ETHYLENE PRODUCTION FROM 9-ALANINE BY AN ENZYME PQWDER

JOHN E. THOBIPSOX AND MARY SPENCER Departments of Plant Scieme and Biochemistry, The kl'niversity of Alberta,

Edmonton, Alberta Reedved June 22, 1966

In the investigation of a pathway for ethylene biosynthesis, intermediates arising from the metabolism of 8-alanine-2-B4C to ethylene by enzyme powders from wax-bean coty8edons were isolated and identified b paper and thin-layer chromatography. Quantitative determinations of the lagel have prwided w- idence for the conversion of 8-alanine through malonic semialdehyde, B-hydroxy- propionic acid, and acrylic acid to ethylene.

Introduction Tlaere are indications that production of ethylene is a general phenomenon

of life (1'). Generation of the volatile compound tends to be associated with distinct phases of tissue morphogenesis (2, 3). For example, i t is apparent that ethylene evolution accompanies maturation and ageing of tissue (4). As well, the gas affects many biochemical processes, such as the degradation of claloro- phyll ( 5 ) but explanations for such effects are still speculative. The present studies are concentrated ow the elucidation of a metabolic pathway for ethylene production. Eshblislament of a pathway may also assist in understanding the effects of the volatile compound.

This laboratory has previously reported several biological systems for ethylene production (1, 4, 6-8), among them a subcellular fraction from tomatoes. The particulate fraction obtained a t 35,080 g was chosen because i t gave the highest yield of ethylene of all cell fractions (7). More recently, partially purified enzyme powders were prepared from a subcellular fraction from cotyledons of the wax bean, Phaseslus vulgaris. In the presence of appropriate cofactors, these powders converted P-ala~aine-2-~" to radioactive ethylene (8). We have noax- been able to separate the metabolic intermediates in this reaction by paper and thin-layer chromatography (t.1.c.) and to quantitatively determine their labels. This information indicated a pathway for the conversion of 6-alanine through malonic seinialdehyde, P-hydroxy- propionic acid, and acrylic acid to ethylene. The scheme provides another significant role for P-alanine in metabolism; i t also relates the production of ethylene by cotyledons to reported (9, PO) increases in malonic acid concen- tration. In consideration of the association of ethylene evolution with ageing 06 tissue ( 5 ) , i t may be significant that P-alanine has been shown to be a product of nucleic acid degradation (1 1, 12).

Materials k%N-celluBose powder 300 and silica gel G were obtained from Canadian

Laboratory Supplies Incorporated, Cab-o-sil from Cabot Corporation, Boston

Canadian Journal of Biochemistry. Volume 45 (1967)

563

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564 CANADIAN JOURNAL OF BIOCHEMISTRY. VOL. 45, 196%

18, hIass., fl-alanine-2-14C from VoJk Radiochemicals Co., reagent grade NH40H from Canadian Industries Ltd., and acetic acid, methanol, and ethanol were Fisher reagent grade. All other chemicals were obtained from Eastman Organic Chemicals.

Methods Preparations of the enzyme powders from wax-bean cotyledons, collections

and quantitative analyses of ethylene, and determinations of the radioactivity sf ethylene were conducted as previously described (8).

Preparaeion o$Samples$sr Chrsmaf~graphy After 24-hour ethylene collection periods, the reaction mixtures containing

the enzyme powder and cofactors as listed in Table HI were prepared for both paper chromatography and t.1.c. in the following manner. Ethylamine salts of acids in the mixture were formed by adjusting the pH to %O or above with 33y0, v/v, ethylamine solution in deionized water. Concentration to semi- dryness was accomplished in a Buchler Rotary Evapo-mix a t 40 "C ; the reaction mixtures were diluted to 4 ml with deionized water, made basic to pH PO or above ~vith 33y0 ethylamine solution, and brought to a final volume of 4.5 ml with deionized water. The concentrated samples were stored a t - BO "C.

I rnmediately prior to spotting, it was necessary to @en trifuge the concentrated solutions to rid them of sugars and pslysaccharides that interfered with chromatogram development. (An International model HN centrifuge was used a t full throttle for 5 inin a t 25 OC.) All spots were dried with a cool air stream.

The proposed intermediates propiolactone, acrylic acid, and acetic acid were not produced in sufficient concentration for detection by indicator spray when separated by t.1.c. When these substances were not included as part of the reaction mixture for ethylene production, known arnounts of their ethylarrnine salts were added as markers prior to concentration of the mixture for analysis. Final concentrations of 70 rnM in the 4 5 ml were used consistently and the amount was sufficient for location on the chromatograms by indicator spray.

