Proc. Natl. Acad. Sci. USAVol. 81, pp. 6613-6617, November 1984Biochemistry

Alkane biosynthesis by decarbonylation of aldehydes catalyzed by aparticulate preparation from Pisum sativum

(plant wax/CO production/hydrocarbon)

T. M. CHEESBROUGH AND P. E. KOLATTUKUDY*

Institute of Biological Chemistry and Biochemistry/Biophysics Program, Washington State University. Pullman, WA 99164-6340

Communicated by P. K. Stumpf, June 29, 1984

ABSTRACT Mechanism of enzymatic conversion of a fat-ty acid to the corresponding alkane by the loss of the carboxylcarbon was investigated with particulate preparations from Pi-sum sativum. A heavy particulate preparation (sp. gr., 1.30g/cm3) isolated by two density-gradient centrifugation stepscatalyzed conversion of octadecanal to heptadecane and CO.Experiments with [1-3H,1_14C]octadecanal showed the stoichi-ometry of the reaction and retention of the aldehydic hydrogenin the alkane during this enzymatic decarbonylation. This de-carbonylase showed an optimal pH of 7.0 and a Km of 35 ,uMfor the aldehyde. This enzyme was severely inhibited by metalion chelators and showed no requirement for any cofactors.Microsomal preparations and the particulate fractions fromthe first density-gradient step catalyzed acyl-CoA reduction tothe corresponding aldehyde. Electron microscopic examina-tion showed the presence of fragments of cell wall/cuticle butno vesicles in the decarbonylase preparation. It is concludedthat the aldehydes produced by the acyl-CoA reductase locatedin the endomembranes of the epidermal cells are converted toalkanes by the decarbonylase located in the cell wall/cuticleregion.

Alkanes are widely distributed in the plant and animal king-doms (1). Biological hydrocarbons usually have an odd num-ber of carbon atoms, suggesting that they are derived by theloss of one carbon atom from fatty acids with even numbersof carbon atoms. Experiments with higher plant tissue slicesstrongly suggested that hydrocarbons are formed by chainelongation of fatty acids followed by loss of the carboxylcarbon, presumably by decarboxylation (2, 3). Subsequentwork with insects (4) and mammals (5) supported this mech-anism for alkane biosynthesis. Experiments with cell-freepreparations from pea leaves showed that oxygen and ascor-bate were required for the conversion of C32 fatty acid toalkane and that metal ion chelators strongly inhibited alkanesynthesis (6). Subsequent studies showed that a major partof this alkane-generating activity was located in a crude mi-crosomal fraction and that C18 to C32 fatty acids could serveas substrates for alkane formation (7). All of these substratesgave rise to mainly alkanes containing two carbon atoms lessthan the parent acid. Evidence was presented suggesting thatin vitro the aldehyde generated from the parent acid by theclassical a-oxidation was the immediate precursor of the al-kane, whereas in vivo aldehydes generated by acyl-CoA re-ductase might be the immediate precursor of alkanes. It wassuggested that the aldehyde might be decarborlylated to al-kane. In the present paper, we describe the isolation of aparticulate fraction devoid of a-oxidation activity, and wepresent direct experimental evidence for enzymatic decar-bonylation of an aldehyde to alkane.

MATERIALS AND METHODSMaterials. Omnifluor, [1-_4C]stearic acid, [U-3H]tetraco-

sanoic acid, [1-'4Cjsteroyl-CoA, and LiAl3H4 were fromNew England Nuclear. Sodium [1-14C]cyanide was fromICN (Chemical and Radioisotopes Division). Pyridiniumchlorochromate, meso-tetraphenylporphyrin, and RhCP3H2O were from Aldrich. LiAIH4, di-t-butylchlorophos-phine, and Ru(CO)12 were from Alfa Products. The rhodiumchelate was synthesized by the procedure of Monson (8).Ruthenium dicarbonyl tetraphenylporphyrin [Ru(CO)2-(TTP)] was synthesized (9) and activated by substitution ofone CO with di-t-butyl-phosphine (10). The bright red crys-tals of RuCO(TTP)[(C4H9)2P1 gave a spectrum identical tothat reported (11). The oxidizing contaminants in Triton X-100 were removed as described (12).

