carboxyl groupby a hydrogen, was just as effective as w-hydrox-ypalmitic acid in inducing cutinase (Table 2,exeriment III).

Since an aliphatic chain with a primary hy-droxyl group appears to be the only requirementfor the induction of cutinase in F. solani f. sp.pisi, the optimal size of the required aliphaticchain could be readily determined (Table 2,experiment IV). C12 and shorter alcohols weretotally ineffective; C16 alcohol was most effective,and other aliphatic alcohols up to C28 were lesseffective in inducing cutinase.The results presented here show that a low

level of cutin hydrolysate induces the synthesisof cutinase. Therefore, it appears that the chem-ical signal which transmits the information con-cerning the nature of the polymer substrate toF. solani f. sp. pisi is one or more componentscontained in cutin hydrolysate. When the fungusis placed in a medium containing cutin as thesole source of carbon, very low levels of hydro-lytic enzymes, including cutinase, are probablyexcreted into the medium. This low level ofcutinase could generate a low level of hydroxy-fatty acids which enter the cell and induce thesynthesis of cutinase. When other nutrients suchas glucose are present in the medium, they actas repressors. Thus, production of cutinase isunder dual control. This might be a generalmechanism by which microorganisms sense thenature of nontransportable polymers availablein their environment and produce extraceilularhydrolases to metabolize such polymers.

Strong evidence that cutinase might be in-volved in the penetration of the plant cuticleby phytopathogenic fungi has been presented(21). The present finding, that fairly simple mol-ecules, including those which are known to bepresent in the plant surface lipids (8), can inducecutinase production by phytopathogenic fungi,raises the possibility that plant surface lipidsmight play a communicative role in pathogen-esis. If the production of cutinase is, in fact,essential for the entry of the pathogen into theplant, the composition of the cuticular wax couldplay a key role in inducing the enzyme in thepathogen which lands on the surface. Alterna-tively, antagonists of the inducers could be pres-ent in the surface wax and thus prevent induc-tion of cutinase and consequently prevent infec-tion. Such possibilities have not yet been tested.

ACKNOWLEDGMENTSThis work was supported in part by grant PCM 74-09351-

A03 from the National Science Foundation.We thank Rodney Croteau for a critical reading of the

manuscript.LITERATURE CITED

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17. Purdy, R. E., and P. E. Kolattukudy. 1975. Hydrolysisof plant cuticle by plant pathogens: purification, aminoacid composition, and molecular weight of two isozymesof cutinase and a nonspecific esterase from Fusariumsolani f. pisi. Biochemistry 14:2824-2831.

18. Purdy, R. E., and P. E. Kolattukudy. 1975. Hydrolysisof plant cuticle by plant pathogens. Properties of cuti-nase I, cutinase II, and a nonspecific esterase isolatedfrom Fusarium solanipisi. Biochemistry 14:2832-2840.

19. Reese, E. T. 1977. Degradation of polymeric carbohydrateby microbial enzymes. Rec. Adv. Phytochem. 11:311-367.

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21. Shaykh, M., C. Soliday, and P. E. Kolattukudy. 1977.Prooffor the production of cutinase by Fusarium solanif. pisi during penetration into its host, Pisum sativum.Plant Physiol. 60:170-172.

22. Shishyama, J., F. Araki, and S. Akai. 1970. Studieson cutin-esterase. II. Characteristics of cutin-esterasefrom Botrytis cinera and its activity on tomato-cutin.Plant Cell Physiol. 11:937-945.

23. Soliday, C. L., and P. E. Kolattukudy. 1976. Isolation

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