Paper and Thin-Layer Chromatog~aphy For paper chromatography, 100 pl sf the reaction mixture was spotted on

Whatman No. 1 filter paper (46 cnl X 57 cm) and two solvent systems were used. Acrylate, acetate, and propiolactone were separated by development with water-saturated n-butanol (13) in a tank that was previously equilibrated with 0.125yc ethylamine in n-butanol-saturated water. For good separation, i t was not necessary to equilibrate the paper. p-Alanine, malonate, and a-ketsglutar- ate were separated by development with 20yo ammonium hydroxide in ethanol ; no equilibration was necessary.

For t.l.c., 0.25-mm layers of either NIN-cellulose powder 300 @@By0 in deionized water) or silica gel G (33y0 in deionized water) were spread on glass plates (20 cm X 20 cm) with a Desaga t.1.c. apparatus. After drying, the plates were activated a t 110 "C under vacuum for 4 hour. Then 5-pi and 40-pl spots

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THOMPSON AND SPENCER: ETHYLENE PRODUCTION 565

of the reaction mixtures were resolved by three different systems. Acrylate, acetate, and propislactone were resolved on cellulose-layered t.l.c. plates, and 8-alanine, malonate, and a-ketoglutarate on silica-gel plates. The solvent systems used in both cases were the same as employed for paper chromatog- raphy. Equilibration of the tank, but not of the plates, was required for the cellulose system and no equilibration was required for the silica-gel system. Since @-hydroxypropionate was masked by 6-alanine, a-ketoglutarate, and rnalonate on cellulose plates developed in water-saturated rt-butanol, i t was separated from all the other acids on silica gel G plates with a solvent system composed of 28% NH40H in ethanol; no equilibration was required. This system also gave improved resolution of a-ketoglutarate from 8-alanine and malonate.

For both paper chromatography and t.l.c., the chromatogranls were allo~ved to develop to almost their entire length, with occasional exceptions when paper chromatograms were run off the end for better resolution. Detection was accomplislaed in both types of chromatography by spraying the chromatograms developed with n-butanol with 0.4% chlorophenol red in 95% ethanol, and those developed with NW40H--ethanol and NH40H-methanol with 0.4y0 chlorophenol red in ethanol made basic to pH 10 or more with dilute sodiu~aa hydroxide so8u tion.

Radioactivity nnd Spec$,c- A ctiaity Determinations Paper chromatograms of reaction imixtures containing labeled components

were scanned with a Nuclear-Chicago recorder model 818009, and Eabitron monitor model 1619A. For radioactivity determinations, spots were scraped from sprayed t.1.c. plates into liquid scintillation-counting vials containing 0.5 g of Cab-o-sil (14). Fifteen milliliters of a mixture containing 8.4y0 2,5- diphenylsxaaole iphengilsxaasle(PP0) and 0.005% 2,2'-j$-phenylenebis(5-phenyloxazole) (POPOP) in toluene were added to each vial. The vials were shaken to suspend the labeled particles in the Cab-o-sil, and counting was done with a Nuclear- Chicago liquid-scintillation unit, model 725, operated a t 5 "C.

For determination of specific activities of acetate and malonate, their concentrations were ascertained spectrophoton~etrically. Ethylarnine salts of the acids were isolated by t.1.c. of 10-pl aliquots of the concentrated reaction mixtures. The spots were scraped from sprayed t.1.c. plates and the acids eluted by shaking witla 4.5 ml of deionized water for I hour. These suspensions were cleared by centrifugation with an International model HN centrifuge a t full throttle for 2 nain a t 25 "C. Aliquots of 4 n ~ l of the solutions sf the acids were made to pH 3 with dilute hydrochloric acid. The solutions were subse- yue~atly quantitatively transferred to 25-ml volumetric flasks containing 5 ml sf a 1 : 1 mixture sf 1.25 X IW3 benzidine hydrochloride (1 rnl of 30y0 acetic acid was required in each 108 ml of this solution) and 3.75 X copper acetate (15). The solutions were made to volume with deionized water prior to reading. Spots serving as blanks were scraped from the t.1.c. plates and treated

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566 CANADIAN JOURNAL O F BIOCHEMISTRY. VOE. 45. 1967

in the same manner. Standard curves were prepared from solutions of the ethylamine salts of the two acids. Absorbancies were read a t 242 mp (15) on a Beckman DU spectrophotometer.