[1-14C]Octadecanal was synthesized from [1-'4C]octade-canol generated by LiAlH4 reduction of the correspondingacid using pyridinium chlorochromate, and the resulting al-dehyde was purified as described (13). [2-14C]Octadecanalwas synthesized by two cycles of nitrile elongation using amicroscale adaptation of the method of Vederas et al. (14),followed by LiAlH4 reduction of the resulting acid and reoxi-dation of the alcohol to the aldehyde. [1-3H]Octadecanal wassynthesized by reduction of methyl octadecanoate withLiAl3H4, followed by oxidation of the resulting alcohol withpyridinium chlorochromate as indicated above. [2-3H]Octa-decanoic acid was synthesized from 2-bromooctadecanoicacid generated by bromination of octadecanoic acid by theHell-Vollard-Zelensky reaction as described (15). Themethyl ester of the bromo acid was reduced with LiAl3H4 intetrahydrofuran, and the resulting [1,2-3H]octadecanol wasoxidized with CrO3 to [2-3H]octadecanoic acid. [U-3H]Te-tracosanoyl-CoA was synthesized from the acid as described(16).Enzyme Preparations. Pisum sativum (var. dark green per-

fection) was grown in a growth room with a 220C day, 15'Cnight, and a 15.5-hr photoperiod of 16,000 Ix. The apicalmeristem and its enclosing unopened leaflets were harvestedfrom 28- to 36-day-old plants. About 15 g of tissue was ho-mogenized three times for 10 sec each in an Omnimix with0.1 M potassium phosphate, pH 7.0/0.3 M sucrose. The ho-mogenate was filtered through two layers of cheesecloth andcentrifuged for 20 min at 10,800 x g. The resulting superna-tant was centrifuged at 165,000 x g, for 1 hr. The microsom-al pellet (P-2), resuspended in 24 ml of 0.1 M potassiumphosphate, pH 7.0/18% (wt/vol) sucrose, was layered on adiscontinuous density gradient. Each tube, containing 6 mlof sample layered on 7 ml of 25%, 6 ml of 30%, 8 ml of 40%,and 6 ml of 60% sucrose solutions in 0.1 M potassium phos-phate (pH 7.0), was centrifuged at 64,000 x g for 3 hr. Thefirst 4 ml of 60% sucrose (G-1) collected from the bottomwas diluted 1:2 with 0.1 M potassium phosphate and centri-

*To whom reprint requests should be addressed.

6613

The publication costs of this article were defrayed in part by page chargepayment. This article must therefore be hereby marked "advertisement"in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 27

, 202

0

6614 Biochemistry: Cheesbrough and Kolattukudy

fuged for 3.5 hr at 64,000 x g on a second discontinuoussucrose density gradient, which consisted of 8 ml of sample,4 ml of 40% sucrose, and 3 ml of 60% sucrose. The first 2.5ml of 60% sucrose collected from the bottom was diluted 1:2with 0.1 M potassium phosphate and centrifuged at 165,000x g for 1.5 hr. The pellet (G-2) was resuspended in 1 ml of0.1 M potassium phosphate buffer (pH 7.0) and used as thesource of the decarbonylase enzyme.

Electron Microscopic and Chemical Examination. The par-ticulate fraction, recovered by a 1:2 dilution and centrifuga-tion at 165,000 x g for 30 min, was fixed with 2% gluteralde-hyde and 2% OS04 and stained with lead citrate/uranyl ace-tate. The particulate fraction recovered from the gradientswas depolymerized with LiAlH4, and the products were ex-amined by combined gas-liquid chromatography and massspectrometry as described (17).Decarbonylase Assays. The reactions were run in 16 x 125

mm test tubes with serum caps through which polypropy-lene cups were fitted. Each cup contained 1 mg of Rh-Cl[(C6H6)3P]3 on a strip of wetted filter paper. Each reactionmixture contained 0.1 M potassium phosphate (pH 7.0), 36A.M octadecanal, and enzyme in 2.0 ml. The substrate solu-tion was prepared by sonicating 72 nmol of octadecanal in0.5 ml of 0.1 M potassium phosphate (pH 7.0) with 0.1%(vol/vol) purified Triton X-100. After incubating the mix-tures at 30'C for 45 min, 200 A.l of 2 M HCl was added, andany bound CO was released by photolysis by placing a 22 Wfluorescent light 4 cm away from the reaction mixture for 2min. The filter paper trap containing the CO adduct wasplaced directly in scintillation fluid and assayed for 14C. Thelipids were recovered from the reaction mixtures withCHCl3/CH30H (2:1; vol/vol), and the alkanes were isolatedby thin-layer chromatography and assayed for radioactivityas described (2). The alkane fraction was analyzed by radiogas-liquid chromatography.

