1

Cutin from Plants

Prof. Dr. Pappachan E. KolattukudyThe Ohio State University, Columbus, Ohio 43210, USA; Tel.: �01-614-292-5682;Fax: �01-614-292-5379; E-mail: kolattukudy.2@osu.edu

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2 Historical Outline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

3 Occurrence and Ultrastructure of Cutin . . . . . . . . . . . . . . . . . . . . . . 3

4 Isolation of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

5 Depolymerization of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55.1 Chemical Depolymerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55.2 Enzymatic Depolymerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

6 Monomer Composition of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

7 Structure of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

8 Biosynthesis of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118.1 Biosynthesis of the C16 Family of Cutin Monomers . . . . . . . . . . . . . . . 128.2 Biosynthesis of the C18 Family of Cutin Monomers . . . . . . . . . . . . . . . 158.3 Synthesis of Cutin from Monomers . . . . . . . . . . . . . . . . . . . . . . . . 17

9 Cutin Biodegradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199.1 Cutin Degradation in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199.2 Degradation of Cutin by Animals . . . . . . . . . . . . . . . . . . . . . . . . . 209.3 Cutin Degradation by Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . 209.4 Fungal Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229.4.1 Purification and Molecular Characterization of Fungal Cutinases . . . . . . . 229.4.2 Catalytic Properties of Cutinase . . . . . . . . . . . . . . . . . . . . . . . . . . . 24

1

10 Function of Cutin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2710.1 Material Exchange with the Environment . . . . . . . . . . . . . . . . . . . . . 2810.2 Low-temperature Adaptation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2810.3 Role of Cutin in the Interaction with Microbes . . . . . . . . . . . . . . . . . 2910.3.1 Regulation of Cutinase Gene Transcription . . . . . . . . . . . . . . . . . . . . 3010.4 Cutin Required for Proper Development of Plant Organs . . . . . . . . . . . 32

11 Potential Commercial Use for Cutin and Cutinase . . . . . . . . . . . . . . . . 33

12 Outlook and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33

13 Patents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33

14 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35

CD circular dichroismCMC critical micellar concentrationCPMAS-NMR cross-polarization magic angle spin nuclear magnetic resonanceCRE cutin- responsive elementCREBP cutin-responsive element binding proteinCTF cutinase transcription factorEDC 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimideFTIR Fourier transform infraredGC/MS gas-liquid chromatography/mass spectrometryHPTLC high-performance thin-layer chromatographyKi inhibition constantLSIMS liquid secondary ion mass spectrometryNMR nuclear magnetic resonancePBP palindrome binding proteinRadio-GLC radioactivity detector gas-liquid chromatographySDS±PAGE sodium dodecyl sulfate±polyacrylamide gel electrophoresisTLC thin-layer chromatography

1

Introduction

The outer envelope of organisms consists ofa polymeric structural component which, interrestrial organisms, is made waterproofwith a complex mixture of nonpolar lipidscollectively called waxes. Proteins or carbo-hydrate polymer (chitin) serves as the struc-tural component in animals, whereas abiopolyester (cutin) ± which is derived from

cellular lipids ± serves as the structuralcomponent of the outer envelope (the cu-ticle) of higher plants. Development of thecuticle, in which cutin is embedded in waxesto make an efficient barrier against desic-cation, allowed plants to move to the landabout 400 million years ago. The widespreadoccurrence of polyesters in plants is notwidely known because textbooks in bio-chemistry and other general fields of biologyrarely mention cutin. Indeed, if the natural

1 Cutin from Plants2

occurrence of polyesters were known, thensynthetic polyesters would have been devel-oped earlier than polyamide polymers.

2

Historical Outline

Studies on the nature of plant cuticle werestarted in the 19th century. In these earlystudies, the thin films which remained aftertreatment of the aerial parts of plants withstrong acids were considered to be the truecuticles. Since the cuticles of leaves wereknown to be readily attacked by alkalinesolutions, the acid-resistant material fromthe leaves was treated with alkali, and thisresulted in the generation of soluble soaps.The term cutin came from the name cutose,this being used originally to describe thematerial that formed part of the epidermis ofleaves andwhich resisted the action of strongacids (Martin and Juniper, 1970).