Results and Bisc=ussion During the entire procedure of concentration and chron~atography, all the

acid intermediates were maintained as ethylamine salts since some of the free acids are volatile. In some experin~ents, ethanol extracts of the reaction mixtures were concentrated for chromatography. The fresh ethanol extracts were relatively free of materials interfering with chronaatography ; but after storage for 10 days, a complex, probably among ethanol, acryilate, and B-hydroxy- propionate, as judged from its label, was noted to be forming in increasing amounts. No evidence of such a complex was found in aqueous concentrates.

Paper chromatography provided adequate separation of the intermediates (Table I). Scanning of the paper chronsatograms indicated labeled malonate and

TABLE I Rr values of ethylarnine salts of acids in a pathway for ethylene biosynthesis

Thin-Bayer chromatography

Paper chromatography Cellulose Silica gel 6

Water-saturated Ethanol- Water-saturated Ethanol- Methanol- n-butanol* NH48Wh n-butanol* NH40H* NH40H*

Acrylate 0.35 0.67 0.45 0.60 0.84 Acetate 0.27' 0.6'2 0.32 0 . 0 0.84 8-Hy.d~oxypropionate 0.04 0.41 0.04 0.3'62 0.76 Propsolactone 0. 17' 0 . .56 0.22 0.48 0.82 Malsnate 0.08 0.24 0.09 0.09 0.48 a-Ketoglu harate 0.06 0.29 0.06 0.15 0.62 8-Alanine 0.02 0.36 0.03 0.22 0.49

*Development solvent.

B-h~7drcsxyprspionate~ but the monitor was not of sufficient sensitivity to detect the radioactivity of other internaediates.

Thin-layer chromatography provided higher resolutiora than did paper chrom- atography, and more versatility, in that two types sf thin layers were used. Typical W1 values for the etlmylamine salts of the acids are given in Table I, but controls were always run since Rf values are somewhat variable with t.l.e.

The enzynae powder was found to be sensitive to small changes in pH, being virtually inactive outside the range 6.7-7.2 (8). An energy source such as adellosine triphosphate was requisite for ethylene production by the enzyme powder. In the absence of substrates and cofactors the powder was inactive (8). A reaction mixture of the normally effective substrates and cofactors and boiled enzyme powder yielded no ethylene. Bacterial contamination in this ethylene- producing system was found to be negligible throughout a 24-hour peri0d.l

Ethylene was produced by the enzyme powder in the presence of @-alanine PW. A. Stinson and M. S. Spencer. Unpublished information.

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THOMPSON AND SPENCER: ETHYLENE PRODUCTION 567

and cofactors coenzyme A, pyridoxal phosphate, ferric: ion, thiamine pyro- phosphate, naagnesiurn ion, and adenosine triphosphate. Such a cofactor con^

plement implicated transamination and decarboxy%ation, and in conjunction with evidence discussed below suggested a pathway for the conversion of 8-ahnine to ethylene as follolvs.

Propiolactsne

(Non-enzymatic conversion) HOH

'I t MaIonic acid

2 1 Acetic acid

'Traa-asanainase activity was further indicated by an increase ii-a ethylene pro- duction when a-ketoglutara te, an amino-group acceptor, was added to the basic reaction mixture (8). From three experiments (e.g. Table HI, sample 1)

TABLE HI Effects sf acrylic acid, propiolactone, and p-alanine on ethylene production by enzyme powders

from wax-bean cotyledons

Ethylene, mpt for collection times (designated periods in hours)

Sample Powder, Additions to number mglml reaction mixture 0-2 2-20 28-21 21-24 Total