Trypsin Digestion of G-2. An aliquot (100 ,ul) of the enzymepreparation was homogenized with 25 strokes in a 2-ml TenBroeck homogenizer with 1.1 ml of 0.1 M potassium phos-phate buffer, pH 7.0/20 ,ug of L-1-tosylamido-2-phenylethylchloromethyl ketone-treated trypsin. After a 2-hr incubationat 30°C, the digest was assayed for decarbonylase activity.

RESULTS

Isolation of an Alkane-Synthesizing Particulate Fraction.When the crude microsomal pellet was resuspended and cen-trifuged on a discontinuous sucrose density gradient, a frac-tion enriched in alkane-synthesizing activity was obtained.a-Oxidation activity was present at high levels in all but the60% sucrose fractions (Fig. 1). These 60% fractions cata-lyzed a-oxidation 2 orders of magnitude slower than that ob-served with crude microsomes, but they readily catalyzedalkane synthesis from fatty acids; aldehydes were, however,generated by all fractions (Fig. 1). To minimize a-oxidation,which is the major reaction interfering with the alkane syn-thesis assays, only the bottom 4 ml of the 60% sucrose (G-1)was used to study alkane synthesis. When the G-1 fractionwas diluted and centrifuged on a second sucrose density gra-dient, the 60%o sucrose fraction of this gradient (G-2) con-tained alkane-synthesizing activity, but no a-oxidation activ-ity was present in these samples (Table 1). Thus, the double-gradient procedure yielded a fraction free of the majorcompeting activity. The specific activity of alkane synthesisincreased from 0.01-0.02 nmol/min per mg of protein incrude extracts to 20-25 nmol/min per mg of protein in G-2.Electron microscopic examination of G-1 showed some vesi-les that appeared to be plastids and fragments of cell wall/

cuticle. In the final preparation, G-2, only the latter could befound (data not shown). Chemical examination of the partic-ulate fractions showed the presence of 10,16-dihydroxyhexa-

1.2

0.6

E 1.2

r-0.6

H=

60 40

Aldehyde

30 25 18

fl n-

0.03p-

0.c)1,0 5 10

Fraction

15

FIG. 1. Sucrose density-gradient fractionation of pea leaf micro-somes. The microsomal suspension was centrifuged on a discontinu-ous sucrose density gradient; the interfaces of the 18%, 25%, 30c,40%, and 60%o sucrose layers are marked by arrows. Each fractionwas assayed for a-oxidation activity with [1-14C]palmitic acid andfor alkane synthesis with [U-3H]tetracosanoic acid. Aldehyde pro-duction in the latter assay was measured by thin-layer chromatogra-phy.

decanoic acid, the major cutin monomer (3), only in the frac-tions that contained decarbonylase activity including G-2,the final particulate preparation.

Substrate for Alkane Synthesis. The relative efficiency ofproduction of alkanes from fatty acid, acyl-CoA, and alde-hyde changed with each step in the preparation of the hydro-carbon-synthesizing particulate preparation (Table 1). Thecrude microsomal preparation (P-2) and the particulate prep-aration from the first gradient converted acyl-CoA 3-4 timesas effectively as fatty acids into alkanes. This fraction alsogenerated labeled octadecanal from octadecanoyl-CoA (datanot shown). The particulate fraction from the second gradi-ent readily converted aldehyde to alkane, whereas acyl-CoAwas a poor substrate and free acid was not converted intoalkane. With the crude microsomal preparation, conversionof aldehyde into alkane was barely detected. Presumably,

Table 1. Relative efficiency of conversions of substrates intoalkane and a-oxidation activity of the particulate preparationsfrom pea leaves

Hydrocarbon formation,Particulate nmol/min per mg a-Oxidation,fraction Fatty acid Acyl-CoA Aldehyde nmol/min per mg

P-2 0.02 0.06 0.0002 1.56 (18.7)G-1 5.0 20.3 5.1 0.73 (0.023)G-2 0.0 0.60 24.0 0

Assays with P-2 and G-1 used [n-3H]tetracosanoic acid or tetra-cosanoyl-CoA prepared from it, whereas for assays with G-2,[1-_4C]octadecanoic acid or the CoA ester prepared from it wasused. [1-3H,1-_4C]Octadecanal was used where aldehyde is in-dicated. a-Oxidation was measured using [1-_4C]palmitic acid. Val-ues in parentheses indicate total activity in nmol/min.