3

Occurrence and Ultrastructure of Cutin

Cutin constitutes the structural componentof the cuticles of higher plants. Even thehigher plants that live under water such asthe sea grass, Zoestra marina, which growssubmerged on coastal shorelines, have cutincomposed of the same type of monomers asthose found on land plants. There is evi-dence that cuticle exists in lower plants suchas mosses, the lycopods, the ferns, and

liverworts (Holloway, 1982a). Extensive ex-amination of the cuticular structures madeduring the modern era demonstrates thatthe cutin-containing layer is attached to theepidermal cell wall with a pectinaceous gluelayer (Figure 1). However, this is an over-simplified general picture, and in reality theboundaries between the cell wall, pectin andcutin layers are not always clearly definedbecause there is usually some intermingling,especially near the boundaries. A micro-scopically distinct cuticular layer can bedistinguished in most plant organs. Cutin-containing layers are found not only on thesurfaces of all aerial parts of plants includingstems, petioles, leaves, flower parts, fruits,and some seed coats, but also on internalparts such as juice sacks of citrus (Kolat-tukudy, 2001). The thickness of the polymerlayer varies among species and amongorgans in the same plant. In higher plantleaves, the thickness ranges from 0.5 to14 �m, with �20 to 600 �g cutin per cm2 ofthe surface area. In some fruits with a well-developed cuticle, the cutin content mayreach 1.5 mg cm�2. In lower plants thecuticle in usually very thin (�0.1 �m).Electron microscopic examination of the

cuticular area shows that the cutin-contain-ing layer has a mostly amorphous appear-ance (Figure 2A). Scanning electron micro-scopic examination of the cuticular surfacethat is attached to the cell wall shows cellularoutlines and protrusions of the polymermatrix into the intercellular junctions, anddemonstrates how the polymer is molded tofit into the intercellular spaces (Figure 2B).Within this matrix, lamellae and fibrillae

3 Occurrence and Ultrastructure of Cutin 3

Fig. 1 Schematic representation ofthe cuticle.

may be found and, when present, suchstructures form an anastomosing system.Based on the presence of such features andtheir sites of occurrence within the cuticle,the cuticular ultrastructures have been clas-sified into six groups (Holloway, 1982b). Thebiological significance of such differentappearances is not clear, but the reticularand the anastomosing structures may play a

role in the transport of materials through thecuticle.

4

Isolation of Cutin

Disruption of the pectinaceous glue thatattaches the cuticle to the epidermal layer by

1 Cutin from Plants4

Fig. 2 (a) Electron micrographsillustrating amorphous (top, Tropaeo-lum majus) and lamellar (bottom,Atriplex semibaccata) cuticle.(b) Scanning electron micrograph ofthe underside of tomato fruit cutinshowing the protrusions that help toanchor the polymer to the fruit byfitting into the intercellular grooves.Cu, cuticle; CW, cell wall.

enzymes or chemicals is required to releasethe cuticular layer. The most commonlyused, gentle methods are treatment withammonium oxalate/oxalic acid or pectin-degrading enzymes (Kolattukudy and Wal-ton, 1973; Holloway, 1982a). The releasedcuticular layer can be physically separatedand subjected to further treatment withcarbohydrate-hydrolyzing enzymes to re-move additional carbohydrates. A thoroughextraction with chloroform is required toremove the soluble waxes embedded in thepolymer matrix. Even after several days ofextraction, apple cutin showed X-ray diffrac-tion caused by the residual waxes thatremained trapped within the polymermatrix(P. E. Kolattukudy, unpublished results).Additional Soxhlet extractions for manymore days gradually decreased the diffrac-tion caused by the crystalline waxes buried inthe polymer matrix. This problem is moresignificant in the case of the thicker cuticlefound in fruits. The final product can bepowdered and used for chemical and/orphysico-chemical studies.