0.5 50 mM8-alanine 0.5 50 m ~ l l propiolactone 1B

50 ml%4B-alanine 2A 0 . 5 50 mMp-alanine 2B 0.5 75 mM propiolactone

75 rnM acrylic acid 50 mM8-alanine

3A 0.75 50 m M 8-alanine 3B 0.75 10 miMp-alanine

16 58 16 15 9 7 M 80 1 54 27 77 184

24hour collection time

4 2 . 0 67.5 lsCi 8-alanine-2-B4C 14 mM@-alanine

5 8.75 67.5 gCi ,~I-alanine-2-~~C 14 rnM B-alanine

6 1 . 0 67.5 lsCi 8-alanine-2-14C 50 mllf propiolactone

15 mrM acrylic acid 29 mil&@-Alanine

Fraction of ethylene derived

Ethylene, Totat d.p.m./rnpl from rnll $.p.m. ethylene 8-ahnine

N s ~ ~ : S a r n p l e numbers 1. 2.3.4, 5, and 6 indicate different batches of powders with differing activities. N = not detectable (less than 5 rn& - d.p.m. = disintegrations per minute. Samples were frozen and thawed before the 21-24 hour ethylene collectiin to release ethylene in solution. 4 1 aampleb were 0.125 M in phosphate buffer (pH '7.8). 1.5 mM in adenosine triphosphate, 2.0 mM in thiamine pyrophosphate, 1.0 mM in Mpn+ added ae MgSOc 0.5 mM in pyridoleal phosphate. 0.2 mM in Feg+ added as Fes(SOc)a, 0.17 mM in coenzyme A, 60 rnlM in a-ketoo glutmate, and 78 rnM in maionic acid. (Additions to this reaction mixture were as indicated in the table.) (The specific activities of 8-alanine were 242 p@l per mmole in samples 4 and 5, and 116 p(3i per mmole in sample 6.)

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568 CANADIAN JOURNAL OF BIOCHEMISTRY. VOE. 45. 1867

it was evident that propiolactone (a ready source of @-hydroxypropionic acid) a t 50 rali?f in the reaction mixture increased ethylene production. The increased production is consistent with the conversion of fl-alanine (presumably by transamination) to malonic semialdehyde, for the latter is quite unstable and could quickly be converted to either fl-hydroxypropionic acid or malonic acid. The presence of acetate in suficient concentration to be detected on a sprayed paper chron~atogram, even in instances where malonic acid was excIuded from the reaction mixture, suggested that malonate was in equilibrium with malonic semialdehyde; acetate could have been derived froill malonate through the action of acetyl-CoA carboxylase (16). Increased ethylene production on addi- tion of xnalonic acid (8) was thought to be attributable to shifts in equilibria favoring production of ,63-hydrsxypropionic acid and ethylene. Hence, prior to label studies, it was apparent that malonic semialdehyde was probably in equilibrium with malonic acid and 0-hydroxypropionic acid.

Acrylic acid in the same system including propiolactone inhibited evolution of the volatile by approximately 70y0 a t 75 mM (Table 11, sample 2) and approximately 25y0 a t 25 n~rW. While this behavior is not expected of an ethylene precursor, the effect sf the acrylic acid may be through a different mechanism. I t is, for example, chemically very active, and may attack some other components of the reaction mixture, such as essential sulfide links on enzymes. (Inhibitor experiments with a subcellular fraction from tomatoes have shown that sulfhydryl groups and probably disulfide links are essential for ethylene biosynthesis (17) .) I t is consistent with the proposed pathway that addition of 50 1n144 propiolactone and 15 mM acrylate decreased the specific activity of ethylene, labeled by 0-alanine-2-14C (Table HI, sample 6), although it is important to bear in mind that different preparations of powder may have differing enzyme activity. In an effort to counteract inhibition by acrylate, malonate was present to shift the equilibria among malonic acid, malonic semialdehyde, and 0-l~ydroxypropionic acid in favor of the latter, and the concentration of 0-alanine was augmented in sample 6. (Ethylene production was greater when the concentration of 0-alanine was 50 mil4 than when it was 10 ITI,W (Table 11, sample 3).) The conversion of 8-alanine to ethylene is approximateIy 0.002% but the proportions of ethylene drived from 0-alanine are approximately 0.6 and 0.75 when the concentrations of the amino acid in the reaction mixture are 14 mM and 29 rnM respectively (Table II, samples 4, 5 , and 6). A sample calculation follows, which is for sample 4 (Table II).

The concentration of 8-alanine was 14 n.mM and the volume of the reaction mixture 20 ml. Therefore the reaction mixture contained (14 X 20)/1000 = 0.28 n~mole of 0-alanine, with a specific activity of 67.5/0.28 = 242 pCi/ mmole. For ethylene, the total count was 2275 d.p.m. ; 1 Ci = 3.7 X 10IQ d.g.s. = 222 X 10l0 d.p.m., and B pCi - 222 X 10q.p .m. Therefore 2275 d.p.m. - 2275/(222 X PO4) $3. The ethylene produced was 168 mpl, or 168/(22,408 X lo3) = 0.075 X lW4 mmole. Its specific activity was {2275/(222 X 1(B4) ) X { 1/(0.875 X ) = 137 pCi/mmole. The proportion of ethylene derived

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THObfVIPSBN AND SPENCER: ETHYLENE PRODUCTHON 569

from /I-alanine can be obtained by dividing the specific activity of the product (ethylene) by that of the substrate ('-alanine) , i.e. 137/242 = 0.565 = 0.57.