Proc. Natl. Acad Sci. USA 81 (1984)

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 27

, 202

0

Biochemistry: Cheesbrough and Kolattukudy

Table 2. Isotopic ratios of alkane generated from dual-labeledoctadecanoic acid and octadecanal by a particulate fractionfrom pea leaves

AlkaneSubstrate (3H/14C)

[2-3H,U-14C]Octadecanoic acid 1.00[2-3H,2- 4C]Octadecanoic acid 0.96[1-3H,2-'4C]Octadecanal 0.74

The G-1 particulate fraction was used for the assay with the acid,and G-2 was used for the assay with octadecanal. In each case, 36/,M substrate was used.

the exogenous aldehyde was diverted into other products bycompeting reactions catalyzed by such preparations. As theparticulate preparation was subjected to density-gradient pu-rification steps, aldehyde became an efficient precursor foralkane and with the final preparation aldehyde was the pre-ferred substrate.

Dual Isotope Labeling Experiments. Previously, a-hydroxyacid was thought to be an intermediate in the conversion of afatty acid to alkane (4, 6), but the present results suggest thatdecarbonylation of the aldehyde generates alkane. To distin-guish between these two possibilities, dual-labeled sub-strates were used, and the results are shown in Table 2. a-Oxidation would be expected to cause loss of at least someof the 3H at the C-2 position of octadecanoic acid during al-kane formation. However, with either [2-3H,U-14C]- or [2-3H,2-14C]octadecanoic acid, the isotopic ratio of the alkanegenerated was identical with that of the substrate, showingthat 3H at C-2 was not lost. In the case of [1-3H,2-14C]octade-canal, the bulk of 3H was retained in the alkane generatedfrom it. These results strongly suggested that conversion ofthe acid to the alkane did not involve a-oxidation but proba-bly involved aldehyde as the intermediate and that the decar-bonylation of the aldehyde took place with retention of 3Hfrom C-1 of the aldehyde in the alkane.

Direct Evidence for Decarbonylation. To detect and mea-sure the amount of CO produced during decarbonylation, wedeveloped a trapping procedure using RhCl[(C6H6)3P]3. Rho-

0

0.~~~~~~~3

C17C16 C18

0 5 10 15Time, min

FIG. 2. Radio gas-liquid chromatography of the alkane generatedfrom [1-'4C,1-3H]octadecanal by the particulate fraction G-2. Thelower tracing shows the thermal conductivity detector response dueto the co-injected n-alkanes indicated. A coiled 3.6-m stainless steelcolumn of 2 mm i.d. packed with 15% FFAP on Chromosorb WHP(Gow-Mac, Bridgewater, NJ) held at 1900C was used, and the efflu-ent was passed through a Nuclear-Chicago radioactivity monitor.

dium and its chelates are known to form stable adducts withCO but have very low affinity for CO2 (18). Under our ex-perimental conditions, 50% of the 14Co generated by chemi-cal decarbonylation of [1-_4C]octadecanal was absorbed bythe Rh complex, but 14CO2 was not trapped by this reagent.When the particulate fraction (G-2) was incubated with [1-3H,1-14C]octadecanal, the Rh complex trapped 14C, whilehyamine hydroxide trapped no radioactivity. Therefore, COand not CO2 was generated from the aldehyde. When thesame reaction mixture was extracted and the products weresubjected to thin-layer chromatography, 3H was found in thealkane fraction. Radio gas-liquid chromatographic analysisof the alkane fraction showed that all of the 3H was con-tained in n-C17 alkane (Fig. 2). Incubation of G-2 with [1-3H,2-14C]octadecanal (3H/ 4C, 1.0) gave C17 alkane with anisotopic ratio of 0.74. These results rule out oxidation of thesubstrate to the acid followed by decarboxylation as well aselongation and decarboxylation.Under the standard assay conditions, an average of 1.28

mol of alkane was produced per mol of CO trapped. Withoutphotolysis, the ratio of alkane to CO was even higher. SinceCO released during the reaction could be binding to some