5

Depolymerization of Cutin

Cutin can be depolymerized by cleaving theester bonds chemically to release free mono-mers or their derivatives, depending on thechemical cleavage method used. This poly-ester can also be depolymerized with en-zymes that catalyze the hydrolysis of esterbonds.

5.1

Chemical Depolymerization

Since cutin is largely a polyester, ester bondcleavage methods can release the mono-mers. Most commonly used methods in-clude alkaline hydrolysis, transesterification

with methanol containing boron trifluorideor sodium methoxide, reductive cleavage byexhaustive treatment with LiAlH4 or LiAlD4

in tetrahydrofuran, or with trimethylsilyliodide in organic solvents. These methodsyieldmonomers or their derivatives based onthe depolymerization method (Figure 3).Cutin can contain functional groups thatare not stable to the depolymerizationmethods such as epoxides and aldehydes.During depolymerization, such functionalgroups may be converted to derivatives thatare useful for identification of the functionalgroup(s) originally present in the polymer.For example, during reductive depoly-merization with LiAlD4, the epoxide andaldehyde would generate D-labeled deriva-tives; subsequently, mass spectrometry canreadily locate the label in the reducedproduct and thus identify the epoxy or oxofunction originally present in the polymer.In fact, such deuterium labeling is whatestablished the 18-hydroxy-9,10-epoxy-C18

acid as a widely occurring cutin component(Walton and Kolattukudy, 1972; Kolattukudyand Walton, 1973). Similarly, �-oxo acid as amajor component of cutin from embryonictissue was discovered by the deuteriumlabeling that occurred during the reductivedepolymerization. Methanolysis of the oxir-ane would generate a methoxy group vicinalto a carbinol identifying the location of theepoxide in the original polymer. Functionalgroups such as aldehydes can also bederivatized, first by making derivatives suchas an oxime, followed by depolymerization,to yield an identifiable derivative (Kolat-tukudy, 1974). Most cutin preparations leavebehind insoluble residues after exhaustivetreatments with ester-hydrolyzing reagents(see Section 7).

5 Depolymerization of Cutin 5

5.2

Enzymatic Depolymerization

Since the majority of monomers are heldtogether by ester linkages, esterases cancleave the polymer. Pancreatic lipase canhydrolyze cutin, thereby releasing oligomersandmonomers. Evidence was presented thatbile salts stabilize the enzyme at the surface

of the insoluble polymer and that theinteraction of the polymer surface withlipase±colipase±bile salt system is similarto that observed with triglycerides (Brownand Kolattukudy, 1978b). A special polyester-hydrolyzing lipase ± cutinase ± has evolvedin microorganisms such as bacteria andfungi (see Section 9). The first cutinase

1 Cutin from Plants6

Fig. 3 Chemical methods used to depolymerize the polyesters (top); thin-layer and gas-liquid chromato-grams (as trimethylsilyl derivatives) of the monomer mixture obtained from the cutin of peach fruits byLiA1H4 treatment (bottom). In the thin-layer chromatogram the five major spots are, from the bottom, C18

tetraol, C16 triol, and C18 triol (unresolved), diols, and primary alcohol.N1, C16 alcohol;N2, C18 alcohol;M1, C16

diol; M2, C18 diol; D1, C16 triol; D2 and D3, unsaturated and saturated C18 tetraol, respectively.

purified and characterized was a fungalcutinase, and this enzyme was shown torelease oligomers and monomers. Sincecutinase preferentially hydrolyzes primaryalcohol esters, enzymatically isolated oligo-mers are suitable for structural studiesinvolving cross-links via secondary alcoholester bonds (see Section 7).