Recovefy of administered radioactivity was approximately 50%. In addition to losses during the three solution transfers in the procedure, scraping sf labeled material from the t.1.c. plate to the counting vials was a source of incomplete recoveq. The radioactivities of the intermediates in samples with differing enzyme activities (Table I I I), supported the proposed sequence of reactions

TABLE 111 Labeling pattern of ethylene production from &1hu-h1e-2-~~C

Compound(s) Sample I Sample 2

Ethylene 2,275' 1,220* Acrylate 64,800 29,700 @-H ydrsxypropisnate

-I- propio%actone 580,800 292,500 8-Alanine 83,481,300 79,141,500 a-Ketoglutarate N N

Specific activity per mmmole X

Sample 1 Sample 2

Malonate Acetate

-

*Values are average d.p.m. for complete reaction mixture. N = not detectable. NOTE: Disintegrations per minute (d.p.m.) = c.p.m. corrected

according to a IdC-quenching curve (for the intermediates) and a 14C standard (for ethylene). Samples I and 2 indicate diaerent batches of powder with differing enzyme activities. The reaction mixture for both samples was 0.125 M phosphate buffer (pH 7.0), 1.5 m.ZI adenosine triphosphate, 14 mM @-ahnine. 2.0 mM thiamine pyrophosphate, 1.0 mM Mglf added as MgSOe. 0.5 mM pyridoxal phosphate. 0.2 mM Fsag added as Fe~(F04)s. 0.17 mM coenzyme A. 50 rnM a-ketoglutarate, 70 mM rnalonic acid, and 67.6 pCi 6-alanine-2-14C.

from P-alanine to ethylene, and specific activities of malonate and acetate indicate their formation in a branch pathway. If acrylic acid had been obtained from acetic acid by a pathway other than indicated, propionic acid ~t~ould likely have been an intermediate; propionic acid was not detectable in these reaction n~ixtures. Tlae absence of label on a-ketoglutarate indicated that a conversion of acetate to a-ketoglutarate by enzymes of the tricarboxylic acid cycle did not occur. Rl'lalonic semialdehyde was not detected, but was probabHy ixnmediately converted to either malonic acid or to P-hydroxypropionic acid. Since the concentration of rnalonic acid was found to affect formation of @-hydroxypropionic acid from malonic semialdehyde, endogenous levels of ~nalonate may indeed be a part of a control mechanism for ethylene biogenesis.

Additional evidence that P-alanine serves as a precursor of ethylene in accordance with the proposed sequence of reactions came from other experi- ments (18) in which ethylene was obtained non-enzymatically by a heat- catalyzed SnelI reaction among P-alanine, pyridoxal hydrochloride, and cupric

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570 CAKADIAN JOURNAL OF BIOCHEMISTRY. VOL. 45, 1967

Burg (19, 201, and \;CTang, Persyn, and Krackov (21), using apple tissue slices and PenicilEiurn digitaturn, respectively, reported that the co~~versio~a of glucose carbon atoms to ethylene may follow the route involving gIycolysis and can- version of acetate to tricarboxylic acid cycle intermediates. Wang, Jacobsen, and Tanaka (22) have more recently suggested that

succinate --+ fumarate --+ malate + acrylate --+ ethylene,

I t is quite plartsible that labeled intermediates of the tricarboxylic acid cycle, in the tissues that they used, cortld have been transformed to pyruvate by the action of malic enzyme or phosphoenolpyruvic carbsxykinase. Pyruvate could have been converted to ethylene through acetate and malonate in accordance with our proposed pathway. Further in this respect, n~alonate may have been derived directly from the tricarboxylic acid cycle since there is evidence for the enzymic conversion of oxaloacetate to malonate in plant tissues (16). Gibson and Young (23) recently suggested, on the basis of label experinaents with P. digitaturn, that only the C-2 sf acetate and the C-3 of pyruvate are incor- porated extensively into the ethylene molecule. They also stated (23) that the rate a t which ethylene is labeled by various radioactive substrates indicates a pathway from the tricarboxylic acid cycle through pyruvate and acetate to another molecule that then leads to ethylene synthesis. Such evidence supports our proposal that acetate and malonate lead into ~e pathway for ethylene formation from 8-alanine. Burg and Burg (24) reported that in apple-tissue slices a 200-nlin collection of ethylene labeled by acetate-2-I4C had a higher specific activity than that labeled by acetate-1-14C ; this further supports the evidence for formation of the symmetrical molecule, malonic acid, via acetyl- CoA carboxylase in the pathway fronn acetate to ethylene. Jacobsen and Wang also detected labeled ethylene when 6%. digitaturn was incubated in medium containing acrylate-2(3) -14C (25).