A100

HC

,75 - \

,50 -

\~.co

Time, min

FIG. 3. (A) Effect of pH on decarbonylase activity. The assayswere run in 0.1 M potassium phosphate (o and *), 0.1 M citratephosphate (e), or 0.1 M Tricine (A), using [1-3H,1-14C]octadecanalas described, and the formation of both CO and alkane (HC) wasmeasured. (B) Time course of alkane and CO formations from [1-3H,1-14C]octadecanal by the decarbonylase preparation.

Proc. Natl. Acad. Sci. USA 81 (1984) 6615

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 27

, 202

0

6616 Biochemistry: Cheesbrough and Kolattukudy

1.8

E oS-u.E 1.2

, 0.6

u

HCfig 44

- II,/ I

-0.05 0 0.1 0.2A1 1/[SI, X10-6

0 20 40Substrate, ,uM

60

FIG. 4. Effect of the concentration [1-'4C,1-3H]octadecanal onthe formation of CO and alkane by the decarbonylase preparation.

components in the reaction mixture, it is not surprising thatthe measured amount of CO was slightly less than theamount of hydrocarbon generated.

Effect of pH, Time, and Concentration of Protein and Sub-strate on Decarbonylation. The decarbonylase showed activi-ty only in a fairly narrow range of pH (Fig. 3A). Little activi-ty was seen below pH 6.0 and above pHI 8.0. Since a fairlysharp pH optimum at -7.0 was observed, all subsequent ex-periments were done at this pH. At this optimal pH, the mea-sured amount of CO released was -80% of the amount ofalkane formed. Alkane formation and CO release from octa-decanal showed near linear increases with time up to -30min of incubation, after which the rate began to slow down(Fig. 3B). The amount of CO released relative to alkaneformed remained constant with the time of incubation. Theamount of alkane formed from octadecanal increased linear-ly with increasing protein concentration up to 4 ug/ml. Therelative amount of CO released, when compared to alkaneformed, decreased with increasing protein concentration,suggesting that binding of CO to some components in theparticulate preparation prevented quantitative trapping ofthe CO generated.Alkane formation increased with increasing octadecanal

concentration, resulting in a typical Michaelis-Menten sub-strate saturation pattern (Fig. 4). Double reciprocal plotswere linear with an apparent Km of 35 uM for octadecanal.Release of CO did not show a typical saturation pattern inthat at very low substrate concentrations the expectedamount of CO was not released, although at higher substrateconcentrations the CO release became normal. With the lowamounts of CO generated at very low substrate concentra-tions, the trapping of CO by the Rh reagent was not effec-tive, probably because of competitive binding of CO bysome components present in the enzyme preparation as not-ed above. However, a double reciprocal plot, obtained with-out including the values acquired at very low substrate con-centration, was linear and showed the same intercept on thev axis.Alkane formation from exogenous fatty acids catalyzed by

previously described cell-free preparations (6, 7) was an indi-rect process involving a-oxidative formation of aldehydes.Therefore, the cofactor requirements observed with suchpreparations cannot be considered valid for the final stepalone. Ascorbate, which was required for alkane formationin the crude preparations (6, 7), was not required for thepresent decarbonylation. Reduced pyridine nucleotides (0.1mM) showed little effect on decarbonylation. Imidazole (1mM), a known inhibitor of a-oxidation (19), inhibited the in-direct alkane formation observed in the crude preparations(6, 7) but showed no effect on decarbonylation. Thiol com-

pounds, such as mercaptoethanol and dithioerythritol at con-

centrations that inhibited alkane formation in crude prepara-tions (6, 20), showed little effect on decarbonylation. How-ever, at a higher concentration (>5 mM), some inhibition ofdecarbonylation was observed. Metal ion chelating agents,such as EDTA (1 mM) and o-phenanthroline (0.1 mM), se-verely inhibited decarbonylation. The enzyme was sensitiveto trypsin treatment and was completely inactivated byfreezing.