6

Monomer Composition of Cutin

The complex mixture of monomers pro-duced by the chemical depolymerizationmethods can be separated by thin-layerchromatography (TLC) on silica gel intodifferent classes (depending on their polar-ity) such as �-hydroxy-, dihydroxy-, trihy-droxy-, and fatty acid derivatives (Kolat-tukudy and Walton, 1973; Holloway,1982a). The selection of the solvent systemsvaries depending on the depolymerizationmethod used. The individual fractions re-covered from TLC are then subjected tocombined gas-liquid chromatography/massspectrometry (GC/MS) after making appro-priate derivatives, again depending on thedepolymerization method used. For exam-ple, transesterificationmethod yieldsmethylesters, and therefore derivatization of thefree hydroxyl groups by trimethylsilylationprovides the appropriate derivatives forcombined GC/MS. Hydrogenolytic cleavagewith LiAlD4 produces reduced monomersthat can be trimethylsilylated before GC/MS.If a hydrolytic method is used, both thecarboxyl groups and the hydroxyl groupsneed to be derivatized. Even though trime-thylsilylation derivatizes carboxyl and hy-droxyl groups, trimethylsilyl (TMSi) deriva-tives of methyl esters give more diagnosticmass spectra. Retention times are useful inidentifying monomers when authenticstandards are available. However, mass

spectrometry is necessary to identify reliablythemonomer structure; henceGC/MS is themethod of choice for determining the struc-ture and composition ofmonomers. Usually,the mixture of monomers obtained by one ofthe depolymerization methods can be deriv-atized and directly analyzed by combinedGC/MS, without preliminary TLC. Thehighly preferred �-cleavage on either sideof the trimethylsiloxyl groups of the mid-chain oxidized fatty acid derivatives makes itrelatively easy to identify the location of themid-chain hydroxyl groups.The results obtained from such analysis

performed on cutin from many plants showthat this plant polyester is composed ofmainly a C16 family and a C18 family ofmonomers (Figure 4) (Kolattukudy andWal-ton, 1973; Holloway, 1982a; Kolattukudy,2001). The most common major compo-nents of the C16 family of monomers are 16-hydroxyhexadecanoic acid and 9 or 10,16-dihydroxyhexadecanoic acid. This dihydroxyacid is usually the dominant component,and usually there is a mixture of mid-chainpositional isomers. In some cases, other C16

derivatives can be significant or majorcomponents. Examples include 16-hydroxy-10-oxo-C16 acid in citrus cutin and 16-oxo-9,or 10-hydroxy-C16 acid in the very youngleaves and embryonic shoots ofVicia faba. Insome cases (especially lower plants), areduced C16 monomer, such as 1,8,16-hexa-decanetriol, is a major component. Theprimitive cutin in ferns and lycopods ischaracterized by large amounts of 16-hy-droxyhexadecanoic acid and other�-hydroxyacids. The most commonmajor members ofthe C18 family of cutin monomers include18-hydroxy-C18-9-enoic acid, 18-hydroxy-C18-9,12-dienoic acid, 18-hydroxy-9,10-epoxy-C18 acid, 18-hydroxy-9,10-epoxy-C18-12-enoic acid, 9,12,18-trihydoxy-C18 acid,and 9,10,18-trihydroxy-C18-12-enoic acid. Insome plants such as Rosmarinus officinalis,

6 Monomer Composition of Cutin 7

9,10,18-trihydroxy-12,13-epoxy-C18 acid and9,10,12,13,18-pentahydroxy-C18 acids arefound (Croteau and Kolattukudy, 1974a).The �-oxo derivatives of the C18 family ofacids may also occur in small quantities.Most plants contain differentmixtures of thetwo families ofmonomers, with each speciesshowing a characteristic mixture of compo-nents. However, the monomer compositiondepends on the anatomic location within aplant. For example, the content of the majorC18 family of cutin monomers in Maluspumila cutin from fruit, leaf, stigma andflower petal was found to be 73%, 35%, 14%,and 12%, respectively (Espelie et al. 1979).Developmental changes in cutin composi-tion can also occur, as demonstrated with thedeveloping V. faba leaves (Kolattukudy,1974). Comparison of cutin composition ofmany plants and organs within them sug-gest that fast-growing plants and organs tendto have a higher content of the C16 family ofmonomers. For example, Arabidopsis leafcutin is composed of 9,16- and 10,16-dihydroxy-C16 acid as a major monomer withless amounts of �-hydroxy-C16 and C18 acidsand 18-hydroxy-9,10-epoxy-C18 acid and9,10,18-trihydroxy-C18 acid as minor compo-nents (P. E. Kolattukudy, unpublished re-