Lieberman, BIapson, and Kunishi (26) have reported labeling of ethylene by methionine-3(4)-'4C in apple-tissue slices. Ethylene was also produced from methionirae non-enzy~natically in a model system (27). Methionine can be converted to propionic acid by known metabolic routes; the C-3 and C-4 of methionine become C-2 and C-3 respectively of propionic acid. I t is known that propionyl-CoA can be transcarboxylated urith oxaloacetate to form methylmalonyl-CoA, which can then be converted by methylmalonyl-CoA isomerase to succinyl-CoA, and presumably through acetate to ethylene. In Clostridi~rn propionisurn there is indication of the conversion of propionyl-CoA to acrylyl-CoA (28). (Both %;CTang (25) and Burg (25) have obtained radioactive ethylene from labeled propionic acid in 6%. digitaturn and apple slices, respec- tively.) Hence l~netabolism of methionine-3(4)-14C could have occurred through propionate t s ethylene.

In concIusion, isolation and quantitative deterininations of Babeled inter- mediates in the conversion of @-alanine-2-14C to radioactive ethylene have indicated the existence of the proposed pathway through n~alonic semialdehyde,

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THOMPSON AND SPEKCER: ETHYLENE PRODUCTION 57 1

P-hydroxyprspisnic acid, and acrylic acid. 0 thers' experiments are consistent with this, although such experiments may, as well, indicate independent systems.

Acknowledgments We are grateful to the National Research Council of Canada for a grant-in-

aid (No. 1451) of this research, and we wish to ackns~vledge the technical assistance sf T. A. Tribe.

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10. K. DUPERON. Compt. Rend., 252,1195 (1961). 11. W. L. BARNES and A. bT. NAYLOR. Plant Physiol. 37,171 (1962). 12. C. S. TSAI and B. AXELROD. Plant PhysioI. 40,39 (1965). 13. M. R. ROBERTS and iV. BUCEK. Aml. Chem. 29,1447 06357). 14. F. SNYDER and N. STEPHENS. Anal. Biochem. 4,128 (1962). 15. Z. B. BRAGANIC. Anal. Chim. Acta, 28,394 (1963). 16. H. E. PATTEE and L. M. SHANNON. Bstan. Gaz. 126,179 (1965). 17. M. MEHERIUK and kI. S. SPENCER. Nature, 204,43 (1964). 18. J. E. THOMPSON, T. A. TRIBE, and ;If. S. SPENCER. Can. J. Biochem. 44,389 (1966). 19. S. P. BURG and M. V. THIMAKN. Arch. Biochem, Biophys. 95,450 (1961). 20. S. P. BURG and E. A. BURG. Science, 148,1190 (1965). 21. C. H. WANG, A. PERSPN, and J. KRACKOV. Nature, 195,1306 (1062). 22. C. H. \VANG, B. ii. JACOBSEN and F. S. TAXAKA. Federation Proe. 224,23 (1964). 23. baa. S. GIBSOK and R. E. YOUNG. Nature, 210,529 (1966). 24. S. P. Bu~Gand E. A. BURG. Nature, 203,869 (1964). 25. B. W. JACOBSEN and C. H. WANG. Plant Physiol. Suppl. 40, xix (1065). 26. M. LIEBERMAN, A. T. KUNHSHI, E. 8V. MAPSON, and B. A. WARDALE. Plant Physiol.

Prac. 40, xix (1965). 27. M. LIEBERMAN, A. T. KUNPSHI, L. W. MAPSON, and D. A. ~VARDALE. Biochem, J. 97,

449 (1965). 28. P. W. VAGELBS, J. M. EARL, and E. R. STADTMAN. J. Biol. Chern. 234,490 (1959).

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