DISCUSSIONWhen [1-14C,2-3H]octadecanal was incubated with the par-ticulate preparation obtained by the two density-gradientcentrifugation steps, labeled heptadecane was the only al-kane generated. Obviously this process involved a loss ofone carbon, which was recovered as 14Co by the Rh com-plex, which specifically binds CO. Although the amount ofCO released under the experimental conditions was too lowto permit direct chemical identification, the above evidencestrongly suggests that CO was the product of the reaction.The amount of CO generated from [1-14C,2-3H]octadecanalwas nearly equal to the amount of alkane formed under al-most all experimental conditions. The recovery of CO wasslightly less than the stoichiometric amount, most probablybecause some components present in the enzyme prepara-tion bound a portion of the CO. This explanation is support-ed by the observation that whenever the ratio ofCO generat-ed to amount of protein in the reaction mixture was low,recovery of CO showed the most drastic departure from stoi-chiometric amounts. In spite of this technical difficulty, thestoichiometry of the products generated strongly suggestedthat the enzyme preparation catalyzed the formation of 1 moleach of CO and heptadecane from each mol of octadecanal.Thus, the enzyme can be called a decarbonylase. An analo-gous organic chemical reaction has been recently described,in which case a Ru porphyrin complex catalyzes decarbonyl-ation of aldehydes to alkanes (21). The present case is a bio-chemical analogue of this reaction.The sensitivity to boiling, freezing, and trypsin treatment

of the conversion of octadecanal to heptadecane and COsuggested that the observed decarbonylation was an enzy-matic process. The time-course, protein concentration de-pendence, pH dependence, and substrate concentration de-pendence showed that the particulate preparation containedan enzyme that catalyzed decarbonylation of octadecanal.The mechanism of decarbonylation is not understood. Al-

though ascorbate and oxygen were found to be required formaximal rates of alkane formation by crude cell-free prepa-rations (6), the present results show that decarbonylation perse does not require ascorbate or oxygen. Most probably,generation of aldehyde from the substrate acid by a-oxida-tion catalyzed by such crude preparations required these co-factors. In the nonenzymatic decarbonylation catalyzed byRu porphyrin complex, the metal ion is thought to partici-pate in coordinating with CQ and the aldehydic hydrogen isretained in the alkane formed (20). Inhibition of the presentenzymatic decarbonylation by metal ion chelating agentsstrongly suggested that a metal ion is involved in this pro-cess. It is possible that the metal is part of a porphyrin com-plex present in the preparation, although no direct evidenceto support this possibility is available. In any case, the metalion requirement indicated by the inhibition by chelators andthe observed retention of the aldehydic hydrogen in the al-kane produced by the present particulate preparation sup-port the conclusion that the mechanism of the enzymatic de-carbonylation is probably analogous to that of the nonenzy-matic decarbonylation previously observed.

Subcellular localization of the decarbonylase is not under-stood. With the crude microsomal preparation, aldehydewas not a preferred substrate and the major alkane generatedcontained two carbon atoms less than the substrate acid (6,

Proc. NatL Acad Sci. USA 81 (1984)

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 27

, 202

0

Biochemistry: Cheesbrough and Kolattukudy

7). Since this preparation was obviously a mixture of mem-branes that indirectly generated alkanes from acids, one can-not draw any conclusions about subcellular location from re-sults obtained with it. After the first density-gradient stepacyl-CoA became a preferred substrate for alkane synthesis,and the alkane formed contained a substantial proportion ofthe homolog containing one carbon less than the acid. Obvi-ously acyl-CoA reductase contained in this preparation musthave generated aldehyde, which must have given the alkaneby decarbonylation. In fact, formation of octadecanal fromoctadecanoyl-CoA was demonstrated with this preparation.The final density-gradient centrifugation gave a particulatepreparation that showed a strong preference for aldehyde as

substrate for alkane formation and contained no detectablea-oxidation activity. The only reaction this relatively heavy(1.30 g/cm3) particulate fraction catalyzed was decarbonyla-tion.