sults). Cutin from lower plants tend to havesome unusual monomers and less hydroxy-latedmonomers. Dicarboxylic acids and verylong-chain (�C18) fatty acid derivatives thatare major components of suberin, are onlyminor components in cutin. Even thoughmost of the monomers are held together byester linkages in the polymer, virtually allcutin preparations leave significant amountsof residual insoluble material after exhaus-tive treatments with all ester-cleaving re-agents as discussed below; the compositionof thismaterial is not well understood (Crisp,1965; Kolattukudy and Walton, 1973).

7

Structure of Cutin

Being an insoluble, amorphous polymer,only a limited number of methods can beapplied to investigate the structure of thispolymer. Transmission Fourier transforminfra-red (FTIR) spectra of isolated cutinshow absorbances indicative of hydroxyl(3300 cm�1), aliphatic C±H stretch(2924 cm�1 and 2852 cm�1), ester carbonyl(1731 cm�1), and C±O ester (1167 cm�1) asexpected (Villena et al., 2000). Some general

1 Cutin from Plants8

Fig. 4 Structure of themost common major mono-mers of cutin.

structural information was obtained by test-ing for the presence of free functionalgroups in the polymer by indirect methods.Two methods were used to modify the freehydroxyl groups present in the polymer,followed by depolymerization and measure-ment of the monomers that had beenmodified in each class of monomer. Onemethod used oxidation of the free hydroxylgroups with CrO3-pyridine followed bydepolymerization with NaOCH3 in anhy-drousmethanol and analysis of the carbonyl-containing monomers (Deas and Holloway,1977). The other involved mesylation of thefree hydroxyl groups in the polymer bytreatment with methanesulfonyl chloridefollowed by depolymerization with LiAlD4

(Kolattukudy, 1977). This procedure resultedin replacement of each free hydroxyl groupin the polymer with a deuterium atom thatcould be located and measured by GC/MSanalysis of the monomers. These methodshave been applied to cutin containing onlythe C16 family of monomers. Both methodsled to similar conclusions. Most of theprimary alcohol groups are in ester linkagesin the polymer, indicating that the polyesteris held together predominantly by primaryalcohol ester linkages. About half of thesecondary alcohols were also found to be inester linkages, indicating branching and/orcross-links. The conclusion from the mesy-lation analysis is that there is about 0.4 freehydroxyl groups/monomer in tomato fruitcutin, consistent with the number of freehydroxyl groups acetylated with radioactiveacetylating reagents (Kolattukudy, 1977).Based on such results amodel was suggestedfor cutin containing mainly the C16 family ofmonomers (Figure 5).More recently, more direct structural

studies have been carried out using NMRapproaches (Zlotnik-Mazori and Stark,1988; Round et al., 2000; Fang et al., 2001).CPMAS-NMR analysis indicated that cutin

is a moderately flexible netting with mo-tional constraints probably at cross-linksites. Even with citrus cutin, which containsconsiderable amounts ofmid-chain carbonylgroups, more than half of the methyleneswere found to be in the rigid category. Incutins containing primarily mid-chain hy-droxyl groups, with a higher potential forcross-linking, even higher portions of themethylenes may be in the rigid category. Togain further insight into the structural de-tails, oligomers enzymatically generatedfrom cutin were subjected to structuralstudies. Pancreatic lipase and fungal cuti-nase preferentially hydrolyze primary alco-hol ester bonds and release soluble oligo-mers, as demonstrated at the time of firstpurification of cutinase (Purdy and Kolat-tukudy, 1975b). Structural studies on enzy-matically released oligomers using solution-state NMR and liquid secondary ion massspectrometry (LSIMS) gave direct proof forthe predicted presence of secondary alcoholesters involving the 10-hydroxy group of themajor cutin monomer, 10,16-dihydroxyC16