Electron microscopic examination of the final decarbonyl-ase preparation showed amorphous structures with no rec-ognizable features characteristic of any subcellular organ-elle. The fragments observed structurally resembled cellwall/cuticle. In fact, chemical examination showed that theheavy particulate fraction, but not the other fractions fromthe gradient, contained cutin. Thus, it is highly likely that thedecarbonylase is associated with cell wall/cuticle matrix.Therefore, it appears probable that the aldehydes generatedby the fatty acid chain elongation and reduction catalyzed bythe endoplasmic reticulum and/or other membranes are se-

creted to the extracellular matrix where the decarbonylase islocated. Thus, the nature of alkane generated depends on thealdehyde available for decarbonylation and not on the speci-ficity of the decarbonylase. In fact, crude cell-free prepara-tions generated alkanes from C16 to C32 fatty acids with no

significant chain-length specificity (7). The high degree ofspecificity observed in vivo probably arises from the speci-ficity of the elongating system that provides the fatty acidsfor reduction and decarbonylation. Until the decarbonylaseis solubilized and characterized, the nature of the enzymeand the mechanism of decarbonylation cannot be elucidated.

We thank Linda Rogers for conducting some of the preliminarywork, Sally Combelic for raising the plants, and Jim Huber for as-

sistance with the electron microscopy. This work was supported in

part by Grant GM-18278 from the U.S. Public Health Service. Thisis Scientific Paper 6851, Project 2001 from the College of AgricultureResearch Center, Washington State University, Pullman, WA.

1. Kolattukudy, P. E., ed. (1976) The Chemistry and Biochemis-try of Natural Waxes (Elsevier/North-Holland, Amsterdam).

2. Kolattukudy, P. E. (1967) Phytochemistry 6, 963-975.3. Kolattukudy, P. E. (1980) in The Biochemistry ofPlants. Lip-

ids: Structure and Function, ed. Stumpf, P. K. (Academic,New York), Vol. 4, pp. 571-645.

4. Chu, A. J. & Bloomquist, G. J. (1980) Comp. Biochem. Physi-ol. 68B, 313-317.

5. Cassange, C., Darriet, D. & Bourre, J. M. (1977) FEBS Lett.82, 51-54.

6. Khan, A. A. & Kolattukudy, P. E. (1974) Biochem. Biophys.Res. Commun. 61, 1379-1386.

7. Bognar, A. L., Paliyath, G., Rogers, L. & Kolattukudy, P. E.(1984) Arch. Biochem. Biophys., in press.

8. Monson, R. S. (1971) Advanced Organic Synthesis: Methodsand Techniques (Academic, New York).

9. Cullen, D., Meyer, E., Srivastava, T. S. & Tsutsui, M. (1972)J. Chem. Soc. Chem. Commun., 584-585.

10. Boschi, T., Bontempelli, G. & Mazzocchin, G.-A. (1979) In-org. Chim. Acta 37, 155-160.

11. Domazetis, G., Tarpey, B., Dolphin, D. & James, B. R. (1980)J. Chem. Soc. Chem. Commun., 939-940.

12. Ashani, U. & Catravas, G. N. (1980) Anal. Biochem. 109, 55-62.

13. Agrawal, V. P. & Kolattukudy, P. E. (1978) Arch. Biochem.Biophys. 191, 452-465.

14. Vederas, J. C., Graf, W., David, L. & Tamm, C. (1975) Helv.Chim. Acta 58, 1886-1898.

15. Kolattukudy, P. E. (1970) Arch. Biochem. Biophys. 141, 381-383.

16. Bishop, J. E. & Hajra, A. K. (1980) Anal. Biochem. 106, 344-350.

17. Walton, T. J. & Kolattukudy, P. E. (1972) Biochemistry 11,1885-1897.

18. Orgel, L. E. (1966) Introduction to Transition Metal Chemistry(Butler & Tanner, London).

19. Martin, R. 0. & Stumpf, P. K. (1959) J. Biol. Chem. 234,2548-2554.

20. Buckner, J. S. & Kolattukudy, P. E. (1973) Arch. Biochem.Biophys. 156, 34-45.

21. Domazetis, G., James, B. R., Tarpey, B. & Dolphin, D. (1981)in Catalytic Activation of Carbon Monoxide, ed. Ford, P. C.(Am. Chem. Soc., Washington, DC), pp. 243-252.

Proc. NatL Acad Sci. USA 81 (1984) 6617

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 27

, 202

0