acid. For example, the oligomer released bypancreatic lipase shown in Figure 6 has all ofthe secondary alcohols in ester linkages (Rayand Stark, 1998). A mild chemical reagent,iodotrimethylsilane, that preferentiallycleaves sterically hindered esters underneutral conditions at room temperature,was used to release soluble products (Rayet al., 1998). Isolation of these compoundsby HPTLC followed by structural studiesusingmultidimensional LSIMS also showedparticipation of the alcohol functions at C-10in ester linkages. These structural studiesprovide direct evidence for the type of cross-links or branching involving the mid-chainhydroxyl groups proposed earlier on thebasis of indirect chemical evidence. How-ever, what fraction of the secondary alcoholfunctions are involved in such linkages wasnot determined in the recent studies.

7 Structure of Cutin 9

The percentage of cutin remaining afterthe ester-cleaving depolymerization treat-ments show species-dependent variation.There is convincing evidence that the residuecontains polymethylenic compounds, sug-gesting that they are derived from fatty acids.For example, NMR studies on the insolubleresidues remaining after exhaustive hydro-genolysis with LiAlH4 treatment of cutinfrom the fruits of apple, pepper, and tomatoshowed the presence of methylenes (Kolat-tukudy, 1996). 13C CP-MAS NMR studies ofthe residue remaining after treatment oflime fruit cutin with TMSI showed thepresence of polymethylene function. Pyro-lysis-coupled GLC/MS of the nonester-

bound residues yielded not only productsexpected from C16 and C18 fatty acids, butalso aliphatic hydrocarbons containing 19±26 carbons (Villena et al., 1996), possiblyderived from cuticular waxes containedwithin the cuticular matrix. The nonester-bound residual materials from Clivia mini-ata and Agave americana were studied byFTIR and 13C-NMR spectroscopic analyses,calorimetry, X-ray diffraction, and exhaustiveozonolysis (Villena et al., 1999). On the basisof the results of such studies, it wasconcluded that this depolymerization-resis-tant core consists of an amorphous three-dimensional network of polymethylene mol-ecules containing double bonds and free

1 Cutin from Plants10

C O CH2(CH2)5 CH(CH2)8 C

(CH2)8

O

CH OH

(CH2)5

O

C

(CH2)8

CH OH

(CH2)5

CH2 O C (CH2)14CH2 O

CH2

O

O

O

C(CH2)8

CH O C CH2)14CH3

O(CH2)5

CH2

C (CH2)8

O

CH (CH2)5CH2O

O

C

CH

CH

OH

C

O

(CH2)8CH(CH2)5CH2

O

O

O

O

O

O

Fig. 5 Models showing the type of structures present in the polymer cutin.

carboxylic functions. That such cores con-tain ether linkages was suggested by theobservation that HI treatment released partof the core as soluble materials (Crisp, 1965;Kolattukudy and Walton, 1973). Since struc-tural studies have not been carried out, theexact nature of these monomers remainunknown. Some carbohydrate materialsmay also be part of the nonester-boundresidue. Since the cuticle is known to containsome phenolics and epidermis containsperoxidases, peroxidatively coupled phenol-icsmight be present in the cuticular polymerand may be part of the depolymerization-resistant residual material.

8

Biosynthesis of Cutin

The early notion that cutin was generated byspontaneous oxidation and polymerizationof cellular lipids was replaced with the ideathat lipoxygenase action on unsaturated fattyacids could generate cutin. However, suchhypotheses were not consistent with eventhe monomer composition of cutin. Forexample, the C16 family of monomers andmuch of the C18 family of monomers inmany plants are saturated and not obviouslyderived from the action of lipoxygenases thatrequire cis-1,4-pentadiene structures in theirsubstrates. Modern biosynthetic studiesstarted with the observation that rapidlyexpanding leaves of Vicia faba incorporated14C-labeled acetate and palmitate into theinsoluble residue remaining after extraction

8 Biosynthesis of Cutin 11

HO CH2 (CH2)4 CH2 CH CH2 (CH2)7 CH2 OH

O

C

(CH2)7

O

CH2

CH O C (CH2)7 CH2 CH CH2

O

(CH2)4 CH3

CH2

(CH2)4

CH2 OH

O

C

(CH2)7

O

CH2

CH O C (CH2)14 CH3

O

CH2

(CH2)4

CH2 OH

Fig. 6 Proposed chemical structure of an isolated soluble product of lime cutin depolymerization bypancreatic lipase.

of all soluble lipids (Kolattukudy, 1970a,b).When the insoluble material was subjectedto exhaustive LiAlH4 treatment, 14C-labeledether-soluble compounds were obtainedwhich could be subjected to TLC and radio-GLC. The 14C-distribution in the monomerclearly showed that the observed incorpora-tion represented cutin biosynthesis. Further-more, the most rapidly expanding youngleaves incorporated the exogenous precur-sors most rapidly into cutin, and incorpo-ration was very low in fully expanded leaves.In expanding fruits such as apple and grapeberries, the incorporation of exogenousprecursors was also dependent on the rateof growth, with little incorporation occurringin fully grown fruits. The epidermis wasshown to be the site of cutin biosynthesis.Excised epidermis from the leaves of V. faba,Senecio odoris (Kleinia odora), and pea in-corporated 14C-labeled acetate and palmitateinto cutin, showing that the epidermal layerof cells contains all of the enzymes neededfor the synthesis of fatty acids, subsequenthydroxylations, and incorporation of mono-mers into the polymer. In fruits, only theskin and not the internal tissue incorporatedlabeled precursors into cutin. Clearly, thebiosynthesis of cutin ± just like the biosyn-thesis of the cuticular waxes ± is a specializedfunction of the terminally differentiatedepidermal cells (Kolattukudy and Walton,1973; Kolattukudy, 1996). Thus, early con-clusions on cutin biosynthesis based onobservations on the effects of fatty acids onwounded tissue (viewed as cutin resynthe-sis) were not valid, as wounding does notcause resynthesis of cutin. Since enzymaticactivities truly involved in cutin synthesiswould be present only in the epidermal cells,enzymes and their mRNA should meet thislocalization criterion before their role incutin biosynthesis can be established.

8.1

Biosynthesis of the C16 Family of CutinMonomers

Chromatographic analysis of the depoly-merization products derived from 14C-la-beled acetate or palmitate in rapidly expand-ing V. faba leaves showed that the major partof the label incorporated into the insolublepolymer was in 9 or 10,16-dihydroxy C16 acid,reflecting the composition of cutin mono-mers in this tissue (Kolattukudy, 1970a,b;Kolattukudy and Walton, 1972). A similarlabeling pattern was observed when young,rapidly expanding pea leaves, S. odoris leafdisks of skin disks from rapidly expandingapple fruits or grape berries were incubatedwith labeled C16 acid (Kolattukudy et al.,1973). In the tissues that have cutin com-posed mainly of C16 monomers, exogenouslabeled stearic acid and oleic acid were hardlyincorporated into cutin, and the very smallamount of label that was incorporated was inunhydroxylated acids or �-hydroxy acids.The time-course of labeling showed that atall times the dihydroxy C16 acid was the mostheavily labeled monomer, with much lesslabel in the �-hydroxy acid. Soluble mono-mers did not accumulate, but they could bedetected at very low levels by autoradiogra-phy. Thus, the monomers are incorporatedinto the polymer as soon as they aresynthesized. Exogenous labeled 16-hydroxy-C16 acid was incorporated directly and aftermid-chain hydroxylation into cutin in V. fabaleaves (Kolattukudy and Walton, 1972).Exogenous labeled dihydroxy-C16 acid wasincorporated into cutin without any modifi-cation, and no other components of cutincontained any label. These results, togetherwith the fact thatmid-chain-hydroxylated C16

acid without any �-hydroxyl group was notfound in V. faba cutin or any other plantcutin, strongly suggested that biosynthesisof the C16 family of monomers involves

1 Cutin from Plants12